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Cultivation of Desmodesmus subspicatus in a tubular photobioreactor for bioremediation and microalgae oil production a

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Pablo Gressler , Thiago Bjerk , Rosana Schneider , Maiara Souza , Eduardo Lobo , Ana a

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Zappe , Valeriano Corbellini & Maria Moraes a

Environmental Technology Program, University of the Santa Cruz do Sul, Santa Cruz do Sul, RS, Brazil Published online: 25 Aug 2013.

To cite this article: Pablo Gressler, Thiago Bjerk, Rosana Schneider, Maiara Souza, Eduardo Lobo, Ana Zappe, Valeriano Corbellini & Maria Moraes (2014) Cultivation of Desmodesmus subspicatus in a tubular photobioreactor for bioremediation and microalgae oil production, Environmental Technology, 35:2, 209-219, DOI: 10.1080/09593330.2013.822523 To link to this article: http://dx.doi.org/10.1080/09593330.2013.822523

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Environmental Technology, 2014 Vol. 35, No. 2, 209–219, http://dx.doi.org/10.1080/09593330.2013.822523

Cultivation of Desmodesmus subspicatus in a tubular photobioreactor for bioremediation and microalgae oil production Pablo Diego Gressler, Thiago Rodrigues Bjerk, Rosana de Cassia Souza Schneider∗ , Maiara Priscilla Souza, Eduardo Alcayaga Lobo, Ana Letícia Zappe, Valeriano Antônio Corbellini and Maria Silvana Aranda Moraes Environmental Technology Program, University of the Santa Cruz do Sul, Santa Cruz do Sul, RS, Brazil

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(Received 25 November 2012; final version received 27 June 2013 ) The microalgae Desmodesmus subspicatus (Chlorophyta) was cultivated in a tubular photobioreactor using effluent from the wastewater treatment plant of the University of Santa Cruz do Sul, Brazil to demonstrate the reactor’s operation. The algae’s ability to remove nutrients from wastewater and the oleaginous potential of the algae’s biomass were also evaluated. Total phosphorus and ammonia nitrogen were measured. The photobioreactor consisted of a system of three acrylic tubes, a reservoir, connections and a CO2 supply. The gas supply was semicontinuous with CO2 added from a cylinder. The culture’s growth was estimated from cell numbers counted on a daily basis. Lipid content in the biomass was analysed using gas chromatography. A maximum cell density of 9.11 × 106 cells mL−1 and a dry weight of 234.00 mg L−1 were obtained during cultivation without CO2 , and these values rose to 42.48 × 106 cells mL−1 and 1277.44 mg L−1 , respectively, when CO2 was added to the cultivation. Differences in the quality of the effluent and the presence of CO2 did not result in different lipid profiles. The presence of palmitic acid and oleic acid was notable. The average extracted oil content was 18% and 12% for cultivation with and without the input of CO2 , respectively. Keywords: microalgae; bioremediation; photobioreactor; oil; Desmodesmus subspicatus

1. Introduction The enormous biodiversity and consequent variability in the biochemical composition of microalgae, as well as the use of genetic improvement and the establishment of largescale cultivation technologies, have allowed microalgae to be utilized in various applications, such as the extraction of beta-carotene, astaxanthin and lutein. Moreover, oils rich in fatty acids of nutritional interest, such as oleic and linoleic acids, can also be obtained from microalgae.[1–5] Several studies on the cultivation of algae have been conducted for the treatment of wastewater from industrial processes, biological detoxification and the removal of heavy metals. Moreover, these organisms are widely used as bioindicators for the detection of nutrients and toxic substances (e.g. detergents and herbicides).[2–7] The applications of algae also extend to other areas, such as the production of molecules with potential use in agriculture as biofertilizer. In addition, algae can mitigate the greenhouse effect by assimilating carbon dioxide (CO2 ) produced from the burning of fossil fuels and wasteful agricultural practices, such as forest burns. Finally, fossil fuel energy is finite, and future economic problems in many countries depend on it; algae may become important in the production of biofuels, such as biodiesel, biogas, hydrogen and alcohol.[3,8–13] ∗ Corresponding

author. Email: [email protected]

© 2013 Taylor & Francis

Oleaginous microorganism production has shown great promise in the area of energy production because these organisms are a lipid source with a fast growth rate. Initiatives that involve the cultivation of algae as an energy source are already installed in various locations and should propel the bioenergy sector in this new millennium with respect to economic and environmental considerations and also with large amounts of water resources.[14,15] However, several concerns still remain in this industry, such as the most economical form of cultivation, the amount of raw materials obtained with algae compared with other oleaginous plant species, the production of algae with low water consumption and the viability of inexpensive inputs (e.g. wastewater and CO2 from the exhaust of power plants).[1,2,16,17] Microalgae are an option for wastewater treatment involving bioremediation promoted by open systems such as high rate algal ponds or by closed systems such as photobioreactors, combining microalgae production with wastewater treatment.[18] In this context, this study aimed to construct and operate a tubular photobioreactor for the cultivation of the microalgae Desmodesmus subspicatus (R. Chodat, E. Hegewald and A. Schmidt (Chlorophyta)) grown in effluent from the wastewater treatment plant of the University of Santa Cruz

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do Sul (WTP-UNISC) in southern Brazil, as well as to evaluate both the algae’s ability to remove nutrients from the wastewater and the oleaginous potential of the algae’s biomass.

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2. Materials and methods 2.1. Effluent and biological materials Effluent was collected at the WTP-UNISC in the municipality of Santa Cruz do Sul, RS, Brazil. The strain of D. subspicatus used in this work was kindly provided by the Laboratory of Ecotoxicology at UNISC. There is not a collection of strains in this lab, so a record number is not included here. The inoculum was peaked weekly by a qualified professional for use in ecotoxicological tests. Taxonomic identification of the species was guaranteed by a legally certified Brazilian laboratory. This species is also found in local water bodies. The strain was maintained in a growth chamber under continuous illumination of 5000 lux at 25◦ C with culture medium Chu12 . The strain was subsequently acclimatized to a solution of 3 g L−1 N:P:K (18:6:18). The microalgae were grown in 75% of the volume of previously sterilized 500 mL Erlenmeyer flasks. Subcultures were homogenized by aeration and maintained under artificial illumination at 2500 lux with no photoperiod. These subcultures were used in tests in the photobioreactors as inoculum. 2.2.

Analytical characterization of the WTP-UNISC effluent All physical and chemical analyses of the effluent were carried out at the Analytical Center of the University of Santa Cruz do Sul, which is registered in the State Foundation of Environmental Protection FEPAM-RS as an Environmental Analysis Laboratory under certificate number 17/2009-DL. All analyses were conducted in accordance with Standard Methods [19] as follows: alkalinity (titrimetric method), biochemical oxygen demand (BOD, incubation at 20◦ C for five days), chemical oxygen demand (COD, open reflux with potassium dichromate/titrimetry), total phosphorus (colorimetric, ascorbic acid method), ammoniacal nitrogen (N-NH3 ) (distillation/titrimetry) and total nitrogen (Kjeldahl method/digestion, distillation, titrimetry). Analyses were conducted according to the methodology outlined by the American Public Health Association.[20] The laboratory was acclimatized at 26◦ C, and the temperature inside the photobioreactor was monitored daily by removing a 50 mL aliquot. The pH was monitored directly in the photobioreactor tank for the entire duration of the experiment and was recorded every 24 h. These parameters were chosen to evaluate the system with respect to the removal of nitrogen, phosphorus and organic material loads present in the effluent. Flocculation was induced in the final samples for analysis of the

effluent using sodium hydroxide. After biomass separation, the supernatant was neutralized; then, the samples were analysed. 2.3. Photobioreactor The photobioreactor presented in this paper was fully developed by the authors. There is a wide variety of styles and concepts in the literature, but this proposal sought a different approach from many past studies, especially the approach found in Tavares and Rocha.[21] The photobioreactor built and used in this study consisted of a system with a reservoir and three acrylic tubes (0.1 m diameter and 1 m height). The photobioreactor was operated in continuous mode. The gas supply was semicontinuous with CO2 added from a cylinder to evaluate CO2 fixation by D. subspicatus. The tubes were connected by plastic hoses to circulate a 30 L volume throughout all of the tubes, as shown in Figure 1. A submerged pump was used in the reservoir for the recirculation of this volume. The reservoir was also equipped with a recirculation system in which part of the submersed pump flow was recirculated in the reservoir to reduce cell sedimentation. The air (environment) was dispersed in the system via a diaphragm compressor with a flow rate of 0.22 vvm that was connected by a hose to a diffuser located in the centre of the first tube. The tubes were fitted with a gas recirculation system in which the first gas outlet was connected to a diaphragm compressor. The compressor sent the gas mixture that contained the volume of non-assimilated CO2 and oxygen resulting from the photosynthetic process of the first tube to the gas diffuser in the base of the second tube. This same arrangement was used between the second and third tubes. The gas output of the third tube was connected to an Erlenmeyer flask that contained a solution of NaOH for the fixation of the CO2 not fixed to the culture by bubbling. This measure of gas recirculation was designed to promote gas exchange in the system, which is fundamental to the success of this procedure, and to force CO2 through the system to promote greater contact with the microalgae. The photobioreactor received artificial light from eight 32 W lamp bulbs that totaled 0.0062 W cm−1 per tube, as shown in Figure 1. The development of the D. subspicatus culture was assessed after the addition of inoculum to the effluent from the WTP-UNISC. CO2 was fed at 6.2% in relation to the air flow (0.22 vvm), which was introduced into the system through a fluxometer to control gas flow. A solenoid valve was used to control the injection of CO2 into the system, which was interspersed in periods of 3 min every 2 h. The laboratory was acclimatized at 26◦ C. 2.4. D. subspicatus growth analysis The growth in cell density of D. subspicatus was recorded daily and was used to calculate growth curves, which also

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Environmental Technology

Figure 1.

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Tubular photobioreactor with bubble column.

involved monitoring the maximum cell density (MCD), cultivation time and growth rate (k) from aliquots taken in triplicate each day from the photobioreactor. Aliquots were collected in the central part of the tubes (not shown in the schematic drawing of the photobioreactor) under homogenization aeration provided by the system. Thus, collection of cells in the areas of highest concentration was avoided. The culture was initiated at a cell density of 4.75 × 106 cells mL−1 . Every 24 h after the onset of the cultures, samples were extracted for the determination of cell density. Counts were performed using an optical microscope with the aid of a Neubauer chamber and a manual cell counter, and the number of cells was calculated as the average of three counts.[21] 2.4.1. Growth curves Growth curves were prepared with the daily cell density of the aliquots obtained in triplicate, where one aliquot was obtained from each tube and also from the reservoir. For the gas flow, the study considered only one input and one output to the gas system in evaluating the assimilation ability of the culture. Therefore, each average cell density value is the result of 12 observations day−1 (three aliquots × four collection sites). 2.4.2. Cultivation time (T) This parameter was determined from the number of days elapsed between the start of cultivation and the day that the culture reached MCD.

2.4.3. Maximum cell density This parameter was defined as the maximum number of cells obtained per millilitre before the culture reached the stationary phase of the growth curve, irrespective of the time elapsed since the beginning of the culture. 2.4.4. D. subspicatus biomass analysis The D. subspicatus biomass was separated from the culture medium by flocculation. After separation and drying, the biomass was subjected to lyophilization, lipid extraction and analysis by gas chromatography coupled to mass spectrometry.[22,23] The biomass was recovered by flocculation with a solution of 6 mol L−1 NaOH at a concentration of 5 mL L−1 of culture. The biomass was decanted into a plastic container, and the supernatant was extracted for effluent sampling. The biomass was then transferred for kiln drying at 65◦ C and lyophilization at −40◦ C under vacuum for 6 h. The dry weight was expressed in milligrams of dry biomass (lyophilized) per litre of culture.[24] 2.4.5. Lipid fraction of D. subspicatus The lipid extraction procedure was performed according to the methodology of Bligh and Dyer.[25] Samples of 1.0 g of lyophilized microalgae were weighed into 12 mL test tubes. Next, 3 mL of a mixture of CHCl3 /CH3 OH (2:1, v/v) and 10 μL of a solution of butylated hydroxytoluene were added. The samples were subsequently subjected to

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sonication (USC 700, frequency 55 Hz) in an ice bath for three cycles of 15 min. The extracts were stored for 24 h at 4◦ C and protected from light (i.e. covered with aluminium foil to prevent photooxidation of samples) for subsequent lipid extraction. The extracts were again subjected to sonication for three cycles of 15 min each and then centrifuged for 10 min. To the separated supernatant, 2 mL of distilled water and 1 mL of chloroform were added. After another centrifugation, the lower phase (chloroform:lipid) was separated and transferred to a pre-weighed glass container. The aqueous phase was washed with 1 mL of chloroform and centrifuged for 10 min, and the lower phase was recovered and incorporated into the previous chloroform extract. After the solvent was evaporated, the lipid fraction was weighed, and the total lipid quantity was determined in % (LB ) of dry biomass and milligrams of lipid per litre of microalgae culture (LC ) according to the following equations: (F2 − F1 ) × 100 , m LB × B L LC = , 100 LB =

(1) (2)

where LB is the total lipids (in % of lyophilized biomass), LC the total lipids (in mg L−1 of microalgae culture), F1 the empty flask weight (mg), F2 the flask weight+total lipids (mg), m the lyophilized biomass sample weight (mg), BL the quantity of dry biomass (lyophilized) per litre of microalgae culture (mg L−1 ). 3.

Results and discussion

3.1. Analysis of the D. subspicatus growth 3.1.1. Cultivation time With respect to the cultivation time required to reach stationary growth phase, the culture in the effluent medium without the addition of CO2 required a cultivation time of six days, reaching stationary phase between days four and six. These results suggest either a limitation of carbon, the precipitation of phosphate or a minimal quantity of nutrients; these hypotheses will be tested in other experiments. To date, the literature contains no data regarding such conditions as those described in this experiment to allow a more accurate comparison. The cultivation time was greater when CO2 was added to the culture medium, and the stationary phase was reached only after the seventh cultivation day. Figure 2 shows that D. subspicatus adapted quickly to the medium. 3.1.2. Maximum cell density The MCD is defined as the maximum number of cells obtained per millilitre of culture before the stationary phase of the growth curve has been reached, irrespective of the time elapsed since the beginning of the cultivation.

Figure 2. Growth curve of D. subspicatus tested in both treatments (with and without added CO2 ). Observations were made at intervals of 48 h during the trial period, and the number of cells were counted in a Neubauer chamber. Table 1. Average cell density, pH and temperature of cultivation in effluent media with and without the addition of CO2 . Cell density (×106 cells mL−1 ) pH

Temperature (◦ C)

Effluent without addition of CO2 4.45 0.00 0.00 7.28 1.40 19.23 8.40 0.74 8.82 8.89 1.07 12.09 9.02 3.22 35.67 9.11 3.99 43.82 7.88 5.48 69.50 7.94 4.75 59.85

7.85 8.53 8.70 9.04 8.65 8.61 8.45 8.20

25.50 26.50 27.50 28.00 28.00 28.00 29.50 29.00

Effluent with addition of CO2 4.45 0.00 0.00 8.39 1.29 15.43 13.90 2.72 19.58 19.26 4.10 21.28 26.66 2.13 7.99 34.52 2.71 7.84 42.48 5.91 13.92 40.65 2.32 5.70

7.13 6.15 5.65 5.30 5.03 4.56 5.20 5.33

26.50 28.00 27.00 27.50 29.00 28.00 27.50 28.00

Average

s±

CV (%)

Note: s±, standard deviation; CV (%), coefficient of variation. The averages for cell density were subjected to the Mann–Whitney U -test (p < 0.05) with p = 0.018.[26]

Table 1 presents the pH, temperature and cell density data collected for the experiment conducted in effluent medium with and without the addition of CO2 . Growth curves are presented for the tested media, both with and without the addition of CO2 , in Figure 2 for visualization of the data trends. In the cultivation of D. subspicatus without the input of CO2 , the MCD was 9.11 × 106 cells mL−1 , which was reached on the sixth day of cultivation. For the cultivation with supplemental of CO2 , the MCD was 42.48 × 106 cells mL−1 , which was reached on the seventh day of cultivation.

Environmental Technology Table 2.

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Analytical characterization parameters of the WTP-UNISC effluent prior to inoculation of D. subspicatus in the photobioreactor. Collection

Parameter Bicarbonate alkalinity Carbonate alkalinity BDO5 COD Total phosphorus Ammonia nitrogen TKN pH

Unit

1st

2nd

3rd

4th

5th

6th

Average

s±

CV (%)

mg L−1 (CaCO3 ) mg L−1 (CaCO3 ) mg L−1 (O2 ) mg L−1 mg L−1 mg L−1 mg L−1

50.50 282.80 20.00 114.00 10.10 36.10 44.50 7.85

133.80 202.10 33.00 283.00 11.30 54.30 70.70 7.78

45.60 307.80 81.00 200.00 4.30 45.60 45.60 7.80

55.50 262.60 22.50 82.00 11.20 56.40 52.50 7.47

85.90 275.50 41.50 103.00 3.75 19.10 22.10 6.79

45.60 248.60 131.00 113.00 1.60 65.80 77.20 7.20

69.48 263.23 54.83 149.17 7.04 46.22 52.10 7.48

34.94 35.98 43.36 77.05 4.31 16.68 19.88 0.42

50.28 13.67 79.07 51.65 61.16 36.08 38.16 5.61

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Note: s±, standard deviation; CV (%), coefficient of variation.

The average values for cell density were subjected to the Mann–Whitney U -test (p < 0.05) with p = 0.018, which confirmed a significant difference between the treatments. This result points to carbon as a limiting growth element, which makes carbon attractive for use in the fixation of atmospheric CO2 , such as that from the burning of coal in power plants. With respect to the MCD of the cultivation without the addition of CO2 , the above result points to a possible lack of carbon in the effluent. The pH values corroborate the MCD results. In the case of cultivation without supplemental CO2 , the pH varied slightly, which indicates low photosynthetic activity over time. In the cultivation with the input of CO2 , the culture medium acidified, with the pH reaching a minimum of 4.56 and increasing to 5.33 at the end of the cultivation. According to Esteves,[27] inorganic carbon in an aqueous medium may be in the form of CO2 , H2 CO3 (carbonic 2− acid), HCO− 3 (bicarbonate) or CO3 (carbonate), and the proportions of these species depend on the pH. With an increase in pH (alkalinization), the proportions of bicarbonate and carbonate increase in the culture medium according to the equilibrium for the neutralization of H2 CO3 . Thus, in an acidic culture, there is a greater availability of CO2 , which constitutes the preferred carbon source for microalgae because this compound rapidly diffuses (through passive adsorption) from the water into the cells and is used directly in fixation processes. Bicarbonate is actively incorporated, which necessitates an expenditure of energy to support this process.[28,29] The pH increased in the cultivation without the input of CO2 . The opposite phenomenon occurred in the cultivation with CO2 , where the pH decreased to approximately 5. 3.2.

Physicochemical analysis of the WTP-UNISC effluent Table 2 presents the results obtained for the analytical characterization parameters of the effluent before it received the inoculum of D. subspicatus in the photobioreactor. The first, second and third collections were from the initial

effluent used in the first three repetitions without the addition of CO2 . The fourth, fifth and sixth collections were from the initial effluent used in the culture with the addition of CO2 . The effluent exhibited great variations in its composition with respect to the water quality parameters, as shown in Table 2. The intervals between collections represent terms in which large number of students were on campus (collections 1–3) and the beginning of the holiday period (collections 4–6), when the number of users associated with the generation of waste was reduced. The reduced amount of waste also reduced the supply of nutrients to the WTP-UNISC effluent. Table 3 gives the analytical characterization parameters of the WTP-UNISC effluent before (initial) and after (final) the cultivation of D. subspicatus in the photobioreactor, first without the input of CO2 and subsequently with the input of CO2 . The hydraulic retention time (HRT) was seven days. The alkalinity of the effluents increased from the beginning to the end of the experiment. According to Hill and Bolte [30] and Bjornsson et al.,[31] anaerobic urban/domestic residual effluents, similar to the effluent used in this study, present high values of alkalinity, and bicarbonate is their major constituent. Autotrophic microalgae preferably require inorganic carbon for their growth, with CO2 being the preferred form used in photobioreactors for microalgae.[32,33] In its absence, the bicarbonate ion is preferably used. However, according to Park et al.,[34] in the presence of NH+ 4, the assimilation of bicarbonate is an active process (i.e. it requires energy). The presence of CO2 is therefore necessary to promote the continued assimilation of NH+ 4 . This assimilation was observed in the culture with the input of CO2 , where the N-NH3 value was reduced. In this experiment, the diaphragm compressor of the first tube pumped ambient air that contained CO2 from the natural environment into the system. The inoculum was adapted to these conditions (carbon source). However, the input of CO2 with a compressor only becomes negligible in the volume used in this work (30 L). As such, the removal of NH+ 4 was impaired in the culture without the input of CO2 .

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Table 3. Analytical characterization parameters of the WTP-UNISC effluent before (initial) and after (final) the cultivation of D. subspicatus in the photobioreactor with an HRT of seven days. Initial

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Parameter

Unit

Culture without the addition of CO2 Bicarbonate alkalinity mg L−1 (CaCO3 ) Carbonate alkalinity mg L−1 (CaCO3 ) BDO5 mg L−1 (O2 ) COD mg L−1 Total phosphorus mg L−1 Ammonia nitrogen mg L−1 TKN mg L−1 pH Culture with the addition of CO2 Bicarbonate alkalinity mg L−1 (CaCO3 ) Carbonate alkalinity mg L−1 (CaCO3 ) BDO5 mg L−1 (O2 ) COD mg L−1 Total phosphorus mg L−1 Ammonia nitrogen mg L−1 TKN mg L−1 pH

Final

Average

s±

CV (%)

Average

s±

CV (%)

p < 0.05

76.633 264.233∗ 44.667 198.5∗ 8.567 45.333∗ 53.600 7.810

49.568 55.242 32.130 84.504 3.743 9.103 14.819 0.036

64.683 20.906 71.933 42.465 43.698 20.080 27.648 0.462

107.900 356.133∗ 52.567 106.6∗ 9.133 71.266∗ 70.500 8.197

83.664 95.296 30.882 42.930 3.420 7.891 12.083 0.208

77.538 26.758 58.748 40.500 37.440 11.072 17.140 2.532

0.400 0.049 0.700 0.035 0.700 0.041 0.200 0.100

62.333∗ 262.233∗ 65.000 99.333∗ 5.517 47.100 50.600 7.153

21.001 13.454 57.942 15.822 5.038 24.700 27.599 0.339

33.691 5.130 89.141 15.928 91.322 52.442 54.544 4.736

304.366∗ 326.433∗ 84.867 157.333∗ 6.400 42.833 45.800 5.787

105.105 33.595 61.128 57.830 4.613 25.787 20.316 0.799

34.532 10.292 72.028 36.756 72.079 60.203 44.359 13.800

0.035 0.049 0.400 0.040 0.400 0.900 0.900 0.100

Note: s±, standard deviation; CV (%), coefficient of variation. ∗ Significantly different at the confidence level of 95% (n = 3).

Because of the ionic balance of inorganic nitrogen in water, nitrogen can be present either in the form of ions (NH+ 4 ) or in a free non-ionized form (NH3 ). An increase in pH and temperature contributes to an increase in the nonionized form (NH3 ) and a reduction in the ionized form (NH+ 4 ), which is relevant because NH3 is extremely toxic. In fresh domestic sewage, approximately 60% of the nitrogen present is in the form of organic nitrogen, and 40% is in the form of N-NH3 .[35] Furthermore, the bacterial decomposition of protein material and the hydrolysis of urea transform organic nitrogen into NH+ 4 .[36] For sewage, the total Kjeldahl nitrogen concentration (TKN) in the effluent has been reported to be approximately 40–60 mg L−g of N; of this total concentration, approximately 75% is −NH3 and 25% is organic nitrogen.[37,38] According to Abeliovich,[39] the N-NH3 concentration in domestic sewage ranges between 70 and 80 mg L−1 , followed by complete decomposition of urea and proteolysis. NH+ 4 is harmless to microalgae because its intracellular transport is controlled by specific mechanisms that do not allow the accumulation of its excess concentration. NH3 freely penetrates biological membranes and is extremely toxic. A combination of 30 mg L−1 of free NH3 at pH 8.2, for example, can inhibit microalgae growth.[40] Martínez et al.,[41] who worked with Scenedesmus obliquus under conditions similar to the present study, found 100% removal of NH+ 4 after 188.33 h of culturing. Kim et al.,[42] who worked with Chlorella vulgaris in effluent with similar characteristics in a closed system, obtained 50% removal in 48 h, which was observed 24 h after the adaptation phase of the culture. Park et al. [34] observed

the removal of N-NH3 on the order of 6.46 mg L−1 day−1 from anaerobic digestion effluent by Scenedesmus sp. at a specific growth rate of 0.038 day−1 . At the beginning of the cultivation without the input of CO2 , exponential growth of the culture occurred, and the increase in photosynthetic rate promoted alkalinization of the medium by removal of CO2 and a shift in the balance. In addition, the temperature increased to a level slightly outside the ideal range for D. subspicatus. This increase in temperature may be related to the operation of the submersible pump, which was not switched off at any time. As highlighted by Grobbelaar,[43] the assimilation of nutrients depends on the key factors that influence microalgal growth, such as light, temperature and turbulence. Temperature, according to Goldman and Carpenter,[44] directly influences enzymatic reactions, and these reactions double with increasing temperature until a certain threshold, which is characteristic for each species, is reached. As mentioned previously, an increase in temperature and pH favours the fraction of free, toxic NH3 ; moreover, the initial values of N-NH3 were greater than 30 mg L−1 . The literature does not contain information concerning the cell development of D. subspicatus in effluents of this nature. The decline phase of the growth curve indicates cell death, either naturally or by hydrodynamic agitation. Cell death promotes the release of protein material, and as previously mentioned, the bacterial decomposition of the protein material and the hydrolysis of urea transform organic nitrogen into NH3 , which may have contributed to the increase in the N-NH3 found in the final analysis of

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Environmental Technology the effluent. However, additional microorganisms possibly present in the medium were not identified because of the exposure of this culture medium at the time of inoculation. The same behaviour was not observed in the experiment with the addition of CO2 , in which an insignificant decrease was observed in the amount of N-NH3 . These results imply that the presence of CO2 contributed to the continued assimilation of N-NH3 . According to Oliveira and Sperling,[45] a phosphorus concentration between 1 and 8 mg L−1 was observed in studies that compared the results of observed and expected total phosphorus concentrations in effluents treated with an upflow anaerobic sludge blanket (UASB) coupled with post-treatment. In contrast, the phosphorus concentration reported in the comparable literature was 4 mg L−1 . Approximate values were obtained by Sousa et al. [46] in domestic effluents treated with an UASB system, where the total phosphorus concentration varied from 5.6 to 7.0 mg L−1 . An increase was observed in the value of total phosphorus in both experiments, although the increase was not significant. This increase may be related to the release of phosphorus from organic matter that resulted from dead cells that were in suspension in the medium. Voltolina et al. [47] achieved a removal of phosphorus on the order of 50% in domestic effluent with Scenedesmus sp., and Zhang et al. [48] reported total phosphate removal by Scenedesmus sp. with 100% efficiency in domestic effluent during a similar time period. Finally, Larsdotter et al. [49] demonstrated that in wastewater treatment, phosphorus assimilation was dependent on algal biomass production. Relative to results published by Samorì et al.,[50] the cultivation of D. subspicatus under the tested conditions without the addition of CO2 was not efficient. This result was especially evident from the decline in growth observed after a brief period of inoculation. Note that the change in pH may have caused changes in the speciation of the compounds, which could not be assessed in this experiment. Additionally, a brief change in temperature from the ideal temperature for cultivation throughout the experiment may also have inhibited growth, given the importance of this parameter. It is assumed that effluent variability contributed to the discrepancies in phosphorus and may contain compounds toxic to microalgae. For cultivation in the presence of supplementary CO2 , a decrease in the total phosphorus in the effluent was observed in both the first and second repetitions. The latter was possibly due to the largest population growth in the cultivation, where a large number of individuals promoted the assimilation of nutrients. In addition, there is a sequestration of phosphorus by calcium content in the flocculation step in high pH. This effect has occurred in all samples because the calcium content in the effluent is near 5.5 mg L−1 due to the quality of water used in the restrooms and kitchens of the university (data from Central of Analysis – UNISC).

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From a kinetic study of a wastewater photobioreactor with C. vulgaris, Ruiz et al. [51] showed that it is possible to produce biomass and remove nutrients as well as could occur with synthetic media. Values for BOD5 and COD increased, except with respect to COD in the experiment without the addition of CO2 , indicating excretion of organic material by the culture rather than sequestration. Organic materials can be present as primary and secondary metabolites, containing carbon, nitrogen and phosphorus excreted by the cell during cultivation. As in Wang et al.,[52] situations similar to this one may indicate metabolic patterns for different species or heterotrophic and autotrophic growth under different conditions (i.e. nutrients, presence or absence of additional CO2 ). It is a common behaviour among species of Chlorophyta. Eny [53] reported that this metabolic pattern can be changed depending on the organic substrate present (e.g. organic acids, glucose), which shows that heterotrophic growth may occur concurrently with autotrophic CO2 as the sole carbon source. The effluent material containing carbon is predominantly inert after the activated sludge process and cannot be used by microalgae. When this scenario occurs, autotrophic growth uses CO2 as the carbon source, and the cells secrete substances of small molecular size, such as glycolic acid, to the environment as the photosynthetic product of reducing carbon.[54] This behaviour could explain the increase in the final value of BOD5/COD after cultivation. Working with Scenedesmus spp., Kim et al. [55] emphasized that the imbalance between N, P and C in batch cultivations can stimulate sequestration of organic carbon in an alternation between autotrophic–heterotrophic metabolism or stimulate the generation of pyrenoids associated with the enzymes of the Calvin cycle for CO2 fixation. Wang et al. [52] also emphasized that the HRT is very important in studies of this nature, and once the maximum growth is reached, the release of cellular material (nutrients) begins to increase, which may occur on the earliest days of cultivation. Therefore, a relatively long period after the cultivation time (maximal growth) can result in increased final reducing BOD5 and COD removal efficiency of the process. In this same study, the retention time could be decreased by introducing bacteria into the system, forming an algal–bacterial symbiosis, which can be used to study the dynamics of nutrients during the exponential growth phase. 3.3.

Biomass and lipid fraction

3.3.1. Fatty-acid composition The amount of lipids in microalgae varies from 1% to 70%; however, the lipid levels of most algae do not exceed 40%. The lipids contained in these organisms are commonly esters and fatty-acid chains that contain between 14 and 22 carbon atoms, both saturated and unsaturated.[40] The average lipid fraction found in the biomass was 18.73 ± 0.25% in the cultivation with the addition of CO2 and 12.00 ± 0.28% without additional CO2 . According to

216 Table 4. ments.

P.D. Gressler et al. Dry weight of D. subspicatus biomass in tested treat-

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Dry weight (mg L−1 ) Treatment

Average

s±

CV (%)

p < 0.05

Effluent Effluent + CO2

234.00 1277.44

91.89 1053.04

39.3 82.4

0.049 0.049

Becker,[56] algal lipids are typically composed of glycerol, sugars or esterified bases with between 12 and 22 carbons, and they may or may not be saturated. In another study, Allard and Templier,[57] working with the same species, found an average extracted oil content of 20% without the input of CO2 under optimum cultivation conditions. Table 4 gives the dry biomass weight for each experiment in mg L−1 . A significant difference was observed between the dry weight of the biomass between treatments with p = 0.049. The treatment with CO2 had a greater dry weight. This result is complemented by the difference in

the obtained cell densities. Table 5 shows the percent averages of fatty acids extracted from the acylglycerol of the D. subspicatus oil. Figure 3 shows the superposition of chromatograms from the experiments with and without the input of CO2 . For both experiments, despite the trends observed in dry weight, the fatty-acid profile was qualitatively similar according to the statistical result, with p < 0.05. Therefore, environmental factors, such as the quality of the effluent and the presence of extra CO2 , did not produce different profiles of fatty acids. Working with the same species without the addition of CO2 and under similar conditions of lipid extraction and chromatography, Allard and Templier [57] found high concentrations of the acids C16:0 (palmitic acid) and C18:1 (oleic acid). The main functions of lipids and fatty acids are in the formation of membranes and cell energy reserves. Most nonpolar lipids in microalgae are triacylglycerols and free fatty acids, whereas the polar lipids are mainly glycerides, phospholipids and glycolipids.

Table 5. Averages (%) of the fatty acids found in the acylglycerols of the extracted oil of the microalgae from experiments in the effluent with and without the input of CO2 . Treatment Analysis fatty acids (%) Myristic (C14:0) Palmitic (C16:0) Stearic (C18:0) Ômega-3 (C18:3) Oleic (C18:1) Eicosanoic (C20:0) Linoleic (C18:2)

Effluent without addition of CO2

Effluent with addition of CO2

Retention time (min)

Average

s±

CV (%)

Average

s±

CV (%)

p < 0.05

16 23.9 32.1 37.2 32.8 40.1 34.7

0.45 28.61 8.28 13.06 28.85 0.78 12.08

0.09 1.39 1.35 0.97 1.44 1.35 1.66

20.63 4.87 16.27 7.44 4.98 173.21 13.70

0.55 27.70 7.40 16.88 24.56 0.07 12.32

0.31 7.18 6.74 3.04 6.17 0.12 2.15

55.91 25.93 91.07 18.04 25.13 173.21 17.46

>0.999 0.7 >0.999 0.2 0.1 >0.500 >0.999

Note: s±, standard deviation; CV (%), coefficient of variation.

Figure 3. Chromatograms of methyl esters related to the fatty acids present in the oil of D. subspicatus produced in effluent with and without the input of CO2 .

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Environmental Technology The biomass analysis of D. subspicatus presented similar fatty-acid profiles under both tested conditions. However, the quantity of stored oil with the presence of CO2 varied. The fatty acids that were observed, as well as the oil content, agree with the previously cited work of Allard and Templier.[57] The same cannot be said for the cultivation with CO2 because the literature does not contain results for the same species under similar cultivation conditions. Considering that other factors can be responsible for the oil quality, we chose the flocculation conditions for preliminary tests. The NaOH flocculation method was used because no differences have been observed between the fatty-acid profiles resulting from electroflotation [58] or flocculation with NaOH at a small scale.[59] NaOH flocculation is simpler than electroflotation, but it is not as rapid. Borges et al. [60] found that cationic and anionic flocculating agents may change the amount of fatty acids present in microalgae biomass compared with the NaOH flocculant. According to these authors, it is very unlikely that the addition of NaOH caused the observed differences in the fatty-acid profiles in treatments with anionic and cationic flocculants. Moreover, it is also possible that NaOH had removed phosphates from the effluent, but the effect was similar in all samples. For bioremediation, NaOH can be considered auxiliary to phosphate flocculation. 3.3.2. Biomass production potentialities According to a personal communication from the technician responsible for the WTP-UNISC, the WTP-UNISC releases 2.42 million litres of effluent into the receptor body of water every month in a batch system. With an estimate of seven days of HRT in the photobioreactor and under the proposed acclimation conditions (in the cultivation room, not in the field), an average work volume of 605,000 L of microalgae cultivation per week would be needed. This work volume would result in approximately 1470 kg of biomass per week. Because the biomass averaged 166.66 mg L−1 in the cultivation with the input of CO2 , it would be possible to obtain approximately 100.40 kg of D. subspicatus oil weekly. The use of this oil in the production of biodiesel would still require an increase in production for the investment to be viable; alternatively, the 1470 kg of biomass could be used for the production of ethanol. Several species of the genera Chlorella, Dunaliella, Chlamydomonas, Scenedesmus and Spirulina contain approximately 50% dry weight of biomass composed of starch and glycogen, which are commonly used as raw materials for the production of ethanol.[61] In the case of microalgal biomass, the existence of complex carbohydrates in the cell wall is known, and these carbohydrates must be converted into simple sugars to enable the action of microorganisms such as yeasts and to produce bioethanol.[62,63] Nguyen et al. [64] reported the acquisition of 58% obtained glucose and approximately 29%

217

of the dry weight converted to ethanol with the species Chlamydomonas reinhardtii. 4.

Conclusions

The results of this study showed the potential and evolution of the use of microalgae, especially D. subspicatus, in the cultivation of WTP-UNISC effluent with concomitant CO2 fixation for the production of biomass. The main findings of the study are summarized as follows: • A tubular photobioreactor with a bubble column in a semicontinuous gas regimen was developed for the production of D. subspicatus biomass. • The cultivation of D. subspicatus in the proposed photobioreactor was possible by inoculation in the WTP-UNISC effluent with the addition of CO2 , achieving an average value of 1277.44 mg L−1 . • The fatty-acid profiles and oleaginous potentials of D. subspicatus cultivated in a tubular photobioreactor with and without the input of CO2 were qualitatively similar. Differences in the quality of the effluent and the presence of CO2 did not result in different fatty-acid profiles. The presence of C16:0 (palmitic acid) and C18:1 (oleic acid) was notable. The extracted oil contents were, on average, 18% and 12% for cultivation with and without the input of CO2 , respectively. Acknowledgements We would like to thank CAPES and AES – Uruguaiana for the scholarships given to P.D.G., M.P.S. and T.R.B. We gratefully acknowledge the support provided by AES – Uruguaiana – and CNPq. We additionally thank the FAP/UNISC program for research support.

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