Cultivation of Chlorella vulgaris in tubular photobioreactors: A lipid source for biodiesel production

Biochemical Engineering Journal 81 (2013) 120–125

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Cultivation of Chlorella vulgaris in tubular photobioreactors: A lipid source for biodiesel production Davide Frumento a , Alessandro Alberto Casazza a , Saleh Al Arni b , Attilio Converti a,∗ a Department of Civil, Chemical and Environmental Engineering (DICCA), Pole of Chemical Engineering, University of Genoa, Via Opera Pia 15, I-16145 Genoa, Italy b Department of Chemical Engineering, King Saud University, P.O. Box 800, Riyadh 11421, Saudi Arabia

a r t i c l e

i n f o

Article history: Received 23 July 2013 Received in revised form 4 October 2013 Accepted 13 October 2013 Available online 22 October 2013 Keywords: Chlorella vulgaris Microalgae Fed-batch culture Bioreactors Growth kinetics Biodiesel

a b s t r a c t Chlorella vulgaris was cultivated in two different 2.0 L-helicoidal and horizontal photobioreactors at 5 klux using the bicarbonate contained in the medium and ambient air as the main CO2 sources. The influence of bicarbonate concentration on biomass growth as well as lipid content and profile was first investigated in shake flasks, where the stationary phase was achieved in about one half the time required by the control. The best NaHCO3 concentration (0.2 g L−1 ) was then used in both photobioreactors. While the fed-batch run performed in the helicoidal photobioreactor provided the best result in terms of biomass productivity, which was (84.8 mg L−1 d−1 ) about 2.5-fold that of the batch run, the horizontal configuration ensured the highest lipid productivity (10.3 mg L−1 d−1 ) because of a higher lipid content of biomass (22.8%). These preliminary results suggest that the photobioreactor configuration is a key factor either for the growth or the composition of this microalga. The lipid quality of C. vulgaris biomass grown in both photobioreactors is expected to meet the standards for biodiesel, especially in the case of the helicoidal configuration, provided that further efforts will be made to optimize the conditions for its production as a biodiesel source. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Chlorella vulgaris is an unicellular photosynthetic organism belonging to the phylum Chlorophyta that utilizes light energy and carbon dioxide to grow, with higher photosynthetic efficiency and biomass production than terrestrial plants [1]. This microalga may be destined to several applications such as purification of wastewater under either heterotrophic or mixotrophic conditions [2], extraction of value-added compounds of interest for the food and pharmaceutical industries, biofuel production or use as food [3]. High value-added compounds can be obtained from microalgae such as fatty acids (palmitic, eicosapentaenoic, arachidonic, ␥-linolenic, docosahexaenoic acids, etc.) [4], pigments and vitamins [5]. Microalgae are also used in aquaculture as food for rotifers and fry of marine fish [6]. The lipid fraction extracted from microalgae could be alternatively used in different processes for energy exploitation such as the combustion in a diesel engine. It has been reported that the best use of this oil, among the studied ones, is its transformation into a biofuel [7]. The carbon content of dry biomass of Chlorella genus species is about 48% [8,9], and all carbon contained in the cell is ordinarily

∗ Corresponding author. Tel.: +39 010 3532593; fax: +39 010 3532586. E-mail address: [email protected] (A. Converti). 1369-703X/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.bej.2013.10.011

earned from carbon dioxide; therefore, when used for biofuel production, these microalgae can virtually employ gaseous industrial emissions as carbon dioxide source, thereby acting as CO2 sequestrants. For such an ability, they are expected to make, in the future, a significant contribution to the control and reduction of the greenhouse effect. Moreover, taking into account that microalgae have shorter doubling time and higher oil content than crops, their biomass could be effectively exploited for large scale oil fuel production combined with CO2 sequestration [10]. As an additional advantage, microalgae may be preferentially cultivated on lands unsuitable for ordinary agriculture such as desert areas or in large saline water ponds [11]. However, for a cost effective microalgae cultivation, the residual biomass, after extraction, should be processed as well, e.g., by anaerobic digestion, combustion, pyrolysis, etc. [12]. Several studies reported that the amount and quality of lipids contained in microalgal cells can differ as an outcome of changes in growth conditions (e.g., temperature, light intensity, mode of operation and reactor type) or growth medium characteristics (concentration of carbon, nitrogen, phosphate, iron, etc.) [13,14]. C. vulgaris is usually cultivated in Bold’s Basal Medium containing bicarbonate or using CO2 as the solely carbon source [15]. Previous studies reported that inorganic carbon transporters (both for CO2 and HCO3 − ) are located at the plasma membrane in microalgae of the Chlorella genus [16]. As is well known, sodium bicarbonate, which

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Fig. 1. Schematics of the photobioreactors utilized to cultivate Chlorella vulgaris. (A) 2.0 L-helicoidal photobioreactor (1 – aerial view; 2 – frontal view) and (B) 2.0 L-horizontal photobioreactor (1 – lateral view; 2 – frontal view); L: fluorescent lamps.

is the chemical species preferred by microalgae as a carbon source [17], is quickly internalized as such by them, while CO2 undergoes additional dissociation steps to be metabolized. Nonetheless, because of its high solubility in water, bicarbonate can accumulate at very high levels in the medium, possibly leading to salt stress associated with high osmotic pressure. Since a similar stress induced by high NaCl levels was shown to stimulate lipid overproduction in C. vulgaris [18], the effect of increasing concentrations of bicarbonate on biomass growth and fatty acid composition was selected as one of the objectives of the present study. Based on these considerations, NaHCO3 was added to the Bold’s Basal Medium at different concentrations and employed by C. vulgaris as the main inorganic carbon source in addition to the CO2 naturally present in the air. This work is the first attempt to increase the lipid content of this microalga acting on the level of the inorganic carbon source in both batch and fed-batch runs in photobioreactors. Lipids extracted from biomass grown either in Erlenmeyer flasks or in two different tubular photobioreactors (the one having a helicoidal configuration, the other a horizontal one) have been analyzed qualitatively and quantified by gas chromatographic and gravimetric methods.

2. Materials and methods 2.1. Microorganism The microalgal species used in this study was C. vulgaris CCAP 211 (Culture Collection of Algae and Protozoa, Argyll, UK),

an eukaryotic photosynthetic microorganism that grows rapidly because of its simple structure. C. vulgaris was maintained in the Bold’s Basal Medium [15], using NaHCO3 and the CO2 contained in air (about 300 ppm) and NaNO3 as the sole sources of carbon and nitrogen, respectively.

2.2. Culture systems Preliminary cultures were done at room temperature (20 ± 0.5 ◦ C) employing different concentrations of bicarbonate (0, 0.2, 0.4 and 0.8 g L−1 ) in 0.5 L-Erlenmeyer flasks each containing 200 mL of medium. The bicarbonate concentration (0.2 g L−1 ) that ensured the best results was then used in batch and fed-batch runs either in a 2.0 L-helicoidal photobioreactor (Fig. 1, panel A) or a 2.0 L-horizontal one (Fig. 1, panel B). The horizontal photobioreactor was a smaller version of that described in previous work [19]. It was composed of 11 1 m-long glass tubes, with inner and outer diameters of 12 and 14 mm, respectively, arranged horizontally with a slight inclination and exposed to light from one side. The culture was continuously mixed by injecting compressed air at the bottom. A 1.0 L-closed Erlenmeyer flask containing about 760 mL of culture was used at the top of the photobioreactor as a reservoir and degasser, giving a total culture volume of 2.0 L and an exposed surface area of 0.24 m2 . The helicoidal photobioreactor was an enlarged version of a previously described one [20]. It consisted of five 1 m-long cylindrically-shaped glass tubes, with inner and outer diameters of 12 and 14 mm, respectively, arranged helicoidally in series and

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slightly inclined. Medium circulation was granted by an airlift mechanism. A 2.0 L-closed Erlenmeyer flask containing about 1.4 L of medium acted as reservoir and degasser, providing a total culture volume of 2.0 L and an exposed surface area of 0.22 m2 . These arrangements ensured an exposed surface/volume ratio (S/V) in the helicoidal photobioreactor (3.88 cm−1 ) that was twice that of the horizontal one (1.94 cm−1 ), despite of a coincident volume and an almost coincident exposed surface; therefore, it was assumed that any variation in the microalga growth or biomass composition could reasonably be ascribed to a different light distribution in the two tubular configurations adopted in this study. Medium, flasks and photobioreactor tubes were sterilized in autoclave for 20 min at 121 ◦ C in order to prevent any contamination during the early stages of growth. Cultivations were performed under fluorescent artificial light ensured by eight 36 W-lamps, whose intensity, measured by a light meter, model LX-107 (Lutron, Taipei, Taiwan), had an average value of 5 klux, corresponding to 60 ␮mol photons m−2 s−1 . CO2 containing air was injected continuously through pumps, model M2K3 (Schego, Offenbach, Germany), at an air flowrate of 350 L h−1 . Seeking for the best conditions to cultivate C. vulgaris as a raw material for biodiesel production, preliminary batch runs were carried out in flasks for 15 days, while those in the helicoidal photobioreactor either in batch or fed-batch mode and those in the horizontal one only in fed-batch mode for 30 days. Fed-batch cultures were performed by daily adding concentrated NaHCO3 solutions, whose volume and concentration was varied according to the need to restore at the same time the starting concentration of this salt and the water loss by evaporation. Different dosages of NaHCO3 were used to evaluate the effect of such an inorganic carbon source on the biomass lipid content. All the runs were carried out in duplicate, the results expressed as mean values, and the errors as standard deviations from the mean values.

2.3. Analytical procedures Biomass concentration (X) was determined by optical density (OD) measurements at 625 nm by a UV–Vis spectrophotometer, model Lambda 25 (Perkin Elmer, Milan, Italy). All the measurements were carried out in triplicate, and biomass concentration was related to optical density by the equation X = 0.2379 OD (R2 = 0.990). After growth, biomass was separated from the medium by centrifugation at 7500 rpm for 10 min, using a centrifuge model PK131 (ALC, Milan, Italy). Biomass was dried at 105 ◦ C for 48 h, pulverized in a mortar and stored at −20 ◦ C for later analysis of the lipid fraction. The lipid fraction was extracted from biomass according to Krienitz and Wirth [21], i.e., with the method of Folch et al. [22] combined with the use of ultrasounds (mod. UP100H, Hielscher, Teltow, Germany) and increasing the extraction time from 1.5 to 5.0 h. All the extractions were performed on dry biomass using chloroform/methanol (2:1, v/v) as a solvent. After sample extraction, washing twice with the extraction solvent and its complete evaporation, gravimetric analyses were done. Part of the lipid fraction was transesterified through the method described by Zunin et al. [23] and qualitatively characterized by a gas chromatograph, model Ultra Trace (Thermo Finnigan, Milan, Italy), equipped with ZB Vax column and FID detector (Thermoscientific, Milan, Italy), using appropriate reference standards (methyl esters of myristic, palmitic, stearic, oleic, linoleic, linolenic and eicosapentaenoic acids) (Sigma–Aldrich, Milan, Italy). The triglyceride content of the lipid fraction was approximately determined as the ratio of the total mass of the above fatty acid methyl esters to the total mass of the lipid fraction.

Fig. 2. Cell concentration of Chlorella vulgaris versus time during batch cultures in flasks. : Control culture; : 0.2 g L−1 NaHCO3 ; : 0.4 g L−1 NaHCO3 ; : 0.8 g L−1 NaHCO3 .

2.4. Kinetic and yield parameters The average specific growth rate was calculated by the equation:

=

1 Xm ln tm X0

(1)

where Xm and X0 are the maximum biomass concentration and that at the beginning of runs, respectively, and tm is the time at which Xm was obtained. The average biomass productivity was defined as: QP =

Xm − X0 tm

(2)

The average lipid productivity () was defined as the ratio of the concentration of lipids at the end of the run to tm , while the yield of lipids (YL ) as the ratio of the mass of extracted lipids to that of dry biomass. 2.5. Statistical analysis of results The influence of bicarbonate concentration in the medium and photobioreactor configuration either on the above kinetic and yield parameters or the fatty acid composition of biomass lipid fraction was assessed by the analysis of variance (ANOVA), and the Tukey’s post hoc test was used for mean discrimination or mean comparison, according to circumstances. Multiple comparison of the means was performed by least significant difference (LSD) test (p ≤ 0.05). The statistically significant differences were illustrated by different letters in tables. The “Statistica” software version 8.0 (StatSoft, Tulsa, OK, USA) was used for analyses. 3. Results and discussion 3.1. Cultivations Results of batch cultivations carried out in flasks (Fig. 2) show that the addition of bicarbonate in the medium at concentrations of 0.2 and 0.4 g L−1 practically suppressed the lag phase. As a result, the stationary growth phase was reached after only 7 days in the former case, giving a final biomass yield 7.16% higher than the control. On the other hand, the highest bicarbonate dosage (0.8 g L−1 ) led to the lag phase appearance, a delay of the stationary phase and a 13% decrease in final biomass concentration compared with the control, which suggests the occurrence of stress conditions induced by high bicarbonate levels in the medium likely due to excess osmotic pressure.

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Table 1 Parameters of Chlorella vulgaris cultivation under different conditions.  (d−1 )a

Run f

g

Control F (B) 0.2 g L−1 NaHCO3 0.4 g L−1 NaHCO3 0.8 g L−1 NaHCO3 0.2 g L−1 NaHCO3 0.2 g L−1 NaHCO3 0.2 g L−1 NaHCO3

0.184 ± 0.015 0.289 ± 0.027b 0.178 ± 0.014a 0.185 ± 0.013a 0.283 ± 0.017b 0.114 ± 0.009a 0.107 ± 0.002a a

F (B) F (B) F (B) HePh (B) HeP (FB)i HoPj (FB)

Qp (mg L−1 d−1 )b

TMC (g)c

20.8 ± 1.2 33.4 ± 4.5a,b 21.6 ± 2.1a 18.8 ± 1.5a 71.7 ± 5.7c 84.8 ± 6.8c 46.2 ± 2.0b

1.08 ± 0.01 1.64 ± 0.02a 1.49 ± 0.01d 0.97 ± 0.01b 1.72 ± 0.03a 5.02 ± 0.03f 3.57 ± 0.01e

a

Initial pH c

7.40 ± 0.02 7.47 ± 0.01a 7.49 ± 0.01a 8.24 ± 0.05b 7.46 ± 0.01a 7.46 ± 0.02a 7.44 ± 0.07a

 (mg L−1 d−1 )e

YL (%)d

Final pH a

7.67 ± 0.02 8.49 ± 0.01b 8.74 ± 0.03b 8.67 ± 0.03b 7.71 ± 0.01a 7.74 ± 0.11a 7.83 ± 0.17a a

13.1 ± 0.1 21.4 ± 0.1a 20.1 ± 0.1a 20.4 ± 0.1a 4.5 ± 1.8c 10.2 ± 0.3b 22.8 ± 1.5a

b

1.42 ± 0.13a 2.36 ± 0.17a 2.19 ± 0.39a 1.65 ± 0.19a 1.37 ± 0.70a 7.86 ± 1.08b 10.3 ± 1.9b

Different letters (a–f) within the columns show significant differences at p < 0.05. a : specific growth rate. b Qp : biomass productivity. c TMC: total metabolized carbon, calculated from the amount of produced biomass assuming a carbon percentage of dry biomass of 48% [8]. d YL : lipids yield. e : lipids productivity as mass percentage on dry biomass. f F: flask. g B: batch. h HeP: helicoidal photobioreactor. i FB: fed-batch. j HoP: horizontal photobioreactor.

Cultures in photobioreactors were performed in different configurations (helicoidal and horizontal) and modes of operation (batch and fed-batch). To test the effect of the photobioreactor configuration under the best growth conditions, the helicoidal configuration was employed in both batch and fed-batch modes, while the horizontal one only in fed-batch mode. The fed-batch culture performed in the helicoidal photobioreactor gave a maximum biomass concentration (Xm ) that was about 4-fold that of the batch one (Fig. 3), probably because of the well-known capability of the fed-batch process to reduce excess substrate inhibition, whereas Xm was in the horizontal configuration little more than one half of that in the helicoidal one. This occurrence was probably due to the different reactor configurations adopted in this study, in that the helicoidal photobioreactor ensured a S/V ratio (3.88 cm−1 ) that was twice that of the other, despite of the same light intensity, photostage volume and exposed surface. Therefore, the culture in the horizontal reactor was likely to suffer a photolimitation. The pH variations reported in Table 1 show that the final pH values were always higher than the initial ones, consistently with the well-known alkalinizing power of photosynthetic metabolism. It has been reported that, when cell incorporates two bicarbonate ions, one is consumed and internalized in the form of carbon dioxide, while the other released as carbonate, leading to a progressive pH increase in the medium [24,25]. Thus, a pH increase can be

Fig. 3. Cell concentration of Chlorella vulgaris versus time during cultures performed in photobioreactors. ♦: Batch culture in the helicoidal photobioreactor; : fed-batch culture in the helicoidal photobioreactor; : fed-batch culture in the horizontal photobioreactor.

considered as an indicator of good performance of a photosynthetic culture. Cells of microalgae belonging to the Chlorella genus have an average carbon content of about 48% (dry weight) under normal conditions [8,9]; therefore, we used this value to approximately estimate the total amount of carbon metabolized (TMC) by the microalga. As shown in Table 1, this parameter was appreciably influenced by both the mode of operation and NaHCO3 concentration in the medium. In flasks, an inversely proportional relationship between bicarbonate concentration and carbon consumption could be observed. Since TMC depends on cell growth, such a trend suggests that the salt stress induced by excess bicarbonate may have somehow affected the metabolism of the microorganism. 3.2. Growth kinetics In flasks, the highest value of the average specific growth rate ( = 0.289 ± 0.027 d−1 ) was observed at a bicarbonate concentration of 0.2 g L−1 , while those obtained at 0.4 and 0.8 g L−1 were comparable with that of the control fed only with air. This result pointed out that 0.2 g L−1 was the optimal bicarbonate dosage to be adopted in subsequent runs in photobioreactors and that the supposed salt stress already occurred at a concentration as low as 0.4 g L−1 . As expected, the batch culture performed in the helicoidal photobioreactor (in which a 0.2 g L−1 dosage was adopted) exhibited a  value (0.283 ± 0.017 d−1 ) almost coincident with that of the corresponding batch culture in flask, while the lower  values in the fed-batch runs performed either in the helicoidal (0.114 ± 0.009 d−1 ) or the horizontal (0.107 ± 0.002 d−1 ) photobioreactors have to be ascribed to the low bicarbonate feeding rate selected in this study to prevent any salt stress along the time. It should in fact be remembered the importance of this parameter in the fed-batch process, for it is responsible for the nutrient availability to the culture as well as the start-up duration [26]. Among the flask runs, biomass productivity (Qp ) reached a maximum value of 33.4 ± 4.5 mg L−1 d−1 with the 0.2 g L−1 bicarbonate dosage, and then decreased at higher levels, confirming the occurrence of stress conditions induced by high bicarbonate levels in the medium. The Qp values of batch and fed-batch cultures in the helicoidal photobioreactor were comparable, owing to the different time periods taken into consideration to calculate this parameter, while that obtained in the horizontal one was significantly lower, likely because of the above-supposed occurrence of photolimitation.

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Table 2 Percentages of fatty acids in the lipid fraction of Chlorella vulgaris biomass. NaHCO3 (g L−1 ) Fatty acid

Control Fa (B)b

0.2 F (B)

0.4 F (B)

0.8 F (B)

0.2 HePc (B)

0.2 HeP (FB)d

0.2 HoPe (FB)

Myristic (%) Palmitic (%) Stearic (%) Oleic (%) Linoleic (%) Linolenic (%) Eicosapentaenoic (%) Triglycerides (%)

2.56 ± 0.57a 39.24 ± 1.3a 2.01 ± 0.57a 9.94 ± 0.22a 14.62 ± 0.86c,d 20.39 ± 0.93a 11.22 ± 0.02a,c,d 14.24 ± 0.22a,b,c

0.00 ± 0.00a 24.52 ± 2.62a,b 0.00 ± 0.00a 29.21 ± 0.39b 14.11 ± 0.32c,d 18.59 ± 0.96a 13.55 ± 2.14a,b,d 16.14 ± 2.15b,c

0.00 ± 0.00a 32.56 ± 4.05a,b 0.00 ± 0.00a 5.47 ± 1.31a 15.40 ± 2.56d 22.21 ± 3.89a 24.33 ± 4.06b 18.29 ± 4.09b,c

1.12 ± 0.21a 31.60 ± 2.58a,b 6.14 ± 0.25b 12.16 ± 0.53a 8.23 ± 0.24a,b,c 17.59 ± 0.53a 23.12 ± 1.39a,b 26.16 ± 1.38c

46.21 ± 3.22b 18.35 ± 2.55a 0.78 ± 0.21a 10.83 ± 2.34a 7.96 ± 1.31a,b 13.39 ± 2.42a 2.43 ± 0.37c,d 2.85 ± 0.70a

0.00 ± 0.00a 25.70 ± 0.21a,b 2.38 ± 0.93a 63.70 ± 4.36b 5.02 ± 0.55a,d 2.76 ± 1.17b 1.01 ± 0.63c 2.16 ± 0.66a

49.88 ± 0.3b 31.86 ± 3.46a,b 0.00 ± 0.00a 1.99 ± 0.03a 0.00 ± 0.00e 0.00 ± 0.00b 16.24 ± 3.08a,b 12.52 ± 3.09a,b

Different letters (a–e) within the rows show significant differences at p < 0.05. a F: flask. b B: batch. c HeP: helicoidal photobioreactor. d FB: fed-batch. e HoP: horizontal photobioreactor.

3.3. Biomass lipid content and composition The lipid yield (YL ) expressed as percentage of lipids on biomass dry weight provides information on possible metabolic responses of C. vulgaris to both the osmotic stress induced by high bicarbonate levels and the different light distribution within the two photobioreactor configurations. Converti et al. [27] did in fact demonstrate that C. vulgaris is able to react to the stress triggered by a temperature increase or nitrogen shortage synthesizing greater amounts of lipids than in cultures performed under normal growth conditions. However, as shown in Table 1, among the batch cultures carried out in flasks, the highest lipid yield (21.4 ± 0.1%) was obtained with the 0.2 g L−1 bicarbonate dosage, but comparable values (20.1–20.4%) were detected at higher bicarbonate concentrations. These results demonstrate that, contrary to the effects of temperature and nitrogen availability [27], the salt stress induced by excess bicarbonate was not able to appreciably increase the total lipid content of C. vulgaris. On the other hand, the effect of photobioreactor configuration was very pronounced, in that biomass obtained in fed-batch cultures showed in the horizontal photobioreactor a YL value (22.8 ± 1.5%) that was more than twice that obtained in the helicoidal one (10.2 ± 0.3%) (Table 1), while the corresponding batch run gave the lowest lipid yield of the whole set of runs. In batch cultures carried out in flasks, the average lipid productivity () was very low (1.65–2.36 mg L−1 d−1 ) and inversely proportional to bicarbonate concentration in the medium, while in the fed-batch cultures it was thrice as high in the helicoidal configuration (7.86 ± 1.08 mg L−1 d−1 ) and four times as high in the horizontal one (10.3 ± 1.9 mg L−1 d−1 ). The higher  value of biomass grown in the horizontal configuration, despite of a 45% reduction of biomass productivity compared to the helicoidal one, was the likely combined result of a lower S/V ratio and a higher YL value. These results could be exploited as useful guidelines to select the environmental conditions suitable to exalt the lipid synthesis or the growth of C. vulgaris, according to the expectations. The analysis of lipid quality reported in Table 2 provides further information on the sensitivity of C. vulgaris to both the salt stress induced by high bicarbonate levels and the bioreactor S/V ratio. The overall percentage of triglycerides in the lipid fraction of biomass cultured in flasks progressively increased from 16.14 ± 2.15% to 26.16 ± 1.38% when the NaHCO3 concentration was raised from 0.2 to 0.8 g L−1 , which means that the salt stress reduced the fraction of lipids other than triglycerides despite of its scarce influence on the overall lipid content. It is noteworthy that the 0.2 g L−1 dosage ensured the highest yield of saturated and monounsaturated fatty acids to be used for biodiesel production (namely myristc, oleic,

palmitic, and stearic acids) [28], while the other runs in flasks did not show any balanced content of these compounds. Among the cultivations made in photobioreactors, only biomass grown in the horizontal configuration had a triglyceride content higher than 12%, even though the lipid fraction was particularly rich in saturated and monounsaturated fatty acids (76.2–91.8%). In the batch culture performed in the helicoidal photobioreactor, it was detected a high content of myristic acid (46.21 ± 3.22%) but poor contents of the others saturated and monounsaturated fatty acids, while in the fed-batch culture the myristic acid was absent and the oleic one the most abundant (63.70 ± 4.36%). Oppositely, biomass grown in the horizontal reactor exhibited the highest content of myristic acid (49.88 ± 0.34%) and <2% of oleic acid, hence suggesting a direct proportionality between fatty acid chain length and S/V ratio.

4. Conclusions Sodium bicarbonate stimulated the growth of C. vulgaris when added at a concentration of 0.2 g L−1 , whereas it acted as a growth inhibitor at higher levels. Such a salt promoted the synthesis of total lipids and triglycerides compared to the control flask, and the 0.2 g L−1 dosage ensured the most balanced content of fatty acids in terms of biodiesel standards. Among the runs performed in tubular photobioreactors, the fed-batch culture in the helicoidal configuration showed the highest biomass productivity (84.8 ± 6.8 mg L−1 d−1 ) and content of saturated and monounsaturated fatty acids (91.8%) suitable for biodiesel production, while that in the horizontal one the highest overall lipid content (22.8 ± 1.5%). Even though other fatty acids may be used for biodiesel production as well, when using hydrotreatment and/or hydrocracking to convert them, these results may constitute a useful basis to continues research efforts in this direction. These results taken together demonstrate that C. vulgaris biomass can be cultivated under different nutritional stress conditions and surface/volume ratios so as to exalt, according to the expectations, the lipid fraction to be used as a raw material for the production of biodiesel or, alternatively, of other cell components to serve as food or food complement.

References [1] X. Miao, Q. Wu, Biodiesel production from heterotrophic microalgal oil, Bioresour. Technol. 97 (2006) 841–846. ˜ [2] R. Munoz, B. Guieysse, Algal–bacterial processes for the treatment of hazardous contaminants: a review, Water Res. 40 (2006) 2799–2815. [3] P. Spolaore, C. Joannis-Cassan, E. Duran, A. Isambert, Commercial applications of microalgae, J. Biosci. Bioeng. 101 (2006) 87–96.

D. Frumento et al. / Biochemical Engineering Journal 81 (2013) 120–125 [4] K.H.M. Cardozo, T. Guaratini, M.P. Barros, V.R. Falcão, A.P. Tonon, N.P. Lopes, S. Campos, M.A. Torres, A.O. Souza, P. Colepicolo, E. Pinto, Metabolites from algae with economical impact, Comp. Biochem. Physiol. C: Toxicol. Pharmacol. 146 (2007) 60–78. [5] E. Baker, J.J.A. McLaughlin, S.S.H. Hutner, B. DeAngelis, S. Feingold, O. Frank, H. Baker, Water-soluble vitamins in cells and spent culture supernatants of Poteriochromonas stipitata, Euglena gracilis, and Tetrahymena thermophile, Arch. Microbiol. 129 (1981) 310–313. [6] F.A. Muller, The role of microalgae in aquaculture: situation and trends, J. Appl. Phycol. 12 (2000) 527–534. [7] Y. Chisti, Biodiesel from microalgae, Biotechnol. Adv. 25 (2007) 294–306. [8] T. Otsuki, M. Yamashita, T. Hirotsu, H. Kabeya, R. Kitagawa, Utilization of microalgae for building materials after CO2 fixation, in: T. Inui, M. Anpo, K. Izui, S. Yanagida, T. Yamaguchi (Eds.), Advances in Chemical Conversions for Mitigating Carbon Dioxide, Proc. 4th Int. Conf. on Carbon Dioxide Utilization, Kyoto, Japan, 7–11 September 1997, Studies in Surface Science and Catalysis, vol. 114, Elsevier Science Serials, Amsterdam, 1998, pp. 479–482. [9] A. Sánchez Mirón, M.C. Cerón García, A. Contreras Gómez, F. García Camacho, E. Molina Grima, Y. Chisti, Shear stress tolerance and biochemical characterization of Phaeodactylum tricornutum in quasi steady-state continuous culture in outdoor photobioreactors, Biochem. Eng. J. 16 (2003) 287–297. [10] B. Wang, Y. Li, N. Wu, C.Q. Lan, CO2 bio-mitigation using microalgae, Appl. Microbiol. Biotechnol. 79 (2008) 707–718. [11] T. Lebeau, J.M. Robert, Diatoms, Cultivation and biotechnologically relevant products. Part I: Cultivation at various scales, Appl. Microbiol. Biotechnol. 60 (2003) 612–623. ˇ stariˇc, D. Klinar, M. Bricelj, J. Golob, M. Beroviˇc, B. Likozar, Growth, lipid [12] M. Soˇ extraction and thermal degradation of the microalga Chlorella vulgaris, New Biotechnol. 29 (2012) 325–331. [13] A.M. Illman, A.H. Scragg, S.W. Shales, Increase in Chlorella strains calorific values when grown in low nitrogen medium, Enzyme Microb. Technol. 27 (2000) 631–635. [14] Z.Y. Liu, G.C. Wang, B.C. Zhou, Effect of iron on growth and lipid accumulation in Chlorella vulgaris, Bioresour. Technol. 99 (2008) 4717–4722. [15] H.W. Bischoff, H.C. Bold, Phycological studies. Some soil algae from enchanted rock and related algal species, University of Texas Publications 6318 (1963) 1–95. [16] C. Rotatore, B. Colman, The localization of active inorganic carbon transport at the plasma membrane in Chlorella ellipsoidea, Can. J. Bot. 69 (1991) 1025–1031.

125

[17] K. Kumar, C.N. Dasgupta, B. Nayak, P. Lindblad, D. Das, Development of suitable photobioreactors for CO2 sequestration addressing global warming using green algae and cyanobacteria, Bioresour. Technol. 102 (2011) 4945–4953. [18] X. Duan, G.Y. Ren, L.L. Liu, W.X. Zhu, Salt-induced osmotic stress for lipid overproduction in batch culture of Chlorella vulgaris, Afr. J. Biotechnol. 11 (2012) 7072–7078. [19] A. Converti, A. Lodi, A. Del Borghi, C. Solisio, Cultivation of Spirulina platensis in a combined airlift-tubular reactor system, Biochem. Eng. J. 32 (2006) 13–18. [20] R.P. Bezerra, E.Y. Ortiz Montoya, S. Sato, P. Perego, J.C.M. Carvalho, A. Converti, Effects of light intensity and dilution rate on the semicontinuous cultivation of Arthrospira (Spirulina) platensis. A kinetic Monod-type approach, Bioresour. Technol. 102 (2011) 3215–3219. [21] L. Krienitz, M. Wirth, The high content of polyunsaturated fatty acids in Nannochloropsis limnetica (Eustigmatophyceae) and its implication for food web interactions, freshwater aquaculture and biotechnology, Limnologica 36 (2006) 204–210. [22] J. Folch, M. Lees, G.H. Sloane Stanley, A simple method for the isolation and purification of total lipids from animal tissues, J. Biol. Chem. 226 (1957) 497–509. [23] P. Zunin, P. Salvadeo, R. Boggia, F. Evangelisti, Sterol oxidation in meat- and fish-based homogenized baby foods containing vegetable oils, J. AOAC Int. 89 (2006) 441–446. [24] Y. Shiraiwa, A. Goyal, N.E. Tolbert, Alkalization of the medium by unicellular green algae during uptake dissolved inorganic carbon, Plant Cell. Physiol. 34 (1993) 649–657. [25] A.G. Miller, G. Colman, Evidence for HCO3 − transport by the blue green alga (cyanobacterium) Coccochloris peniocystis, Plant Physiol. 65 (1980) 397–402. [26] J.C.M. Carvalho, R.P. Bezerra, M.C. Matsudo, S. Sato, Cultivation of Spirulina (Arthrospira) platensis by fed-batch process, in: J.W. Lee (Ed.), Advanced Biofuels and Bioproducts, Springer, New York, 2013, pp. 781–806, Chapter 33. [27] A. Converti, A.A. Casazza, E.Y. Ortiz, P. Perego, M. Del Borghi, Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production, Chem. Eng. Process. 48 (2009) 1146–1151. [28] S.K. Hoekman, A. Broch, C. Robbins, E. Ceniceros, M. Natarajan, Review of biodiesel composition, properties, and specifications, Renew. Sust. Energ. Rev. 16 (2012) 143–169.