Evaluacion del potencial de microalgas endemicas para el cultivo de artemia franciscana.
Feeding marine organisms under culture conditions is one of the most important efforts to improve performance and ensure production processes. Supplying live food is an irreplaceable activity at early culture stages, so efforts have been made to improve them. The microcrustacean, Artemia spp. is the most used species as live food for larval fish and crustacean species in aquaculture, from nauplius stage to adulthood, used by 85% of the marine species cultured (Godinez et al, 2004, Cisneros & Vinatea, 2009). The employment of Artemia in aquaculture has many advantages; however, the nauplius stages are a source of incomplete food since they have low content of eicosapentaenoic (EPA, 20:5 n-3) and docosahexaenoic (DHA, 22:6 n-3) fatty acids, both important for the development of organisms in aquaculture (Sorgeloos et al., 1986; Mary-Leema et al., 2010), and which should be supplied in feed because of their limited ability to synthesize EPA and DHA. Fortunately, Artemia can be enriched by feeding it with microalgal species that have these fatty acids.
Various marine microalgae are used as food for Artemia, which must meet several nutritional and morphological features to enable their use. Tropical ecosystems have a diversity of these species that can be grown under different environments, and which may have different fatty acid composition (Luong-Van et al., 1999). Some microalgae have a high percentage of polyunsaturated fatty acids (PUFAs) that exceed the sources traditionally used because they include carotenoids and antioxidant vitamins naturally, which are bio-encapsulated in their cell wall (Patil et al, 2007).
In tropical climates microalgal species are usually grown in controlled environmental conditions at 19[degrees]C; however, the optimum temperature for cultivating Artemia is 24[degrees]C (Browne & Wanigasekera, 1999). This temperature difference causes growth inhibition and thermal stress to the microalgae. The use of microalgal strains isolated from tropical temperatures may enhance growth of cultured Artemia and avoids the need for temperature controlled environments. Mass production of marine microalgae under temperaturelight controlled and hatchery-type environments has a high production cost for Artemia spp. The use of algal strains from local environments facilitates cultivation and reduces production costs in outdoor mass scale hatchery systems installed in tropical temperatures, which shows the importance of researching new species of marine microalgae.
Due to the increase of aquaculture activities and the need for alternative fish protein and lipid sources, it is necessary to isolate, characterize, and assess microalgal strains as live food. Our study reports five marine species of microalgae that were partially characterized by using conventional taxonomic tools and further evaluated as live food for Artemia franciscana.
MATERIALS AND METHODS
Isolation and characterization of microalgal strains
Sea water samples were collected from three coastal locations in Bahia de La Paz, Mexico. Isolation of microalgal cells was done by micropipetting, streaking on direct spray, and making serial dilutions in agar culture media. The selection of specific colonies and monospecific cultures was made taking into account some cell characteristics: size, shape, mobility, and pigments. Once monospecific microalgal cultures were obtained, they were processed with a mixture of streptomycin and dicloxacillin, each one at a concentration of 0.222 mg m[L.sup.-1] for 24 h to obtain axenic strains. Strains were incubated at 19 [+ or -] 1[degrees]C under continuous illumination with daylight lamps and cultured in 1 [micro]m filtered sea water, UV irradiated in F/2 enriched medium (Guillard, 1975).
For strain identification, a morphological observation of the cells was performed with a compound microscope (Cupp, 1943; Ruggiero et al, 2009; Spaulding et al, 2010). For individual dimensions of the cells forming the strains obtained, 30 different measurements were taken using Nikon camera equipment Sight Ds-L1 equipped with software previously calibrated and mounted to a Nikon compound microscope Optiphot-2.
For outdoor cultures of Navicula sp., Nitzschia sp., Grammatophora sp., and Rhabdonema sp. F/2 medium was used; they were cultured in 2.8 L flasks as above but incubated at outdoor conditions and then transferred to 18 L carboys, with filtered seawater (1 [micro]m) sterilized with 0.1% sodium hypochlorite for 24 h, and any residual chlorine was neutralized with a sodium thiosulfate solution before use.
Agitation of the microalgal culture was established by means of constant aeration with filtered air through a 0.2 membrane. The number of cells was counted daily using a Neubauer hemocytometer under a conventional compound light microscope. After the fifth day of culture, the microalgae were maintained in a semi-continuous system at the rate of 25% dilution per day. The control treatment (C. muelleri) was maintained at 20[degrees]C with constant aeration and illumination.
Biochemical composition of microalgae
Microalgal composition was determined in cells harvested from outdoor cultures. Samples were collected in triplicate on the fourth day of culture, freeze-dried, and then stored at -40[degrees]C until further analysis. Total weight was calculated according to Sorokin (1973), protein content according to Lowry et al. (1951), and procedure as modified by Malara & Charra (1972). Carbohydrate content was determined according to White (1987) and Dubois et al. (1959). Lipids were determined colorimetrically according to Bligh & Dyer (1959) and procedure as modified by Chiaverina (1972).
To determine the fatty acid profile of the strains, a lipid extraction from the cells was done with hexane by stirring at 22oC for 24 h. Suspensions of the microalgae were then filtered, and supernatants were concentrated under reduced pressure. The methylated fatty acid esters were prepared according to AOAC International (AOAC, 1990), using 14% w/v boron trifluoridemethanol. The fatty acid profile was determined by gas chromatography coupled to mass spectrum (GC-MS) Saturn 2200 (Varian Inc.) ion trap in a SP-2380 capillary column 0.25 mm ID x 0.25 100 mx film (Supelco) using helium gas as carrier fluid. Signals were identified according to their mass spectrum. For fatty acid quantification, areas under the curve of the corresponding signal and the sum of the fatty acids were considered. Each sample was analyzed in triplicate.
Artemia franciscana cultures
Cultivation of A. franciscana was performed on a series of fiberglass conical-bottom vessels with 15 L of sea water and covered with polyethylene sheets. Seawater was filtered through cotton yarn cartridges 20, 10, 5, and 1 [micro]m of porosity and sterilized by means of an UV system and sodium hypochlorite. Sodium thiosulfate was used for further neutralization. A. franciscana cysts (Biogrow, Premium) were decapsulated and incubated following Sorgeloos et al. (1986). After hatching, counting was conducted to adjust a density of nauplii m[L.sup.-1] in the experimental units. Each experimental diet was used to feed groups in triplicate. Feeding started after 10 h-post hatching to ensure food availability in the tanks. Food was provided twice daily throughout the test, adjusting to a concentration of 70,000 cells m[L.sup.-1] for each treatment. Daily temperature, dissolved oxygen, salinity, and pH were recorded. Seven treatments were decided to run the experiment for twelve days. For the first five, a single culture of native species: Schizothyrium sp., Nitzschia sp., Rhabdonema sp. and Navicula sp., was retained to feed A. franciscana. For the sixth one, a mix of Navicula sp. and Schizothyrium sp. was chosen given previous results, and for the seventh treatment Chaetoceros muelleri was retained as control, given the wide utilization of this species as food for crustaceans. Twenty Artemia were sampled from each experimental unit at the beginning and end of the experiment and fixed in 5% formaldehyde to estimate their growth by measuring total length and dry weight (45[degrees]C for 36 h).
To determine cell size, 30 samples per microalgal species were measured, and their average size was calculated. In this experiment, the treatments were analyzed and compared in triplicate; before developing the statistical analysis, experimental data were tested for variance normality and homogeneity. Differences in the content of proteins, lipids, carbohydrates, and fatty acids were determined by one-way variance analysis (ANOVA). Similarly, the effect of the different microalgal species on survival, length, and weight gain in A. franciscana was evaluated by one-way (ANOVA) variance analysis for each case. Percentage data obtained were arcsine-square-root transformed prior to analysis. In cases where significant differences were observed, a Tukey HSD post hoc analysis was applied. Statistic 8.0 for Windows (StatSoft, USA) and [alpha] = 0.05 was used for the statistical analysis.
The results of isolating and purifying the strains are shown in Table 1. Five species of microalgae were isolated showing various cell sizes with transeptical axial lengths from 7.26 to 44.65 [micro]m in apical length for diatoms while Schizochytrium sp. showed zoospores of 9.06 [micro]m (Table 1). Regarding major cellular constituents, the largest proportion of ashes was found in Navicula sp. The highest protein content was shown in Rhabdonema sp. and Chaetoceros muelleri. Regarding lipid content, Grammatophora sp. showed a greater percentage and lower proportion in C. muelleri. The highest carbohydrate content was recorded in Rhabdonema sp. and in Schizochytrium sp., the lowest content was found in Navicula sp. (Fig. 1). Three types of lipids showed a variation of fatty acid composition of the microalgae; Grammatophora sp. showed a lower content of saturated fatty acids while the highest content was found in Nitzschia sp. In the case of monounsaturated fatty acids, Rhabdonema sp. showed a higher content while the smallest proportion was recorded in Nitzschia sp. The PUFA profile showed a variation in the microalgal species isolated where the percentage of docosahexaenoic acid (DHA) was higher in Grammatophora sp. and C. muelleri, but it was not found in Nitzschia sp. and Schizochytrium sp. whereas eicosapentaenoic acid (EPA) was higher in Gramma tophora sp., and it was not found in Schizochytrium sp. (Table 2).
Growth kinetics of outdoor-cultured microalgae
The trends of the microalgae's growth kinetics until the fifth day of culture showed the biggest cell density for Rhabdonema sp. (2,134.3 x [10.sup.3] cell m[L.sup.-1]), Grammatophora sp. (2,128.7 x [10.sup.3] cell m[L.sup.-1]), and control culture of Chaetoceros muelleri (2,028.5 x [10.sup.3] cell m[L.sup.-1]). While the least cell density was obtained in Nitzschia sp. (595.5 x [10.sup.3] cell m[L.sup.-1]) (Fig. 2). This results show significant differences (P < 0.05, ANOVA-One way) among microalgae. Temperature for the different outdoor cultures ranged from 17[degrees] to 29[degrees]C and pH was 8.2 to 9.0.
Feeding Artemia franciscana with microalgal strains
The results on cultured A. franciscana survival after 12-day feeding trials showed significant differences (P > 0.05, ANOVA-One way) by using different strains (Fig. 3). Tukey HSD test showed that A. franciscana fed with Nitzschia sp. was significantly smaller than the survival rates of the other treatments. The results on size increments, starting from nauplius to adult stage, showed significant differences (P > 0.05, ANOVA-One way) (Fig. 4).
After a 12-day culture, instead of the microalga used as food, dry weight of A. franciscana showed significant differences (P > 0.05, ANOVA-One way) from the beginning to the end of the experiment (Fig. 5); the mixed diet of Navicula sp. and Schizothyrium sp. yielded the highest weight while feeding trials with Rhabdonema sp. and Nitzschia sp. resulted in less weight compared to the control diet with C. muelleri.
Water physical and chemical culture parameters are shown in Table 3; all of them were within normal and acceptable ranges for cultivating A. franciscana.
Microalgal cells with a wide range of forms, sizes, shapes, composition, and habits are found in nature, which may have a valuable application for aquaculture. In our study, five species of microalgae with variability in shape and size due to differences in taxon were isolated. One of the main differences that usually occur between microalgae belonging to different genera is their biochemical composition. According to the classification, those identified as Navicula sp., Nitzschia sp., Grammatophora sp., and Rhabdonema sp. belong to the class Bacillariophyceae (diatoms) from which a wide range of cell size is found, such as the genera isolated while the size of Schizothyrium sp. may range from 2.5 to 18 [micro]m depending on its life cycle. Among the various microalgae assessed in our work, Nitzschia sp. was the strain with the largest size (44.9 [micro]m), but the survival of A. franciscana fed by this microalga was affected. The size of the microalgae could determine the ingestion rate by the metanauplius A. franciscana since it has shown preference for microalgal cells of less than 16 [micro]m (Diaz et al, 2006).
Significant differences (P > 0.05) were observed among ash, protein, lipid, and carbohydrate contents in the five strains. However, these variations are attributable to the different genera to which they belong, as their composition can vary by gender and culture conditions (Brown et al, 1997). Among the various microalgal constituents, the total carbohydrate content in Schizotyhrium sp. was the highest compared to the other four strains because they do not belong to Bacillariophyceae, which generally have lower carbohydrate content. However, this carbohydrate content makes them attractive for feeding oysters and scallops (Enright et al, 1986; Whyte et al, 1989). The feed control C. muelleri showed a low lipid percentage due to the stable culture conditions; outdoor cultivated microalgae had higher lipid percentages because under stress conditions the metabolism is oriented to over produce lipids (Courchesne et al, 2009). Values less than 8% in lipids are recommended to feed crustaceans; in our work higher values did not affect growth for A. franciscana. The reported data on growth of Artemia sp. is difficult to compare due to differences in rearing parameters that also affect the composition of microalgae (Seixas et al, 2009). In addition, slight changes in factors as digestibility, palatability and composition will give variations in utilizing nutrients (Glencross et al, 2007).
Water quality parameters in Artemia cultures showed significant variations. Among these variables, oxygen, salinity, and temperature are the factors that can impact its growth and morphological features (BenNaceur et al, 2011). Artemia cultures can withstand and tolerate a wide range of variations in physical and chemical parameters and dissolved oxygen concentrations ranging from 1.0 mg [L.sup.-1] to oxygen saturation. In the experimental units we registered concentrations above 3.8 mg [L.sup.-1]. Salinity levels were within the required values for Artemia (Van Hoa et al., 2011). Temperature was the parameter in which major variations were registered, and it was attributed to environmental variations throughout the day but within the tolerance range for Artemia (Sorgeloos et al., 1986) cultivation. The pH was maintained in cultures without major changes around pH 8, as recommended by Treece (2000).
Our results on A. franciscana survival fed with the various species of microalgae showed similar values to the control group, except when Nitzschia sp. was used as food. These values are higher than those found by Atashbar et al. (2010), and similar to those obtained by Evjemo & Olsen (1999) with microalgae cultured under controlled ambient conditions. In our work, having no control over temperature, temperature variations were observed from 18.6 to 31.9[degrees]C. Artemia parthenogenetica cultures have been successfully maintained in rustic culture tanks at 35[degrees]C (Van Hoa et al, 2011) although Van Hoa (2002) mentions that growth and survival of A. franciscana can be influenced by temperature and salinity. Despite there was no control of environmental variables (outdoors) in our study, our results are comparable to those obtained under controlled culture conditions.
The results of protein, lipid, and carbohydrate contents of the various strains could not be correlated with survival and growth; feed conversion rate values could confirm the best algal feed to Artemia sp. (Maldonado-Montiel & Rodriguez-Canche, 2005).
The low dry weight in Artemia fed wth Nitzschia sp. is attributed to the low consumption of algae and consequently a low chitin synthesis which is the main constituent of the exoskeleton. However, our results on feeding A. franciscana with Rhabdonema sp. did not follow a pattern, requiring further evaluation to determine the cause of its low dry weight. Other authors suggest that energy content of these species of microalgae can affect the growth of Artemia (GarciaUlloa et al, 1999; Godinez et al, 2004).
Among the different strains tested to feed A. franciscana, the ones fed with Grammatophora sp., Schizothyrium sp. and Rhabdonema sp. had a greater size than those fed with the control species (C. muelleri), reaching larger sizes as reported by Atashbar et al. (2010). Utilization of endemic microalgal species from tropical weather, suitable for cultivating Artemia, may facilitate the feeding process since adaptation to these temperatures allows microalgal cultures to be kept in better physiological condition and be utilized as food integrally. The results obtained from cultures of Thalassiosira weissflogii revealed that the diatoms changed their biochemical composition and decreased their growth rate when undergoing salt stress conditions at salinities greater than 25 (Garcia et al, 2012). The same phenomenon may be related to the final composition of C. muelleri when it is ingested at high salt conditions as shown in our work. Additionally, the culture temperature (20[degrees]C) of C. muelleri is below to the Artemia culture temperatures, the temperature difference can affect the growth and cell composition of microalgae, and thus cause a decrease in size. An evaluation of the nutritional quality of cold water species of microalgae showed that an increase in temperature decreased the amount of essential fatty acids and their growth rate, but the work with the microalgal strains isolated from temperate climates showed no variations were found in their cellular constituents due to their plasticity (Ming-Li et al., 2013). The various microalgal strains evaluated in our study can be exempted of this kind of event because of their origin.
Previous works in species of microalgae have shown the variation in their fatty acid profile (i.e., Luong-Van et al, 1999; Patil et al, 2007), and our results are in acordance with these authors. Regarding the total content of saturated fatty acids, the lowest proportion was registered in Grammatophora sp. and the highest content was measured in Nitzschia sp. The content of monounsaturated fatty acids among the various species was in the range from 22.6% in Nitzschia sp. to 51.0% in Schizothyrium sp.
Regarding PUFAs, docosaexanoic (DHA) acid was not found in our cultures of Nitzschia sp. and Schizothyrium sp. However, Chatdumrong et al. (2006) has reported that this fatty acid is considered as a major constituent in Schizothyrium sp. and in a lesser amount in Nitzschia sp., and that its content in both species is increased under heterotrophic culture conditions. Because in our study both species were cultured under autotrophic conditions, it may be the reason for the absence of these fatty acids.
The temperate marine species of microalgae as those isolated in our study may have a high lipid content, the presence of polyunsaturated fatty acids, and docosahexanoic eicosapentanoic acids. The use of endemic species for aquaculture purposes may favor their cultivation under semi-controlled conditions, reducing their production costs. In addition, they can be used as live food. Navicula sp., Grammatophora sp., Rhabdonema sp., and Schizothyrium sp. are better than C. muelleri. According to the results in our study, the use of Nitzschia sp. for feeding and development of A. franciscana is not recommended.
This work was supported by grants from CIBNOR (AC0.3) to F.A. and from UABCS to M.A.C-R. Our thanks to the National Council of Science and Technology of Mexico (CONACyT) for supporting Juan M. Pacheco-Vega, postdoctoral student with a fellowship award. The authors would like to express their gratitude to Nathaniel Rivera-Reyes and Elizabeth Perez Bravo for invaluable technical support and to Diana Dorantes for editorial services.
Association of Official Analytical Chemists (AOAC). 1990. Official methods of analysis. Association of Official Analytical Chemists. Method 92307. Washington, DC., USA.
Atashbar, B., N. Agh & E. Kmerani. 2010. Intensive culture of Artemia urmiana in semi-flow through system feeding on algae Dunaliella and wheat bran. Int. J. Aquacult. Sci., 1: 3-7.
Barclay, W.R., Meager, K.M. & J.R. Abril. 1994. Heterotrophic production of long chain omega-3 fatty acids utilizing algae and algae-like microorganisms. J. Appl. Phycol., 6: 123-129.
Ben-Naceur, H., A.B. Rejeb Jenhani & M.S. Romdhane. 2011. Report d'une nouvelle population d'Artemia (Branchiopoda, Crustacea) en Tunisie: sabkhet halk el menzel. Bull. Inst. Natn. Scien. Tech. Mer de Salammbo, 38: 73-82.
Bligh, E.G. & W.J. Dyer. 1959. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol., 37: 911-917.
Brown, M.R., S.W. Jeffrey, J.K. Volkman & G.A. Dunstan. 1997. Nutritional properties of microalgae for mariculture. Aquaculture, 151: 315-331.
Browne, R.A. & G. Wanigasekera. 1999. Combined effects of salinity and temperature on survival and reproduction of five species of Artemia. J. Exp. Mar. Biol. Ecol., 244: 29-44.
Chatdumrong, W., W. Yongmanitchai, S. Limtong & W.Worawattanamateekul. 2006. Optimization of docosahexaenoic acid (DHA) production and improvement of astaxanthin content in a mutant Schizochytrium limacinum isolated from mangrove forest in Thailand. Mar. Biotechnol., 8: 319-327.
Chiaverina, J. 1972. Techniques d'extraction et analyses des lipids. Universite de Paris, Station Zoologique, Villefranche-sur-Mer, Notes de Travail, 12: 12.
Cisneros, C. & E. Vinatea. 2009. Produccion de biomasa de Artemia franciscana (Kellog, 1906) utilizando diferentes dietas. Ecol. Apl., 8: 9-14.
Courchesne, N.M.D., A. Parisien, B. Wang & C.Q. Lan. 2009. Enhancement of lipid production using biochemical, genetic and transcription factor engineering approaches. J. Biotechnol., 141(1): 31-41.
Cupp, E.E. 1943. Marine plankton diatoms of the west coast of North America. Bull. Scripps Inst. Oceanogr. Tech. Ser., 15: 1-238.
Diaz, A.H., A. Ramirez-Ayvar, D. Godinez-Siodia & C. Gallo-Garcia. 2006. Efecto del tamano de las microalgas sobre la tasa de ingestion en larvas de Artemia franciscana (Kellog, 1906). Zootec. Trop., 24: 1-7.
Dubois, M., K.A. Gilles, J.K. Hamilton, P.A. Rebers & F. Smith. 1956. Colorimetric method for determination of sugars and related substances. Anal. Chem., 28: 350-356.
Enright, C.T., G.F. Newkirk, J.S. Craigie & J.D. Castell. 1986. Evaluation of phytoplankton as diets for juvenile Ostrea edulis L. J. Exp. Mar. Biol. Ecol., 96: 1-13.
Evjemo, J.O. & Y. Olsen. 1999. Effect of food concentration on the growth and production rate of Artemia franciscana feeding on algae (T-iso). J. Exp. Mar. Biol. Ecol., 242: 273-296.
Garcia, N., J.A. Lopez-Elias, A. Miranda, P.N. Huerta & A. Garcia. 2012. Effect of salinity on growth and chemical composition of the diatom Thalassiosira weissflogii at three culture phases. Lat. Am. J. Aquat. Res., 40: 435-440.
Garcia-Ulloa, G.M., J. Gamboa, J.L. Zavala, T. Ogura & P. Lavens. 1999. Influence of diets on length and biomass production of brine shrimp Artemia franciscana (Kellog, 1906). Biol. Mar. Coast. Res., 28: 7-18.
Glencross, B.D., M. Booth & G.L. Allan. 2007. A feed is only as good as its ingredients-a review of ingredient evaluation strategies for aquaculture feeds. Aquacult. Nutr., 13(1): 17-34.
Godinez, D.E., M.C. Gallo R. Gelabert, A.H. Diaz, J. Gamboa, V. Landa & E.M. Godinez. 2004. Crecimiento larvario de Artemia franciscana (Kellog, 1906) alimentada con dos especies de microalgas vivas. Zootec. Trop., 22: 265-275.
Guillard, R.R.L. 1975. Culture of phytoplankton for feeding marine invertebrates. In: W.L. Smith & M.H. Chantey (eds.). Culture of marine invertebrate animals. Plenum Publishers, New York, pp. 29-60.
Lowry, O.H., N.J. Rosebrough, A.L. Farr & R.J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Biochem., 193: 265-275.
Luong-Van, T., S.M. Renaud & D.L. Parry. 1999. Evaluation of recently isolated Australian tropical microalgae for the enrichment of the dietary value of brine shrimp, Artemia nauplii. Aquaculture, 170: 161173.
Malara, G. & R. Charra. 1972. Dosages des proteines partiulaires selon la methode de Lowry. Universite de Paris, Station Zoologique, Villefrenche-Sur-Mer, Notes de Travail, 6: 11.
Maldonado-Montiel, T.D. & L.G. Rodriguez-Canche. 2005. Biomass production and nutritional value of Artemia sp. (Anostraca: Artemiidae) in Campeche, Mexico. Rev. Biol. Trop., 53(3-4): 447-454.
Mary-Leema, J.T., M. Vijayakumaran, R. Kirubagaran & K. Jayaraj. 2010. Effects of Artemia enrichment with microalgae on the survival and growth of Panulirus homarus phyllosoma larvae. J. Mar. Biol. Assoc. India, 52: 1-7.
Ming-Li, T., P. Siew-Moi & C. Wan-Loy. 2013. Response of Antarctic, temperate, and tropical microalgae to temperature stress. J. Appl. Phycol., 25: 285-297.
Patil, V., T. Kallqvist, E. Olsen, G. Vogt & H.R. Gislerod. 2007. Fatty acid composition of 12 microalgae for possible use in aquaculture feed. Aquacult. Int., 15: 19.
Ruggiero, M., D. Gordon, N. Bailly, P. Kirk & D. Nicolson. 2009. The catalogue of life taxonomic classification. Edition 2, Part A. Species 2000 & ITIS Catalogue of Life, 3 February 2012. DVD; Species 2000: Reading, UK.
Seixas, P., M. Rey-Mendez, L.M. Valente & A. Otero. 2010. High DHA content in Artemia is ineffective to improve Octopus vulgaris paralarvae rearing. Aquaculture, 300(1): 156-162.
Sorgeloos, P., P. Lavens, P. Leger, W. Ackaert & D. Versichele. 1986. Manual for the culture and use of brine shrimp Artemia in aquaculture. State University of Ghent, Belgium, 319 pp.
Sorokin, C. 1973. Dry weight, packed cell volume and optical density. In: J.R. Stein (ed.). Handbook of phycological methods. Culture methods and growth measurements. Cambridge University Press, Cambridge, 321-343 pp.
Spaulding, S.A., D.J. Lubinski & M. Potapova. 2010. Diatoms of the United States. http://westerndiatoms. colorado.edu. Reviewed: 28 April 2013.
Treece G.D. 2000. Artemia production for marine larval fish culture. Southern Regional Aquaculture Center, Publ., 702: 1-8.
Van Hoa, N. 2002. Seasonal farming of the brine shrimp Artemia franciscana in artisanal ponds in Vietnam: effects of temperature and salinity. PhD Thesis. University of Ghent, Belgium, 184 pp.
Van Hoa, N., T. Anh-Thu, N.T. Ngoc-Anh & H. ThanhToi. 2011. Artemia franciscana Kellogg, 1906 (Crustacea: Anostraca) production in earthen pond: improved culture techniques. Int. J. Artemia Biol., 1: 13-28.
White, J.N.C. 1987. Biochemical composition and energy content of six species of phytoplankton used in mariculture of bivalves. Aquaculture, 60: 231-241.
Whyte, J.N.C., N. Bourne & C.A. Hodgson. 1989. Influence of algal diets on biochemical composition and energy reserves in Patinopecten yessoensis (Jay) larvae. Aquaculture, 78: 333-347.
Received: 4 October 2013; Accepted: 20 August 2014
Juan M. Pacheco-Vega (1,2), Marco A. Cadena-Roa (2), Felipe Ascencio (1) Carlos Rangel-Davalos (2) & Maurilia Rojas-Contreras (2)
(1) Centro de Investigaciones Biologicas del Noroeste (CIBNOR), Av. Instituto Politecnico Nacional No. 195 Colonia Playa Palo de Santa Rita, La Paz, BCS 23096, Mexico
(2) Universidad Autonoma de Baja California Sur (UABCS), Unidad Pichilingue Apartado Postal 19-B, La Paz, BCS 23080, Mexico
Corresponding author: Juan M. Pacheco-Vega (firstname.lastname@example.org)
Corresponding Editor: Beatriz Modenutti
Caption: Figure 2. Mean values and standard deviations (SDs, n = 3) of cell concentrations (106 cells mL-1) of five strains of microalgae cultures outdoors and control (Chaetoceros muelleri). Superscripts (a, b, c) express statistical significant differences among microalgae.
Table 1. Some characteristics of microalgae isolated from Bahia de La Paz, B.C.S., Mexico. Number indicate median values [+ or -] SD (n = 30). Microalgae (genus) Family Apical length ([micro]m) Schizochytrium sp. Thraustochytriaceae 9.06 [+ or -] 1.41 Nitzschia sp. Nitzschiaceae 44.65 [+ or -] 3.93 Rhabdonema sp. Fragilarioidea 7.26 [+ or -] 1.01 Navicula sp. Naviculaceae 11.45 [+ or -] 1.47 Grammatophora sp. Fragilarioidea 11.58 [+ or -] 2.59 Microalgae (genus) Axial length Seta length transeptic ([micro]m) ([micro]m) Schizochytrium sp. -- -- Nitzschia sp. 6.91 [+ or -] 1.00 44.94 (0.91) Rhabdonema sp. -- -- Navicula sp. 7.87 [+ or -] 0.82 -- Grammatophora sp. 3.94 [+ or -] 0.69 -- Table 2. Fatty acid composition (%) from different microalgae. Values represent the mean. Different superscript letters (a, b, c, d, e) indicate on each column indicate significant differences (P < 0.05) between species. DHA: docosahexaenoic, EPA: eicosapentaenoic acids. Fatty acid Chaetoceros Grammatophora sp. muelleri (control) Saturated 42.83 (b) 34.97 (c) Monounsaturated 31.66 (b) 36.67 (b) Polyunsaturated C18:2 cis,trans 2.20 (a) 1.00 (b) C 18:2 n-6 5.39 (b) 5.08 (b) C 18:3 n-6 0.00 (c) 1.12 (b) C20:2 n-6 0.00 (b) 0.84 (a) C20:3 n-6 0.00 (c) 0.89 (b) C22:4 n-6 4.02 (a) 0.00 (b) C22:6 n-3 (DHA) 2.36 (a) 2.83 (a) C20:5 n-3 (EPA) 11.55 (b) 16.61 (a) Sum polyunsaturated 25.51 (b) 28.36 (a) Fatty acid Rhabdonema sp. Schizochytrium sp. Saturated 57.62 (a) 37.51 (b) Monounsaturated 35.85 (b) 51.02 (a) Polyunsaturated C18:2 cis,trans 0.00 (c) 0.00 (c) C 18:2 n-6 0.00 (c) 8.24 (a) C 18:3 n-6 0.00 (c) 2.19 (a) C20:2 n-6 0.00 (b) 0.00 (b) C20:3 n-6 0.00 (c) 1.04 (b) C22:4 n-6 0.00 (b) 0.00 (b) C22:6 n-3 (DHA) 1.68 (b) 0.00 (c) C20:5 n-3 (EPA) 4.85 (c) 0.00 (d) Sum polyunsaturated 6.53e 11.47 (d) Fatty acid Nitzschia sp. Navicula sp. Saturated 60.41 (a) 41.54 (b) Monounsaturated 22.62 (c) 33.86 (b) Polyunsaturated C18:2 cis,trans 0.00 (c) 0.00 (c) C 18:2 n-6 3.51 (b) 5.68 (b) C 18:3 n-6 0.00 (c) 0.00 (c) C20:2 n-6 0.00 (b) 0.00 (b) C20:3 n-6 0.00 (c) 5.10 (a) C22:4 n-6 0.00 (b) 0.00 (b) C22:6 n-3 (DHA) 0.00 (c) 1.41 (b) C20:5 n-3 (EPA) 13.46 (b) 12.41 (b) Sum polyunsaturated 16.97 (c) 24.60 (b) Table 3. Water quality parameters in cultures of Artemia franciscana fed with different microalgae, lower and maximum values. Microalgae Oxygen Salinity (mg m[L.sup.-1]) Chaetoceros muelleri 4.3-6.6 36.6-46.0 Navicula sp. 5.1-6.3 39.8-43.7 Grammatophora sp. 4.3-7.7 40.1-45.5 Schizothyrium sp. 4.1-5.1 40.8-46.6 Navicula sp. + Schizothyrium sp. 4.5-8.3 40.3-43.5 Rhabdonema sp. 4.3-5.3 40.3-46.9 Nitzschia sp. 3.8-4.7 41.7-45.1 Microalgae Temperature pH ([degrees]C) Chaetoceros muelleri 19.9-30.8 7.4-7.8 Navicula sp. 18.7-31.9 7.7-8.0 Grammatophora sp. 20.9-30.8 7.5-8.4 Schizothyrium sp. 19.3-30.8 7.6-7.9 Navicula sp. + Schizothyrium sp. 19.3-30.12 7.7-8.0 Rhabdonema sp. 18.6-30.4 7.6-8.2 Nitzschia sp. 18.6-30.2 7.6-8.8 Figure 1. Biochemical composition (%) per species of microalgae isolated. Values represent the mean. Ash Protein Lipids Carbohydrate Nitzschia sp. 36.4 31.4 28.5 3.8 Rhakdonema sp. 28.7 46.4 22.7 4.8 Schizochytrium sp. 30.3 26.3 25.6 19.5 Navicula sp. 46.9 12.7 31.7 7.7 Grammalophora sp. 34.9 22.7 32.3 9.3 Chaetoceros muelleri 28.1 46.4 13.2 12.9 Note: Table made from bar graph. Figure 3. Survival of Artemia franciscana after 12 days, fed with different species of marine microalgae. Vertical bars indicate median values [+ or -] SDs Horizontal bars (n = 3). Superscripts (a, b) express significant differences between treatments. Microalgae Chaetoceros muelleri 78.9 (a) Navicula sp. 74.5 (a) Grammatophora sp. 71.2 (a) Navicula sp. + Schizothyrium sp. 77.5 (a) Schizoruim sp. 78.3 (a) Rhadonema sp. 71.5 (a) Nitzschia sp. 23.6 (a) Note: Table made from bar graph. Figure 4. Length (mm) of Artemia franciscana fed with different species of marine microalgae after 12 days. Vertical bars indicate median values [+ or -] SDs Horizontal bars (n = 3). Superscripts (a, b) express significant differences between treatments. Microalgae Nauplius 0.49 (b) Chaetoceros muelleri 2.81 (a) Navicula sp. 3.05 (a) Grammatophora sp. 3.43 (a) Navicula sp. + Schizothyrium sp. 4.22 (a) Schizoruim sp. 4.75 (a) Rhadonema sp. 23.6 (a) Nitzschia sp. 2.69 (a) Note: Table made from bar graph. Figure 5. Dry weight (pg) of Artemia franciscana fed with different species of marine microalgae after 12 days. Vertical bars indicate median values [+ or -] SDs Horizontal bars (n = 3). Superscripts (a, b, c, d, e) express significant differences between treatments. Microalgae Nauplius 11.0 (e) Chaetoceros muelleri 170.7 (e) Navicula sp. 273.3 (b) Grammatophora sp. 262.5 (b) Navicula sp. + Schizothyrium sp. 329.5 (a) Schizoruim sp. 205.8 (c) Rhadonema sp. 97.1 (d) Nitzschia sp. 119.5 (d) Note: Table made from bar graph.
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|Title Annotation:||Research Article|
|Author:||Pacheco-Vega, Juan M.; Cadena-Roa, Marco A.; Ascencio Carlos Rangel-Davalos, Felipe; Rojas-Contreras|
|Publication:||Latin American Journal of Aquatic Research|
|Date:||Mar 1, 2015|
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