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Growth of oil accumulating microalga Neochloris oleoabundans under alkaline–saline conditions
Bioresource Technology 104 (2012) 593–599
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Growth of oil accumulating microalga Neochloris oleoabundans under alkaline–saline conditions A.M. Santos a,b,⇑, M. Janssen b, P.P. Lamers b, W.A.C. Evers b, R.H. Wijffels b a b
Wetsus – Center of Excellence for Sustainable Water Technology, P.O. Box 1113, 8900 CC Leeuwarden, The Netherlands Bioprocess Engineering, Wageningen University and Research Center, P.O. Box 8129, 6700 EV Wageningen, The Netherlands
a r t i c l e
i n f o
Article history: Received 28 July 2011 Received in revised form 20 October 2011 Accepted 22 October 2011 Available online 6 November 2011 Keywords: Neochloris oleoabundans Alkalinity Growth rate Lipid accumulation Nitrogen starvation
a b s t r a c t The effect of elevated pH and salt concentration on the growth of the freshwater microalga Neochloris oleoabundans was investigated. A study was conducted in 24-well plates on the design of a growth medium and subsequently applied in a photobioreactor. An artificial seawater medium with reduced Ca2+ and PO34 could prevent mineral precipitation at high pH levels. Growth was characterized in this new medium at pH 8.1 and at pH 10.0, with 420 mM of total salts. Specific growth rates of 0.08 h 1 at pH 8.1 and 0.04 h 1 at pH 10.0 were obtained under controlled turbidostat cultivation. The effect of nitrogen starvation on lipid accumulation was also investigated. Fatty acids content increased not only with nitrogen limitation but also with a pH increase (up to 35% in the dry biomass). Fluorescence microscopy gave visual proof that N. oleoabundans accumulates oil bodies when growing in saline conditions at high pH. Ó 2011 Elsevier Ltd. All rights reserved.
1. Introduction The use of microalgal lipids as a feedstock for a sustainable production of biofuels is a hot topic within the scientific community. This is related to the high lipid content of some species and to the fact that lipid synthesis can be easily modulated by manipulating their cultivation conditions. Moreover, due to their high photosynthetic efficiency and faster growth rates in comparison to higher plants, lipid-rich microalgae can be considered as potentially one of the most efficient biological oil producers (Chisti, 2007; Li et al., 2008a,b; Mata et al., 2010; Wijffels and Barbosa, 2010). The total composition of lipids in microalgae can vary between 1% and 85% of their dry weight (Chisti, 2007) achieving values up to 40–70% under nutrient limitation (Roessler, 1990; Rodolfi et al., 2009). Nevertheless, a microalgae-based fuel production technology which can economically compete with the ones currently on the market still needs to be developed. The main limitations still associated with this technology are the selection of strains which can grow and produce high quantities of lipids at the same time, and the high energy inputs necessary for pumping, mixing and harvesting the produced biomass (Hu et al., 2008; Rodolfi et al., 2009; Norsker et al., 2011).
⇑ Corresponding author at: Bioprocess Engineering, Wageningen University and Research Center, P.O. Box 8129, 6700 EV Wageningen, The Netherlands. Tel.: +31 (0) 317 48 50 12; fax: +31 (0) 317 48 22 37. E-mail address: [email protected] (A.M. Santos). 0960-8524/$ - see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.biortech.2011.10.084
Substantial energy is required for the supply of carbon dioxide (CO2) to large-scale cultivation systems by means of gas transfer. During microalgal cultivation, CO2 is usually introduced to the system by CO2 enrichment of the inlet air flow, but it is envisioned to use CO2-rich gasses as for example flue gas (Zeiler et al., 1995; Brown, 1996). However, a considerable amount of CO2 which fails to be transferred into the liquid phase will still be wasted to the effluent gas. When growing microalgae at a pH higher than 8.5, the cultivation medium could be saturated with inorganic carbon as bicarbonate (HCO3 ) prior to entering the cultivation system. The energy necessary for CO2 transfer from the gas to the liquid phase could thus be reduced by optimizing CO2 transfer in a separate gas transfer unit at elevated pH (>8.5). Moreover, also in traditional systems where CO2 is supplied in the actual growth stage the efficiency of CO2 transfer could be increased when operating the system at a pH of 10 or more (Yagi and Yoshida, 1977). Such process strategies are only possible if there are microalgae able to efficiently grow and produce lipids under alkaline conditions at high salt concentrations (i.e. bicarbonate, carbonate, and associated counter-ions). The alkalinity of an aqueous solution is regulated by the distribution of the bicarbonate and carbonate species (HCO3 and CO23 ). These salts also determine the system’s buffering capacity. For this reason, they must be present at high enough levels to supply the necessary CO2 for microalgal growth and to keep the pH from increasing too much. Algal growth in culture also depends on the adequate supply of essential macro and micronutrients. These nutrients are usually
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highly soluble salts which dissociate in ions and are either present in high concentrations, as in the case of magnesium (Mg2+) and calcium (Ca2+) in seawater, or can be added in amounts sufficient to support good algal growth. At high pH (pH > 8.0) especially divalent ions have the tendency to precipitate with phosphate or hydroxide as insoluble salts. For this reason, the concentrations of ions such as Ca2+, Mg2+ and phosphate (PO34 ) should be kept as low as possible without limiting microalgae growth. The accumulation of large quantities of lipids by microalgal species has already been reported (Williams and Laurens, 2010). Among these species is Neochloris oleoabundans, known as a freshwater species capable of producing oil bodies composed of triacylglycerides (TAG) at a level of up to 80% of its total lipid composition. Most of those lipids are composed of saturated fatty acids (Tornabene et al., 1983), suitable for the production of biofuels. A more recent screening study performed by Gouveia and Oliveira (2008) also showed the potential of N. oleoabundans as a source of lipids for biofuels production. A considerable increase in oil quantity (50% of the dry weight) was claimed during growth under N-depletion. However, the possible application of N. oleoabundans for biofuels production under alkaline and saline conditions has not been covered in literature so far. The present work aimed to test whether N. oleoabundans could also be grown under alkaline and saline conditions while still being able to accumulate lipids. Experiments were performed in microtiter plates and in a lab-scale photobioreactor to determine the specific growth rate of this microalga under a range of alkalinities and salinities. Besides nutrient-replete growth, nitrogen deprivation was applied to induce lipid accumulation and to check whether this accumulation was affected by the alkalinity and the salinity of the growth medium. Moreover, the development of a stable inorganic growth medium for alkaline–saline conditions is discussed. 2. Methods 2.1. Microalgal strain N. Oleoabundans UTEX 1185 was pre-cultivated in 250 mL shake flasks containing 100 mL of medium. The cultures were placed on an orbital shaker incubator (Innova 44, New Brunswick Scientific, New Jersey, USA) under a 2% CO2-enriched headspace and a light intensity of 30 lmol photons m 2 s 1 provided by fluorescent lamps. Temperature was controlled at 25 °C and pH was 7.8. 2.2. Medium design A seawater-type medium (Table 1) was developed based on the elemental biomass composition of several marine species (Ho et al., 2003) and designed to support at least 1 g L 1 of biomass. Non-biological precipitation experiments were performed in 24well microtiter plates (Falcon 3047, BD Biosciences, CA, USA). The plates were fixed inside the incubator and rotated at 150 rpm (orbit 2.54 cm). In a total volume of 1.5 mL per well, different Mg2+/ Ca2+ combinations in the medium were tested. The ratio between both elements was either kept constant (13:1), or varied with a constant minimal Ca2+ concentration (0.01 mM). Nitrate was kept in a ratio of 14:1 with phosphate, which was added as a potassium polyphosphate solution insensitive to high pH levels (Super FK, YARA, Vlaardingen, The Netherlands). Six different pH values were tested in duplicate for each medium composition: 9.0, 9.5, 10.0, 10.3, 10.5 and 10.8. The desired pH was obtained by buffering with different 300 mM NaHCO3–Na2CO3 mixtures, as described in Table 2. The stability of the medium was assessed by following the optical density at 750 nm (OD750) using
Table 1 Modified medium composition (in 1 L). X and Y are the concentrations of magnesium and calcium in artificial seawater, respectively. Values for these concentrations were determined in this study and are given in Section 3.1. Compound
[mM]
Macronutrients KNO3 Super FK EDTA ferric sodium salt
4.4 0.32 29.28 lM
Vitamins Thiamine Biotin B12
200 lg L 1 lg L 1 1 lg L 1
Micronutrients EDTA-MnNa2 EDTA-ZnNa2 EDTA-CuNa2 Na2MoO42H2O CoCl26H2O
1.5 2.0 7.0 7.7 3.7
Artificial seawater NaCl MgCl26H2O CaCl22H2O Na2SO4
420.0 X Y 22.53
1
E-02 E-03 E-03 E-04 E-04
a multilabel plate reader (VICTORTM X3, PerkinElmer, MA, USA) and wells with demineralised water as blanks. An increase in optical density signifies salt crystals formation due to the light scattering by these insoluble particles. The growth of N. oleoabundans on the modified medium was studied under the same conditions, except for the additional supply of light (fluorescent light tubes, 30 lmol photons m 2 s 1) and CO2 (4% in headspace). Temperature was controlled at 30 °C. The wells (total volume 1.5 mL) were inoculated with 75 lL from 2 weeks old shake flask cultures. The optical density at 750 nm was measured daily to follow biomass growth (medium served as control).
2.3. Growth assessment on alkalinity and salinity The influence of a range of alkalinities and salinities on growth was also studied in 24-well microtiter plates. The wells (total volume 1.5 mL) were inoculated with 150 lL from 2 weeks old shake flask cultures and incubated as described in Section 2.2, with the exception that the light intensity was set at 55 lmol photons m 2 s 1. Microalgal growth was assessed at four pH values: 8.1, 9.0, 10.0 and 10.5, which were set by applying different NaHCO3–Na2CO3 buffer mixtures (Table 3) and by adjusting the CO2 fraction in the headspace. In four sets of experiments, different concentrations of NaCl in the medium were combined with the NaHCO3–Na2CO3 mixtures to check the algae’s salt tolerance Table 2 Molar concentrations of NaHCO3 and Na2CO3 in the buffering mixtures used during the medium design experiments. For each pH tested, the combined concentration of both salts was 300 mM. pH
NaHCO3 [mM|
Na2CO3 [mM]
9.0 9.5 10.0 10.3 10.5 10.8
270 210 150 90 60 30
30 90 150 210 240 270
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Table 3 Molar concentrations of NaHCO3 and Na2CO3 in the buffering mixtures used during growth assessment experiments. For each pH tested, the combined concentration of both salts in the medium was either 300 mM or 150 mM. pH
Total 300 [mM]
9.0 10.0 10.5
Total 150 [mM]
NaHCO3 [mM]
Na2CO3 [mM]
NaHCO3 [mM]
Na2CO3 [mM]
270 150 60
30 150 240
135 75 30
15 75 120
(Table 4). At pH 8.1 medium with solely 12 mM NaHCO3 was used. All sets of experiments were performed in four replicates. Repeated batch cultivation was applied to assess the specific growth rates for each medium. Every 24 h the cultures were diluted to the starting optical density (as represented in Fig. 1), whilst keeping the culture volume at 1.5 mL. The dilution series were continued until a constant specific growth rate was observed. 2.4. Growth assessment in a bench-scale panel photobioreactor The microalgae were grown in a flat panel photobioreactor (FMT150, Photon Systems Instruments, Brno, Czech Republic) with a detachable rectangular cuvette 20 cm high, 10 cm wide and 2.5 cm optical path (working volume 385 mL). The photobioreactor (PBR, Fig. 2) was equipped with temperature control (set to 30 °C) and a light source composed of red light emitting diodes, LEDs (kmax 627 nm). The incident light flux was measured at 15 different places equally distributed over the PBR’s light-exposed surface with a LI190 2p PAR quantum sensor (Li-CORÒ, Lincoln, USA). The average light intensity at the surface was set to 300 lmol photons m 2 s 1. A built-in dual-wavelength densitometer (680 nm and 735 nm) allowed us to monitor the culture growth and the chlorophyll absorption using the PBR itself as cuvette. Mixing was performed by continuous sparging at 650 mL min 1 of a CO2-enriched air stream through a gas-distributor over the bottom of the panel. The gas was saturated with water before entering the vessel. The influence of the pH on growth was studied at pH 8.1 and pH 10.0. Growth was started batch wise using 30 mL inoculum from a 2 weeks old shake flask culture and 355 mL medium with optimal Ca2+ and Mg2+ salt concentrations (X and Y in Table 1, see Section 3). The concentration of NaCl in the medium was 420 mM at pH 8.1 and 270 mM at pH 10.0. At pH 8.1, 12 mM NaHCO3 was in equilibrium with 1% v/v CO2. At pH 10.0, a mixture of 75 mM NaHCO3 and 75 mM Na2CO3 was applied, while gassing with 0.1% v/v CO2. An OD735 of 0.2 was defined as the start and set-point for the turbidostat control. In turbidostat mode, the OD735 was maintained constant by the automatic addition of fresh medium and concomitant harvesting of surplus culture. The dilution rate (D), being equal to the specific growth rate of the microalgae, was assessed daily. The correspondent biomass concentrations (Cx) were determined by dry weight measurements. At both pH the algae were deprived of nitrogen, after a constant growth rate at nitrogen-replete conditions was obtained, to induce the accumulation of lipids. To this end, the feed medium was replaced with
Fig. 1. Schematic representation of dilution steps in 24-well microtiter plates.
nitrogen-free medium resulting in a gradual but complete depletion of nitrogen in the reactor. 2.5. Biomass dry weight determination The biomass concentration was determined according to Zhu and Lee (1997). Triplicate samples (10 mL) were diluted in ammonium formate (0.5 M) and filtrated with a constant vacuum (0.44 bar, absolute) over pre-dried and pre-weighted Whatman GF/F glass fiber filters (Ø 55 mm, pore size 0.7 lm). The filters were rinsed twice with ammonium formate, dried (24 h, 95 °C), cooled down in a dessicator (2 h) and weighted again. The biomass dry weight was determined as the difference between the weights of the dried filters containing the samples and the dried empty filters. 2.6. Visualization of intracellular neutral lipid bodies The accumulation of intracellular lipid bodies was checked by fluorescence microscopy. The lipophilic fluorescent dye BODIPY 505/515 (4,4-difluoro-1,3,5,7-tetramethyl-4-bora-3a,4a-diazasindacene; Invitrogen Molecular Probes, Carlsbad, CA) was used for staining the cells, according to Cooper et al. (2010). A 40 lM BODIPY 505/515 stock solution was prepared by dissolving the dye in 0.2% (v/v) of anhydrous dimethyl sulfoxide (DMSO). Aliquots of this solution were added directly to the suspensions, having a final dye concentration of 1 lg mL 1 and 0.02% DMSO. After 5 min of contact, slides were prepared for observation under an inverted fluorescence microscope (Leica DMI6000, Leica Microsystems B.V., Rijswijk, The Netherlands). Excitation was done under a blue excitation light through a band-pass filter (450–490 nm) and emission wavelengths were imaged through a long-pass filter (515 nm). 2.7. Fatty acids quantification
Table 4 Sodium chloride combinations with the NaHCO3 and Na2CO3 buffering mixtures tested during growth assessment experiments at different pH values. Set
NaCl [mM]
NaHCO3 + Na2CO3 [mM]
1 2 3 4
420 – 120 270
300 300 300 150
Duplicate reactor samples were centrifuged for 5 min at 2500 rpm and 15 °C and the cell pellets were stored at 80 °C. On a later stage, the pellets were transferred to bead beating tubes (Lysing Matrix E, MP Biomedicals) and lyophilized overnight followed by gravimetric biomass dry weight determination. Upon addition of a chloroform/methanol mixture (2:2.5), containing 47 lg mL 1 of the internal standard tripentadecanoin (Sigma– Aldrich T4257), complete cell disruption was achieved by bead
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Fig. 2. Schematic representation of photobioreactor FMT150. The green parts correspond to the additional connections when operating in turbidostat mode.
beating for 40 min. The homogenates were then transferred to 12 mL glass tubes. Complete recovery of the homogenate was assured by rinsing the bead beating tubes with an additional 3 mL of the chloroform/methanol mixture. The next steps of the extraction were adapted from a protocol optimized by Lamers et al. (2010) and are available as supplemental material. Gas chromatography (GC) analysis was done using a HP 6890 (Hewlett Packard Inc.) with FID detector. A volume of 1 lL was injected (split ratio 0.1:1) onto a Supelco NucolTM column (355 33-03A, 30 m 530 lm 1.0 lm film thickness) with helium as the carrier gas (constant total column flow 20 mLmin 1). The column oven was programmed at an initial temperature of 90 °C for 0.5 min, followed by a ramp of 20 °Cmin 1 to 200 °C and a final step of 24 min at 200 °C. The FAMEs were identified by comparison of their retention times with those of authentic standards (Sigma Aldrich, MO, USA). Quantification of the identified lipids was done by correlating their individual peak areas to the one of the internal standard, taking into account the amount of internal standard added per sample and the response factors that were calibrated for each fatty acid using authentic standards. Samples that did not contain biomass, but were otherwise subjected to the entire protocol, served to correct for three minor peaks that were extracted from the plastic bead beater tubes.
3. Results and discussion 3.1. Growth of N. oleoabundans in a modified medium The preparation of cultivation media with high concentrations of salts and high pH needs careful attention regarding the issue of precipitation. Compounds which precipitate are no longer available for microalgal growth. For instance, the binding capacity of calcium to phosphate is pH dependent, increasing at elevated values. A chemical equilibrium model for the calculation of saturation indices in aqueous solutions (Visual MINTEQ, Gustafsson, 2009) was the base for developing a medium with low levels of precipitation. A simulation showed for example high formation of a calcium phosphate complex, hydroxyapatite, for our modified medium at pH 10.5 when containing Ca2+ and PO34 in concentrations of 3.61 mM and 5 mM, respectively. One way to reduce the formation of hydroxyapatite is to decrease the concentrations of Ca2+ and/or PO34 . It is difficult to get reliable data on the need of calcium for microalgae growth. Our non-biological experiments showed that using Ca2+ in a concentration of 0.01 mM results in a medium with very low complexation levels at high pH values. Besides calcium, also
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magnesium can complex and precipitate at high pH, though at a lesser degree. Furthermore, the amount of magnesium needed by the cells seems to be higher than calcium. Magnesium is essential for cell growth, while the need for calcium can be questionable (Walker, 1994). In our experiments, a ratio of 1:15 between Ca2+ and Mg2+ could be translated in minimal precipitation, magnesium being in a concentration of 0.15 mM. Based on these findings we developed a new medium with 0.01 mM CaCl2 (X in Table 1), 0.15 mM MgCl2 (Y in Table 1), 4.4 mM KNO3 and 0.32 mM ‘P’ from the polyphosphate solution (Super FK). Additional experiments indeed confirmed that this medium resulted in negligible precipitation. In Fig. 3A the optical density shows the level of precipitation at different pH values. In the first 100 h the OD hardly increases. Only at elevated pH (>10) and after extended time (150 h) limited precipitation occurs. For continuous microalgae production this should not cause any problems since the rate of consumption will be faster than the rate of precipitation. Moreover, continuous consumption will result in even lower concentrations of Ca2+, Mg2+ and PO34 , further reducing the risk of precipitation. Although in the new medium there was sufficient nitrogen and phosphate available to sustain a biomass concentration of 1 g L 1 it was not known whether the final concentrations of calcium and magnesium were sufficient to support algae growth up that value. However, during experiments with N. oleoabundans growing batch wise in this modified medium in the FMT150 photobioreactor (work not presented) a final biomass concentration of 1.8 g L 1 was obtained, confirming the calculation that at least 1 g L 1 can be sustained by this cultivation medium. Fig. 3B shows the increase in optical density during growth in the modified medium at different pH values. As expected, growth of N. oleoabundans was not limited by the low concentrations of Ca2+ and Mg2+, although there was a general decrease with the increase of pH, especially at values above 10.0. 3.2. Influence of pH and salt concentration on specific growth rate 3.2.1. Microalgae growth in 24-well microtiter plates The results presented in the previous section show that growing N. oleoabundans in a saline medium with a high pH is possible. In this section it is discussed how the combination of pH and salt concentration affects the growth of this microalga. For all the pH values tested (Table 3), no growth of N. oleoabundans was observed with the highest total salt concentration of
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720 mM (Set 1, Table 4). Furthermore, very poor growth was observed at pH 10.5, for all sets of experiments. Growth rates were not determined for this value. At higher pH levels with less HCO3 /CO23 (total 150 mM), the combination with more NaCl (270 mM/Set 4) seems to be better for the algae than with less NaCl (absent/Set 2 and 120 mM/Set 3) combined with more HCO3 /CO23 (total 300 mM). A medium with 270 mM NaCl and 150 mM HCO3 / CO23 was therefore the only formulation capable of supporting growth at pH 10.0. An average growth rate of 0.0212 h 1 was found for the reference experiment at pH 8.1 (Fig. 4A), being the highest amongst all the experiments. However, this value was not considerably different for the one found at pH 10.0, where the specific growth rate reached 0.0177 h 1 (Fig. 4B). A correlation between the pH and the salt concentration seems evident as both have an impact on microalgal growth. Although the specific growth rates are all relatively low, due to light limitation inside the incubators (55 lmol photons m 2 s 1), they give a good insight on which should be the best conditions to grow N. oleoabundans at high pH: a marine medium with 270 mM NaCl, 75 mM NaHCO2 and 75 mM Na2CO3. 3.2.2. Microalgae growth in turbidostat cultivation and lipid accumulation under nitrogen depletion To estimate the growth under high pH and salt concentration under controlled conditions in a continuous system, two turbidostat experiments were performed at pH 8.1 and pH 10.0. The light intensity applied was of saturating nature (300 lmol photons m 2 s 1) and allowed to assess the maximum specific growth rate without any light limitation. The total salt concentration was 420 mM according to the study performed in the microtiter plates. The respective biomass concentrations (Cx) and dilution rates (D) were determined under steady-state. In steady state the specific rate of cell growth equals the rate of dilution. Growth rates of 0.08 h 1 and 0.04 h 1 were found at pH 8.1 and pH 10.0 (Fig. 5A and B, respectively). The biomass concentration Cx was constant under steadystate at 0.1 g L 1 in both experiments. Although the growth rate we obtained at pH 10.0 was twice as small as that at pH 8.1, it is still comparable with the one found by Pruvost et al. (2009). In their work, they reached a biomass concentration of around 0.1 g L 1 at a specific growth rate of 0.05 h 1 under the same light intensity, but in freshwater medium (bold basal medium) at pH 7.5 and 25 °C. The influence of nitrogen starvation on the accumulation of lipids was also investigated. Nitrogen starvation is known to be a
Fig. 3. Optical density at 750 nm (OD750) of (A) modified medium at different pH values where the increase in OD750 is a measure of medium precipitation and (B) N. oleoabundans in the modified medium at different pH values as a measure of growth. Lowest and highest pH values tested are represented by white and black triangles, respectively. Error bars represent the standard deviation from the average of the replicates.
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Fig. 4. Growth of N. oleoabundans in dilution series at (A) pH 8.1 and (B) pH 10.0. Growth rates determined 24 h after each dilution step.
Fig. 5. Growth of N. oleoabundans in turbidostat cultivation at (A) pH 8.1 and (B) pH 10.0. The vertical dotted lines represent the point after what depletion of nitrogen from the medium was applied.
strong inducer of lipid accumulation (Tornabene et al., 1983; Li et al., 2008a,b; Pruvost et al., 2011; Yeh and Chang, 2011). After reaching a constant growth rate, the cultivation medium was replaced by nitrogen-free medium. This resulted in a sharp decrease in l, to 0.02 h 1 at pH 8.1 and 0.01 h 1 at pH 10.0, and this decrease is obviously caused by the absence of nitrogen.
Cells growing in the photobioreactor at pH 8.1 and 10.0 were also analyzed for their total fatty acid content and composition, both under nitrogen-replete conditions and after 1 day of nitrogen depletion. Table 5 shows the microalgal content in each of the identified fatty acid, as well as their relative percentage of the total fatty acid pool. As we expected, there was an increase in the total fatty acid content of the nitrogen-starved cells one day after switching from N-replete medium to N-free medium. This increase was even more pronounced for the cells growing at pH 10.0 (up to 35% of dry weight), which may suggest that an additional stress such as high pH could promote lipid accumulation. A similar conclusion was taken by Guckert and Cooksey (1990) when they studied the accumulation of TAG during the growth of Chlorella at high pH. They suggested that such high pH-related stress can even be nitrogen independent, resulting in cell-cycle inhibition and triggering TAG accumulation. This was recently supported by Gardner et al. (2010), who suggested that high pH and nitrogen depletion have a synergistic effect on TAG accumulation in microalgae. But, the fatty acid contents of microalgae are also dependent on other cultivation conditions (i.e. temperature, salinity and incident light) (Hu et al., 2008), which makes the comparison of the data presented in Table 5 with values reported by others difficult. In addition, literature usually presents the total lipid content of microalgae instead of their fatty acid content. Values between 25% and 54% lipids (w/w) are common during nitrogen-depleted growth of N. oleoabundans in freshwater conditions, mostly at pH 7.5 and 25 °C (Tornabene et al., 1983; Gatenby et al., 2003; Li et al., 2008a,b; Gouveia et al., 2009; Pruvost et al., 2011). From this perspective, the fatty acid content of 35% (w/w) obtained under alkaline–saline conditions, in nitrogendepleted medium and during an experiment that was not yet fully optimized for lipid accumulation, justifies further studies on the kinetics of lipid accumulation in N. oleoabundans cultivated under extreme conditions. As described by Hu et al. (2008) the most abundant fatty acids under nitrogen depletion are the C16:0 and the C18:1. This was also confirmed in this study since a two-fold increase of the relative content of C18:1 fatty acids was found under nitrogen deplete conditions for both pH values (Table 5). The relative content of unsaturated fatty acids, and especially C18:3, on the other hand, decreased substantially. The effect of nitrogen on the accumulation of lipids and their unsaturation has been reported previously (Tornabene et al., 1983). But, our results show that N. oleoabundans can also accumulate lipids under high pH and high salt concentrations which makes this microalga a very versatile species for large-scale production of lipid-rich biomass.
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Table 5 Fatty acid content (% w/w) of N. oleoabundans grown at pH 8.1 and pH 10.0 during turbidostat experiments, without nitrogen limitation (N+) and with nitrogen depletion (N-). The correspondent relative compositions (%rel of the total) are also shown. Content (% w/w) Fatty acid
pH 8.1 (N+)
%rel
C16:0 C16:l C18:0 C18:l C18:2 C18:3 Total
2.41 0.22 0.17 1.00 2.14 1.80 7.74
31.1 2.8 2.2 12.9 27.6 23.6
pH 8.1 (N ) 3.92 – 0.35 4.00 3.08 1.88 13.23
These findings were supported through the visualization of the neutral lipid bodies inside the cells, which was possible using fluorescence microscopy. After a fast diffusion of BODIPY 505/515, the intracellular lipid bodies emitted a strong green fluorescence and could be easily distinguished from the chloroplasts. The chloroplasts appeared red due to chlorophyll fluorescence under the same excitation wavelength. During the course of both experiments the cells showed green fluorescent spherical bodies, either with or without nitrogen in the medium. However, that accumulation was more evident upon nitrogen starvation, with the majority of the cells showing a sharp and strong green fluorescence. Fig. 6 (Supplementry figure) shows the accumulation of those bodies in microalgae cells growing under pH 10.0. 4. Conclusions The growth of the ‘freshwater’ microalga N. oleoabundans was investigated under alkaline–saline conditions. A stable cultivation medium was developed under these conditions based on the minimal growth requirements and maximal solubility of nutritious salts. The growth rate obtained in this modified medium with 420 mM of salts at pH 10.0 brings new possibilities for cultivating N. oleoabundans and for the possible improvement of CO2 transfer in photobioreactors. Its potential as an alternative source for biofuels production is also shown, as lipids rapidly accumulate under those conditions. Lipid accumulation even seemed more pronounced when nitrogen starvation was applied at high pH. Acknowledgements This work was performed in the TTIW-cooperation framework of Wetsus, Center of Excellence for Sustainable Water Technology (http://www.wetsus.nl). Wetsus is funded by the Dutch Ministry of Economic Affairs, the European Union Regional Development Fund, the Province of Fryslan, the City of Leeuwarden and the EZ/ Kompas program of the ‘‘Samenwerkingsverband Noord-Nederland’’. The authors like to thank the participants of the research theme ‘‘Algae’’ for the discussions and their financial support. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.biortech.2011.10.084. References Brown, L.M., 1996. Uptake of carbon dioxide from flue gas by microalgae. Energy Convers. Manage. 37, 1363–1367. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306. Cooper, M.S., Hardin, W.R., Petersen, T.W., Cattolico, R.A., 2010. Visualizing ‘‘green oil’’ in live algal cells. J. Biosci. Bioeng. 109, 198–201.
%rel
pH 10.0 (N+)
29.6 – 2.6 30.2 23.3 14.2
2.74 0.31 0.39 3.34 3.23 1.51 11.52
%rel
pH 10.0 (N )
%rel
23.8 2.7 3.4 29.0 28.0 13.1
7.79 0.14 0.76 15.22 9.25 1.72 34.88
22.3 0.4 2.2 43.6 26.5 4.9
Gardner, R., Peters, P., Peyton, B., Cooksey, K.E., 2010. Medium pH and nitrate concentration effects on accumulation of triacylglycerol in two members of the chlorophyta. J. Appl. Phycol. doi:10.1007/s10811-010-9633-4. Gatenby, C.M., Orcutt, D.M., Kreeger, D.A., Parker, B.C., Jones, V.A., Neves, R.J., 2003. Biochemical composition of three algal species proposed as food for captive freshwater mussels. J. Appl. Phycol. 15, 1–11. Gouveia, L., Oliveira, A.C., 2008. Microalgae as a raw material for biofuels production. J. Ind. Microbiol. Biotechnol. 36, 269–274. Gouveia, L., Marques, A.E., da Silva, T.L., Reis, A., 2009. Neochloris oleoabundans UTEX #1185: a suitable renewable source for biofuel production. J. Ind. Microbiol. Biotechnol. 36, 821–826. Guckert, J.B., Cooksey, K.E., 1990. Triglyceride accumulation and fatty acid profile changes in Chlorella (Chlorophyta) during high pH-induced cell cycle inhibition. J. Phycol. 26, 72–79. Gustafsson, J.P., 2009. Visual MINTEQ (Version 2.61) [Software]. Available from. Ho, T.-Y., Quigg, A., Finkel, Z.V., Milligan, A.J., Wyman, K., Falkowski, P.G., Morel, F.M.M., 2003. The elemental composition of some marine phytoplankton. J. Phycol. 39, 1145–1159. Hu, Q., Sommerfeld, M., Jarvis, E., Ghirardi, M.L., Posewitz, M., Seibert, M., Darzins, A., 2008. Microalgal tiyacylglycerols as feedstocks for biofuels production: perspectives and advances. Plant J. 54, 621–639. Lamers, P.P., van de Laak, C.C.W., Kaasenbrood, P.S., Lorier, J., Janssen, M., de Vos, R.C.H., Bino, R.J., Wijffels, R.H., 2010. Carotenoid and fatty acid metabolism in light-stressed Dunaliella salina. Biotechnol. Bioeng. 106 (4), 638–648. Li, Y., Horsman, M., Wu, N., Lan, C.Q., Dubois-Calero, N., 2008a. Biofuels from microalgae. Biotechnol. Prog. 24, 815–820. Li, Y., Horsman, M., Wang, B., Wu, N., Lan, C.Q., 2008b. Effects of nitrogen sources on cell growth and lipid accumulation of green alga Neochloris oleoabundans. Appl. Microbiol. Biotechnol. 81, 629–636. Mata, T.M., Martins, A.A., Caetano, N.S., 2010. Microalgae for biodiesel production and other applications: a review. Renew. Sustain. Energy Rev. 14, 217–232. Norsker, N.-H., Barbosa, M.J., Vermue, M.H., Wijffels, R.H., 2011. Microalgal production – a close look at the economics. Biotechnol. Adv. 29, 24–27. Pruvost, J., Van Vooren, G., Cogne, G., Legrand, J., 2009. Investigation of biomass and lipids production with Neochloris oleoabundans in photobioreactor. Bioresour. Technol. 100, 5988–5995. Pruvost, J., Van Vooren, G., Le Gouic, B., Couzinet-Mossion, A., Legrand, J., 2011. Systematic investigation of biomass and lipid productivity by microalgae in photobioreactors for biodiesel application. Bioresour. Technol. 102, 150–158. Rodolfi, L., Zittelli, G.C., Bassi, N., Padovani, G., Biondi, N., Bonini, G., Tredici, M.R., 2009. Microalgae for oil: strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol. Bioeng. 102, 100–112. Roessler, P.G., 1990. Environmental control of glycerolipid metabolism in microalgae: commercial implications and future research directions. J. Phycol. 26, 393–399. Tornabene, T.G., Holzer, G., Lien, S., Burris, N., 1983. Lipid composition of the nitrogen starved green alga Neochloris oleoabundans. Enzyme Microb. Technol. 5, 435–440. Walker, G.M., 1994. The roles of magnesium in biotechnology. CRC critical reviews in biotechnology 14 (4), 311–354. Williams, P.J. le B., Laurens, L.M.L., 2010. Microalgae as biodiesel & biomass feedstocks: review & analysis of the biochemistry, energetics & economics. Energy Environ. Sci. 3, 554–590. Wijffels, R.H., Barbosa, M.J., 2010. An outlook on microalgal biofuels. Science 329, 796–799. Yagi, H., Yoshida, F., 1977. Desorption of carbon dioxide from fermentation broth. Biotechnol. Bioeng. 19, 801–819. Yeh, K.-L., Chang, J.-S., 2011. Nitrogen starvation strategies and photobioreactor design for enhancing lipid production of a newly isolated microalga Chlorella vulgaris ESP-31: implications for biofuels. Biotechnol. J. doi:10.1002/ biot.201000433. Zeiler, K.G., Heacox, D.A., Toon, S.T., Kadam, K.L., Brown, L.M., 1995. The use of microalgae for assimilation and utilization of carbon dioxide from fossil fuelfired power plant flue gas. Energy Convers. Manage. 36, 707–712. Zhu, C.J., Lee, Y.K., 1997. Determination of dry weight of marine microalgae. J. Appl. Phycol. 9, 189–194.
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Growth of oil accumulating microalga Neochloris oleoabundans under alkaline–saline conditions A.M. Santos a,b,⇑, M. Janssen b, P.P. Lamers b, W.A.C. Evers b, R.H. Wijffels b a b
Wetsus – Center of Excellence for Sustainable Water Technology, P.O. Box 1113, 8900 CC Leeuwarden, The Netherlands Bioprocess Engineering, Wageningen University and Research Center, P.O. Box 8129, 6700 EV Wageningen, The Netherlands
a r t i c l e
i n f o
Article history: Received 28 July 2011 Received in revised form 20 October 2011 Accepted 22 October 2011 Available online 6 November 2011 Keywords: Neochloris oleoabundans Alkalinity Growth rate Lipid accumulation Nitrogen starvation
a b s t r a c t The effect of elevated pH and salt concentration on the growth of the freshwater microalga Neochloris oleoabundans was investigated. A study was conducted in 24-well plates on the design of a growth medium and subsequently applied in a photobioreactor. An artificial seawater medium with reduced Ca2+ and PO34 could prevent mineral precipitation at high pH levels. Growth was characterized in this new medium at pH 8.1 and at pH 10.0, with 420 mM of total salts. Specific growth rates of 0.08 h 1 at pH 8.1 and 0.04 h 1 at pH 10.0 were obtained under controlled turbidostat cultivation. The effect of nitrogen starvation on lipid accumulation was also investigated. Fatty acids content increased not only with nitrogen limitation but also with a pH increase (up to 35% in the dry biomass). Fluorescence microscopy gave visual proof that N. oleoabundans accumulates oil bodies when growing in saline conditions at high pH. Ó 2011 Elsevier Ltd. All rights reserved.
1. Introduction The use of microalgal lipids as a feedstock for a sustainable production of biofuels is a hot topic within the scientific community. This is related to the high lipid content of some species and to the fact that lipid synthesis can be easily modulated by manipulating their cultivation conditions. Moreover, due to their high photosynthetic efficiency and faster growth rates in comparison to higher plants, lipid-rich microalgae can be considered as potentially one of the most efficient biological oil producers (Chisti, 2007; Li et al., 2008a,b; Mata et al., 2010; Wijffels and Barbosa, 2010). The total composition of lipids in microalgae can vary between 1% and 85% of their dry weight (Chisti, 2007) achieving values up to 40–70% under nutrient limitation (Roessler, 1990; Rodolfi et al., 2009). Nevertheless, a microalgae-based fuel production technology which can economically compete with the ones currently on the market still needs to be developed. The main limitations still associated with this technology are the selection of strains which can grow and produce high quantities of lipids at the same time, and the high energy inputs necessary for pumping, mixing and harvesting the produced biomass (Hu et al., 2008; Rodolfi et al., 2009; Norsker et al., 2011).
⇑ Corresponding author at: Bioprocess Engineering, Wageningen University and Research Center, P.O. Box 8129, 6700 EV Wageningen, The Netherlands. Tel.: +31 (0) 317 48 50 12; fax: +31 (0) 317 48 22 37. E-mail address: [email protected] (A.M. Santos). 0960-8524/$ - see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.biortech.2011.10.084
Substantial energy is required for the supply of carbon dioxide (CO2) to large-scale cultivation systems by means of gas transfer. During microalgal cultivation, CO2 is usually introduced to the system by CO2 enrichment of the inlet air flow, but it is envisioned to use CO2-rich gasses as for example flue gas (Zeiler et al., 1995; Brown, 1996). However, a considerable amount of CO2 which fails to be transferred into the liquid phase will still be wasted to the effluent gas. When growing microalgae at a pH higher than 8.5, the cultivation medium could be saturated with inorganic carbon as bicarbonate (HCO3 ) prior to entering the cultivation system. The energy necessary for CO2 transfer from the gas to the liquid phase could thus be reduced by optimizing CO2 transfer in a separate gas transfer unit at elevated pH (>8.5). Moreover, also in traditional systems where CO2 is supplied in the actual growth stage the efficiency of CO2 transfer could be increased when operating the system at a pH of 10 or more (Yagi and Yoshida, 1977). Such process strategies are only possible if there are microalgae able to efficiently grow and produce lipids under alkaline conditions at high salt concentrations (i.e. bicarbonate, carbonate, and associated counter-ions). The alkalinity of an aqueous solution is regulated by the distribution of the bicarbonate and carbonate species (HCO3 and CO23 ). These salts also determine the system’s buffering capacity. For this reason, they must be present at high enough levels to supply the necessary CO2 for microalgal growth and to keep the pH from increasing too much. Algal growth in culture also depends on the adequate supply of essential macro and micronutrients. These nutrients are usually
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highly soluble salts which dissociate in ions and are either present in high concentrations, as in the case of magnesium (Mg2+) and calcium (Ca2+) in seawater, or can be added in amounts sufficient to support good algal growth. At high pH (pH > 8.0) especially divalent ions have the tendency to precipitate with phosphate or hydroxide as insoluble salts. For this reason, the concentrations of ions such as Ca2+, Mg2+ and phosphate (PO34 ) should be kept as low as possible without limiting microalgae growth. The accumulation of large quantities of lipids by microalgal species has already been reported (Williams and Laurens, 2010). Among these species is Neochloris oleoabundans, known as a freshwater species capable of producing oil bodies composed of triacylglycerides (TAG) at a level of up to 80% of its total lipid composition. Most of those lipids are composed of saturated fatty acids (Tornabene et al., 1983), suitable for the production of biofuels. A more recent screening study performed by Gouveia and Oliveira (2008) also showed the potential of N. oleoabundans as a source of lipids for biofuels production. A considerable increase in oil quantity (50% of the dry weight) was claimed during growth under N-depletion. However, the possible application of N. oleoabundans for biofuels production under alkaline and saline conditions has not been covered in literature so far. The present work aimed to test whether N. oleoabundans could also be grown under alkaline and saline conditions while still being able to accumulate lipids. Experiments were performed in microtiter plates and in a lab-scale photobioreactor to determine the specific growth rate of this microalga under a range of alkalinities and salinities. Besides nutrient-replete growth, nitrogen deprivation was applied to induce lipid accumulation and to check whether this accumulation was affected by the alkalinity and the salinity of the growth medium. Moreover, the development of a stable inorganic growth medium for alkaline–saline conditions is discussed. 2. Methods 2.1. Microalgal strain N. Oleoabundans UTEX 1185 was pre-cultivated in 250 mL shake flasks containing 100 mL of medium. The cultures were placed on an orbital shaker incubator (Innova 44, New Brunswick Scientific, New Jersey, USA) under a 2% CO2-enriched headspace and a light intensity of 30 lmol photons m 2 s 1 provided by fluorescent lamps. Temperature was controlled at 25 °C and pH was 7.8. 2.2. Medium design A seawater-type medium (Table 1) was developed based on the elemental biomass composition of several marine species (Ho et al., 2003) and designed to support at least 1 g L 1 of biomass. Non-biological precipitation experiments were performed in 24well microtiter plates (Falcon 3047, BD Biosciences, CA, USA). The plates were fixed inside the incubator and rotated at 150 rpm (orbit 2.54 cm). In a total volume of 1.5 mL per well, different Mg2+/ Ca2+ combinations in the medium were tested. The ratio between both elements was either kept constant (13:1), or varied with a constant minimal Ca2+ concentration (0.01 mM). Nitrate was kept in a ratio of 14:1 with phosphate, which was added as a potassium polyphosphate solution insensitive to high pH levels (Super FK, YARA, Vlaardingen, The Netherlands). Six different pH values were tested in duplicate for each medium composition: 9.0, 9.5, 10.0, 10.3, 10.5 and 10.8. The desired pH was obtained by buffering with different 300 mM NaHCO3–Na2CO3 mixtures, as described in Table 2. The stability of the medium was assessed by following the optical density at 750 nm (OD750) using
Table 1 Modified medium composition (in 1 L). X and Y are the concentrations of magnesium and calcium in artificial seawater, respectively. Values for these concentrations were determined in this study and are given in Section 3.1. Compound
[mM]
Macronutrients KNO3 Super FK EDTA ferric sodium salt
4.4 0.32 29.28 lM
Vitamins Thiamine Biotin B12
200 lg L 1 lg L 1 1 lg L 1
Micronutrients EDTA-MnNa2 EDTA-ZnNa2 EDTA-CuNa2 Na2MoO42H2O CoCl26H2O
1.5 2.0 7.0 7.7 3.7
Artificial seawater NaCl MgCl26H2O CaCl22H2O Na2SO4
420.0 X Y 22.53
1
E-02 E-03 E-03 E-04 E-04
a multilabel plate reader (VICTORTM X3, PerkinElmer, MA, USA) and wells with demineralised water as blanks. An increase in optical density signifies salt crystals formation due to the light scattering by these insoluble particles. The growth of N. oleoabundans on the modified medium was studied under the same conditions, except for the additional supply of light (fluorescent light tubes, 30 lmol photons m 2 s 1) and CO2 (4% in headspace). Temperature was controlled at 30 °C. The wells (total volume 1.5 mL) were inoculated with 75 lL from 2 weeks old shake flask cultures. The optical density at 750 nm was measured daily to follow biomass growth (medium served as control).
2.3. Growth assessment on alkalinity and salinity The influence of a range of alkalinities and salinities on growth was also studied in 24-well microtiter plates. The wells (total volume 1.5 mL) were inoculated with 150 lL from 2 weeks old shake flask cultures and incubated as described in Section 2.2, with the exception that the light intensity was set at 55 lmol photons m 2 s 1. Microalgal growth was assessed at four pH values: 8.1, 9.0, 10.0 and 10.5, which were set by applying different NaHCO3–Na2CO3 buffer mixtures (Table 3) and by adjusting the CO2 fraction in the headspace. In four sets of experiments, different concentrations of NaCl in the medium were combined with the NaHCO3–Na2CO3 mixtures to check the algae’s salt tolerance Table 2 Molar concentrations of NaHCO3 and Na2CO3 in the buffering mixtures used during the medium design experiments. For each pH tested, the combined concentration of both salts was 300 mM. pH
NaHCO3 [mM|
Na2CO3 [mM]
9.0 9.5 10.0 10.3 10.5 10.8
270 210 150 90 60 30
30 90 150 210 240 270
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Table 3 Molar concentrations of NaHCO3 and Na2CO3 in the buffering mixtures used during growth assessment experiments. For each pH tested, the combined concentration of both salts in the medium was either 300 mM or 150 mM. pH
Total 300 [mM]
9.0 10.0 10.5
Total 150 [mM]
NaHCO3 [mM]
Na2CO3 [mM]
NaHCO3 [mM]
Na2CO3 [mM]
270 150 60
30 150 240
135 75 30
15 75 120
(Table 4). At pH 8.1 medium with solely 12 mM NaHCO3 was used. All sets of experiments were performed in four replicates. Repeated batch cultivation was applied to assess the specific growth rates for each medium. Every 24 h the cultures were diluted to the starting optical density (as represented in Fig. 1), whilst keeping the culture volume at 1.5 mL. The dilution series were continued until a constant specific growth rate was observed. 2.4. Growth assessment in a bench-scale panel photobioreactor The microalgae were grown in a flat panel photobioreactor (FMT150, Photon Systems Instruments, Brno, Czech Republic) with a detachable rectangular cuvette 20 cm high, 10 cm wide and 2.5 cm optical path (working volume 385 mL). The photobioreactor (PBR, Fig. 2) was equipped with temperature control (set to 30 °C) and a light source composed of red light emitting diodes, LEDs (kmax 627 nm). The incident light flux was measured at 15 different places equally distributed over the PBR’s light-exposed surface with a LI190 2p PAR quantum sensor (Li-CORÒ, Lincoln, USA). The average light intensity at the surface was set to 300 lmol photons m 2 s 1. A built-in dual-wavelength densitometer (680 nm and 735 nm) allowed us to monitor the culture growth and the chlorophyll absorption using the PBR itself as cuvette. Mixing was performed by continuous sparging at 650 mL min 1 of a CO2-enriched air stream through a gas-distributor over the bottom of the panel. The gas was saturated with water before entering the vessel. The influence of the pH on growth was studied at pH 8.1 and pH 10.0. Growth was started batch wise using 30 mL inoculum from a 2 weeks old shake flask culture and 355 mL medium with optimal Ca2+ and Mg2+ salt concentrations (X and Y in Table 1, see Section 3). The concentration of NaCl in the medium was 420 mM at pH 8.1 and 270 mM at pH 10.0. At pH 8.1, 12 mM NaHCO3 was in equilibrium with 1% v/v CO2. At pH 10.0, a mixture of 75 mM NaHCO3 and 75 mM Na2CO3 was applied, while gassing with 0.1% v/v CO2. An OD735 of 0.2 was defined as the start and set-point for the turbidostat control. In turbidostat mode, the OD735 was maintained constant by the automatic addition of fresh medium and concomitant harvesting of surplus culture. The dilution rate (D), being equal to the specific growth rate of the microalgae, was assessed daily. The correspondent biomass concentrations (Cx) were determined by dry weight measurements. At both pH the algae were deprived of nitrogen, after a constant growth rate at nitrogen-replete conditions was obtained, to induce the accumulation of lipids. To this end, the feed medium was replaced with
Fig. 1. Schematic representation of dilution steps in 24-well microtiter plates.
nitrogen-free medium resulting in a gradual but complete depletion of nitrogen in the reactor. 2.5. Biomass dry weight determination The biomass concentration was determined according to Zhu and Lee (1997). Triplicate samples (10 mL) were diluted in ammonium formate (0.5 M) and filtrated with a constant vacuum (0.44 bar, absolute) over pre-dried and pre-weighted Whatman GF/F glass fiber filters (Ø 55 mm, pore size 0.7 lm). The filters were rinsed twice with ammonium formate, dried (24 h, 95 °C), cooled down in a dessicator (2 h) and weighted again. The biomass dry weight was determined as the difference between the weights of the dried filters containing the samples and the dried empty filters. 2.6. Visualization of intracellular neutral lipid bodies The accumulation of intracellular lipid bodies was checked by fluorescence microscopy. The lipophilic fluorescent dye BODIPY 505/515 (4,4-difluoro-1,3,5,7-tetramethyl-4-bora-3a,4a-diazasindacene; Invitrogen Molecular Probes, Carlsbad, CA) was used for staining the cells, according to Cooper et al. (2010). A 40 lM BODIPY 505/515 stock solution was prepared by dissolving the dye in 0.2% (v/v) of anhydrous dimethyl sulfoxide (DMSO). Aliquots of this solution were added directly to the suspensions, having a final dye concentration of 1 lg mL 1 and 0.02% DMSO. After 5 min of contact, slides were prepared for observation under an inverted fluorescence microscope (Leica DMI6000, Leica Microsystems B.V., Rijswijk, The Netherlands). Excitation was done under a blue excitation light through a band-pass filter (450–490 nm) and emission wavelengths were imaged through a long-pass filter (515 nm). 2.7. Fatty acids quantification
Table 4 Sodium chloride combinations with the NaHCO3 and Na2CO3 buffering mixtures tested during growth assessment experiments at different pH values. Set
NaCl [mM]
NaHCO3 + Na2CO3 [mM]
1 2 3 4
420 – 120 270
300 300 300 150
Duplicate reactor samples were centrifuged for 5 min at 2500 rpm and 15 °C and the cell pellets were stored at 80 °C. On a later stage, the pellets were transferred to bead beating tubes (Lysing Matrix E, MP Biomedicals) and lyophilized overnight followed by gravimetric biomass dry weight determination. Upon addition of a chloroform/methanol mixture (2:2.5), containing 47 lg mL 1 of the internal standard tripentadecanoin (Sigma– Aldrich T4257), complete cell disruption was achieved by bead
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Fig. 2. Schematic representation of photobioreactor FMT150. The green parts correspond to the additional connections when operating in turbidostat mode.
beating for 40 min. The homogenates were then transferred to 12 mL glass tubes. Complete recovery of the homogenate was assured by rinsing the bead beating tubes with an additional 3 mL of the chloroform/methanol mixture. The next steps of the extraction were adapted from a protocol optimized by Lamers et al. (2010) and are available as supplemental material. Gas chromatography (GC) analysis was done using a HP 6890 (Hewlett Packard Inc.) with FID detector. A volume of 1 lL was injected (split ratio 0.1:1) onto a Supelco NucolTM column (355 33-03A, 30 m 530 lm 1.0 lm film thickness) with helium as the carrier gas (constant total column flow 20 mLmin 1). The column oven was programmed at an initial temperature of 90 °C for 0.5 min, followed by a ramp of 20 °Cmin 1 to 200 °C and a final step of 24 min at 200 °C. The FAMEs were identified by comparison of their retention times with those of authentic standards (Sigma Aldrich, MO, USA). Quantification of the identified lipids was done by correlating their individual peak areas to the one of the internal standard, taking into account the amount of internal standard added per sample and the response factors that were calibrated for each fatty acid using authentic standards. Samples that did not contain biomass, but were otherwise subjected to the entire protocol, served to correct for three minor peaks that were extracted from the plastic bead beater tubes.
3. Results and discussion 3.1. Growth of N. oleoabundans in a modified medium The preparation of cultivation media with high concentrations of salts and high pH needs careful attention regarding the issue of precipitation. Compounds which precipitate are no longer available for microalgal growth. For instance, the binding capacity of calcium to phosphate is pH dependent, increasing at elevated values. A chemical equilibrium model for the calculation of saturation indices in aqueous solutions (Visual MINTEQ, Gustafsson, 2009) was the base for developing a medium with low levels of precipitation. A simulation showed for example high formation of a calcium phosphate complex, hydroxyapatite, for our modified medium at pH 10.5 when containing Ca2+ and PO34 in concentrations of 3.61 mM and 5 mM, respectively. One way to reduce the formation of hydroxyapatite is to decrease the concentrations of Ca2+ and/or PO34 . It is difficult to get reliable data on the need of calcium for microalgae growth. Our non-biological experiments showed that using Ca2+ in a concentration of 0.01 mM results in a medium with very low complexation levels at high pH values. Besides calcium, also
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magnesium can complex and precipitate at high pH, though at a lesser degree. Furthermore, the amount of magnesium needed by the cells seems to be higher than calcium. Magnesium is essential for cell growth, while the need for calcium can be questionable (Walker, 1994). In our experiments, a ratio of 1:15 between Ca2+ and Mg2+ could be translated in minimal precipitation, magnesium being in a concentration of 0.15 mM. Based on these findings we developed a new medium with 0.01 mM CaCl2 (X in Table 1), 0.15 mM MgCl2 (Y in Table 1), 4.4 mM KNO3 and 0.32 mM ‘P’ from the polyphosphate solution (Super FK). Additional experiments indeed confirmed that this medium resulted in negligible precipitation. In Fig. 3A the optical density shows the level of precipitation at different pH values. In the first 100 h the OD hardly increases. Only at elevated pH (>10) and after extended time (150 h) limited precipitation occurs. For continuous microalgae production this should not cause any problems since the rate of consumption will be faster than the rate of precipitation. Moreover, continuous consumption will result in even lower concentrations of Ca2+, Mg2+ and PO34 , further reducing the risk of precipitation. Although in the new medium there was sufficient nitrogen and phosphate available to sustain a biomass concentration of 1 g L 1 it was not known whether the final concentrations of calcium and magnesium were sufficient to support algae growth up that value. However, during experiments with N. oleoabundans growing batch wise in this modified medium in the FMT150 photobioreactor (work not presented) a final biomass concentration of 1.8 g L 1 was obtained, confirming the calculation that at least 1 g L 1 can be sustained by this cultivation medium. Fig. 3B shows the increase in optical density during growth in the modified medium at different pH values. As expected, growth of N. oleoabundans was not limited by the low concentrations of Ca2+ and Mg2+, although there was a general decrease with the increase of pH, especially at values above 10.0. 3.2. Influence of pH and salt concentration on specific growth rate 3.2.1. Microalgae growth in 24-well microtiter plates The results presented in the previous section show that growing N. oleoabundans in a saline medium with a high pH is possible. In this section it is discussed how the combination of pH and salt concentration affects the growth of this microalga. For all the pH values tested (Table 3), no growth of N. oleoabundans was observed with the highest total salt concentration of
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720 mM (Set 1, Table 4). Furthermore, very poor growth was observed at pH 10.5, for all sets of experiments. Growth rates were not determined for this value. At higher pH levels with less HCO3 /CO23 (total 150 mM), the combination with more NaCl (270 mM/Set 4) seems to be better for the algae than with less NaCl (absent/Set 2 and 120 mM/Set 3) combined with more HCO3 /CO23 (total 300 mM). A medium with 270 mM NaCl and 150 mM HCO3 / CO23 was therefore the only formulation capable of supporting growth at pH 10.0. An average growth rate of 0.0212 h 1 was found for the reference experiment at pH 8.1 (Fig. 4A), being the highest amongst all the experiments. However, this value was not considerably different for the one found at pH 10.0, where the specific growth rate reached 0.0177 h 1 (Fig. 4B). A correlation between the pH and the salt concentration seems evident as both have an impact on microalgal growth. Although the specific growth rates are all relatively low, due to light limitation inside the incubators (55 lmol photons m 2 s 1), they give a good insight on which should be the best conditions to grow N. oleoabundans at high pH: a marine medium with 270 mM NaCl, 75 mM NaHCO2 and 75 mM Na2CO3. 3.2.2. Microalgae growth in turbidostat cultivation and lipid accumulation under nitrogen depletion To estimate the growth under high pH and salt concentration under controlled conditions in a continuous system, two turbidostat experiments were performed at pH 8.1 and pH 10.0. The light intensity applied was of saturating nature (300 lmol photons m 2 s 1) and allowed to assess the maximum specific growth rate without any light limitation. The total salt concentration was 420 mM according to the study performed in the microtiter plates. The respective biomass concentrations (Cx) and dilution rates (D) were determined under steady-state. In steady state the specific rate of cell growth equals the rate of dilution. Growth rates of 0.08 h 1 and 0.04 h 1 were found at pH 8.1 and pH 10.0 (Fig. 5A and B, respectively). The biomass concentration Cx was constant under steadystate at 0.1 g L 1 in both experiments. Although the growth rate we obtained at pH 10.0 was twice as small as that at pH 8.1, it is still comparable with the one found by Pruvost et al. (2009). In their work, they reached a biomass concentration of around 0.1 g L 1 at a specific growth rate of 0.05 h 1 under the same light intensity, but in freshwater medium (bold basal medium) at pH 7.5 and 25 °C. The influence of nitrogen starvation on the accumulation of lipids was also investigated. Nitrogen starvation is known to be a
Fig. 3. Optical density at 750 nm (OD750) of (A) modified medium at different pH values where the increase in OD750 is a measure of medium precipitation and (B) N. oleoabundans in the modified medium at different pH values as a measure of growth. Lowest and highest pH values tested are represented by white and black triangles, respectively. Error bars represent the standard deviation from the average of the replicates.
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Fig. 4. Growth of N. oleoabundans in dilution series at (A) pH 8.1 and (B) pH 10.0. Growth rates determined 24 h after each dilution step.
Fig. 5. Growth of N. oleoabundans in turbidostat cultivation at (A) pH 8.1 and (B) pH 10.0. The vertical dotted lines represent the point after what depletion of nitrogen from the medium was applied.
strong inducer of lipid accumulation (Tornabene et al., 1983; Li et al., 2008a,b; Pruvost et al., 2011; Yeh and Chang, 2011). After reaching a constant growth rate, the cultivation medium was replaced by nitrogen-free medium. This resulted in a sharp decrease in l, to 0.02 h 1 at pH 8.1 and 0.01 h 1 at pH 10.0, and this decrease is obviously caused by the absence of nitrogen.
Cells growing in the photobioreactor at pH 8.1 and 10.0 were also analyzed for their total fatty acid content and composition, both under nitrogen-replete conditions and after 1 day of nitrogen depletion. Table 5 shows the microalgal content in each of the identified fatty acid, as well as their relative percentage of the total fatty acid pool. As we expected, there was an increase in the total fatty acid content of the nitrogen-starved cells one day after switching from N-replete medium to N-free medium. This increase was even more pronounced for the cells growing at pH 10.0 (up to 35% of dry weight), which may suggest that an additional stress such as high pH could promote lipid accumulation. A similar conclusion was taken by Guckert and Cooksey (1990) when they studied the accumulation of TAG during the growth of Chlorella at high pH. They suggested that such high pH-related stress can even be nitrogen independent, resulting in cell-cycle inhibition and triggering TAG accumulation. This was recently supported by Gardner et al. (2010), who suggested that high pH and nitrogen depletion have a synergistic effect on TAG accumulation in microalgae. But, the fatty acid contents of microalgae are also dependent on other cultivation conditions (i.e. temperature, salinity and incident light) (Hu et al., 2008), which makes the comparison of the data presented in Table 5 with values reported by others difficult. In addition, literature usually presents the total lipid content of microalgae instead of their fatty acid content. Values between 25% and 54% lipids (w/w) are common during nitrogen-depleted growth of N. oleoabundans in freshwater conditions, mostly at pH 7.5 and 25 °C (Tornabene et al., 1983; Gatenby et al., 2003; Li et al., 2008a,b; Gouveia et al., 2009; Pruvost et al., 2011). From this perspective, the fatty acid content of 35% (w/w) obtained under alkaline–saline conditions, in nitrogendepleted medium and during an experiment that was not yet fully optimized for lipid accumulation, justifies further studies on the kinetics of lipid accumulation in N. oleoabundans cultivated under extreme conditions. As described by Hu et al. (2008) the most abundant fatty acids under nitrogen depletion are the C16:0 and the C18:1. This was also confirmed in this study since a two-fold increase of the relative content of C18:1 fatty acids was found under nitrogen deplete conditions for both pH values (Table 5). The relative content of unsaturated fatty acids, and especially C18:3, on the other hand, decreased substantially. The effect of nitrogen on the accumulation of lipids and their unsaturation has been reported previously (Tornabene et al., 1983). But, our results show that N. oleoabundans can also accumulate lipids under high pH and high salt concentrations which makes this microalga a very versatile species for large-scale production of lipid-rich biomass.
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Table 5 Fatty acid content (% w/w) of N. oleoabundans grown at pH 8.1 and pH 10.0 during turbidostat experiments, without nitrogen limitation (N+) and with nitrogen depletion (N-). The correspondent relative compositions (%rel of the total) are also shown. Content (% w/w) Fatty acid
pH 8.1 (N+)
%rel
C16:0 C16:l C18:0 C18:l C18:2 C18:3 Total
2.41 0.22 0.17 1.00 2.14 1.80 7.74
31.1 2.8 2.2 12.9 27.6 23.6
pH 8.1 (N ) 3.92 – 0.35 4.00 3.08 1.88 13.23
These findings were supported through the visualization of the neutral lipid bodies inside the cells, which was possible using fluorescence microscopy. After a fast diffusion of BODIPY 505/515, the intracellular lipid bodies emitted a strong green fluorescence and could be easily distinguished from the chloroplasts. The chloroplasts appeared red due to chlorophyll fluorescence under the same excitation wavelength. During the course of both experiments the cells showed green fluorescent spherical bodies, either with or without nitrogen in the medium. However, that accumulation was more evident upon nitrogen starvation, with the majority of the cells showing a sharp and strong green fluorescence. Fig. 6 (Supplementry figure) shows the accumulation of those bodies in microalgae cells growing under pH 10.0. 4. Conclusions The growth of the ‘freshwater’ microalga N. oleoabundans was investigated under alkaline–saline conditions. A stable cultivation medium was developed under these conditions based on the minimal growth requirements and maximal solubility of nutritious salts. The growth rate obtained in this modified medium with 420 mM of salts at pH 10.0 brings new possibilities for cultivating N. oleoabundans and for the possible improvement of CO2 transfer in photobioreactors. Its potential as an alternative source for biofuels production is also shown, as lipids rapidly accumulate under those conditions. Lipid accumulation even seemed more pronounced when nitrogen starvation was applied at high pH. Acknowledgements This work was performed in the TTIW-cooperation framework of Wetsus, Center of Excellence for Sustainable Water Technology (http://www.wetsus.nl). Wetsus is funded by the Dutch Ministry of Economic Affairs, the European Union Regional Development Fund, the Province of Fryslan, the City of Leeuwarden and the EZ/ Kompas program of the ‘‘Samenwerkingsverband Noord-Nederland’’. The authors like to thank the participants of the research theme ‘‘Algae’’ for the discussions and their financial support. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.biortech.2011.10.084. References Brown, L.M., 1996. Uptake of carbon dioxide from flue gas by microalgae. Energy Convers. Manage. 37, 1363–1367. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306. Cooper, M.S., Hardin, W.R., Petersen, T.W., Cattolico, R.A., 2010. Visualizing ‘‘green oil’’ in live algal cells. J. Biosci. Bioeng. 109, 198–201.
%rel
pH 10.0 (N+)
29.6 – 2.6 30.2 23.3 14.2
2.74 0.31 0.39 3.34 3.23 1.51 11.52
%rel
pH 10.0 (N )
%rel
23.8 2.7 3.4 29.0 28.0 13.1
7.79 0.14 0.76 15.22 9.25 1.72 34.88
22.3 0.4 2.2 43.6 26.5 4.9
Gardner, R., Peters, P., Peyton, B., Cooksey, K.E., 2010. Medium pH and nitrate concentration effects on accumulation of triacylglycerol in two members of the chlorophyta. J. Appl. Phycol. doi:10.1007/s10811-010-9633-4. Gatenby, C.M., Orcutt, D.M., Kreeger, D.A., Parker, B.C., Jones, V.A., Neves, R.J., 2003. Biochemical composition of three algal species proposed as food for captive freshwater mussels. J. Appl. Phycol. 15, 1–11. Gouveia, L., Oliveira, A.C., 2008. Microalgae as a raw material for biofuels production. J. Ind. Microbiol. Biotechnol. 36, 269–274. Gouveia, L., Marques, A.E., da Silva, T.L., Reis, A., 2009. Neochloris oleoabundans UTEX #1185: a suitable renewable source for biofuel production. J. Ind. Microbiol. Biotechnol. 36, 821–826. Guckert, J.B., Cooksey, K.E., 1990. Triglyceride accumulation and fatty acid profile changes in Chlorella (Chlorophyta) during high pH-induced cell cycle inhibition. J. Phycol. 26, 72–79. Gustafsson, J.P., 2009. Visual MINTEQ (Version 2.61) [Software]. Available from