Effects of photoperiod, salinity and pH on cell growth and lipid content of Pavlova lutheri

Abstract

The aim of the present work was to study the effects of photoperiod, salinity and pH on growth and lipid content of Pavlova lutheri microalgae for biodiesel production in small-scale and large-scale open-pond tanks. In a 250-mL flask, the cultures grew well under 24 h illumination with maximum specific growth rate, μ _max , of 0.12 day⁻¹ and lipid content of 35 % as compared to 0.1 day⁻¹ and 15 % lipid content in the dark. The salinity was optimum for the cell growth at 30–35 ppt, but the lipid content of 34–36 % was higher at 35–40 ppt. Algal growth and lipid accumulation was optimum at pH 8–9. Large-scale cultivation in 5-L and 30-L tanks achieved μ _max of 0.13–0.14 day⁻¹ as compared to 0.12 day⁻¹ in small-scale and 300L cultures.

Introduction

Microalgae are unicellular photosynthetic organisms that use light energy and carbon dioxide, and show higher photosynthetic efficiency than plants for the production of biomass (Miao and Wu 2006). Lipids from microalgae can be turned into biodiesel through transesterification and used for energy development including simple combustion in a boiler or diesel engine (Vasudevan and Briggs 2008). Biodiesel produced from microalgae will not compromise the production of food and other products derived from crops. As an alternative to corn and sugar cane, many believe that algae are ideal to replace all other biofuel feedstocks as the cheapest, easiest, and most environmentally friendly way to produce liquid fuel (Antoni et al. 2007). They grow rapidly, accumulate large amounts of oil inside the cells leading to high biomass productivity, and do not need much land for cultivation as compared to oil crops and terraneous plants (Chisti 2007).

The entire production process ranging from the cultivation of high-lipid microalgae to the production of biodiesel from microalgal oil has been explored. A typical algal culture system can generate 150–400 barrels of oil per acre per year, which is 30 times more than those produced via typical oilseed crops. The production cost of algal oil depends on many factors, such as yield of biomass from the culture system, oil content, scale of production systems, and cost of recovering oil from algal biomass. Currently, algal-oil production is still far more expensive than petroleum diesel fuels. The production cost of algal oil from a photobioreactor with an annual production capacity of 1,000 tonnes per year, assuming the oil content of approximately 30 %, is estimated to be $2.80 per liter ($10.50 per gallon), while the petroleum-diesel price is at $3.80–4.50 per gallon (Chisti 2007). This does not include the cost of converting algal oil to biodiesel, distribution and marketing costs for biodiesel, and taxes.

Oil levels of 20–50 % are common in microalgae (Chisti 2007), though the ratios of lipid, carbohydrates and proteins are species-dependent. In some species, lipids can be up to 60 % of the algal dry weight (Griffiths and Harrison 2009). The quantity and quality of lipids within the cell may vary as a result of changes in growth conditions (temperature and light intensity) or nutrient media concentrations (nitrogen, phosphates and iron) (Liu et al. 2008). Factors such as temperature, irradiation, and nutrient availability are crucial to microalgal metabolism and high lipid productivity (Rodolfi et al. 2009; Mandal and Mallick 2009). There is a relationship between substrate inhibition and maintenance energy (Chen and Johns 1996). Algae have a trigger which, when put into stressful environments such as nutrient deprivation, may switch the use of carbon uptake from reproduction to energy storage.

Pavlova lutheri, a phytoflagellate, is a good example of high lipid-producing marine microalgae. It is a common member of the Pavlovophyceae (Haptophyta), often used as a food source for aquatic filter-feeders and cultured in laboratories to produce high levels of polyunsaturated fatty acids (PUFAs). A size of 4–6 μm, cell density of 0.4–2.5 g L⁻¹ and 15–35 % lipid content have been reported, with doubling time of 3–4 days (Guihéneuf et al. 2011; Carvalho et al. 2009). Manipulation of processing conditions (temperature, salinity, light, pH and nutrients), as well as culture duration allows modulation of cell growth and biochemical and lipid composition for consequent optimization of overall yield and productivity (Carvalho et al. 2009). With technological advances and more knowledge on algal biology, the commercialization of algal biofuel production will be feasible in the not too distant future once the issues related to production methods, i.e. growth, harvest of the algal culture, extraction of the lipid and conversion into diesel, are properly addressed.

The aim of this study was to evaluate the influence of environmental conditions such as photoperiod, salinity and pH on the growth and lipid content of P. lutheri. The cell concentration, dry weight and lipid content were analyzed. Comparison of the kinetics of cell growth between 250-mL and large-scale cultures in 1-, 5-, 30-, and 300-L batch cultures were made, to identify optimization parameters for scale-up.

Materials and methods

Algal culture and maintenance

Algal culture and maintenance were established as batch cultures. For small-scale cultivation (from 250 to 1 L), 10 % (v/v) of inocula (cell density of 12.0 × 10⁶cells mL⁻¹) was added into 100 mL of Conway media in 250-mL Erlenmeyer flasks. All media constituents were added aseptically after sterilization (MacLachlan 1973; Hamilton 1973). All chemicals and solvents were obtained from Merck (Darmstadt, Germany). Cultures were subcultured on a fortnightly basis, and grown on an orbital shaker at 80 rpm, at 28 ±2 °C, under 24 h illumination of 90–130 μmol photons m² s⁻¹ intensity from white fluorescent tube (Philips).

Effects of photoperiod, salinity and pH

Unless stated otherwise, the standard conditions for culture were 24 h illumination, 30 ppt NaCl and initial pH 7.5. The effects of photoperiod tested were 24 and 12 h illuminations and dark conditions. The effects of salinity were tested at 15, 20, 25, 30, 35 and 40 ppt of NaCl. The effects of pH studied were pH 5, 6, 7, 8, 9 and 10. Cultures were grown for 16 days, and removed in duplicate at 3- to 4-day intervals, and subjected to analysis of cell number, dry weight and lipid.

Large scale cultivation

Large-scale cultivation (from 5 to 300 L), as shown in Fig. 1, was carried out in a batch system using TMRL Enrichment media (AQUACOP 1984). Optimized culture conditions (obtained at the 250-mL scale at pH 8, salinity of 35 ppt, photoperiod of 24 h and illumination of 130 μmol photons m⁻² s⁻¹ intensity) were applied in large-scale cultivation, except that sunlight illumination was used for the 300-L cultivation. For different culture scales, 1-L Erlenmeyer flasks, 5-L plastic containers, and 30- and 300-L spherical tanks were used. Aeration were provided non-aseptically. Flasks were covered with rubber stoppers pierced with two holes for the air inlet and outlet.

Fig. 1

Cultivation of P. lutheri in a 5-L container, b 30-L tank, c 300-L tank

Plastic containers of 1 and 5 L were sterilized empty and later filled with microsterilized seawater. Nutrients were added at the time of inoculation and flasks were inoculated with 8- to 11-day-old cultures from 250-mL or 1-L flasks, respectively. Cultures were illuminated with white fluorescent tube (Philips) of 130 μmol photons m² s⁻¹ intensity.

Thirty liter tanks were disinfected with chlorox and filled with microfiltered seawater. Media enrichment was made at the time of inoculation and tanks were inoculated with 8- to 11-day-old stock cultures from 5-L containers. PVC distribution pipes were fixed with a channel to allow condensing water to be purged. The distribution of air into the culture tanks was made through 6-mm-diameter glass tubes. Cultures were illuminated with white fluorescent tube (Philips) of 130 μmol photons m² s⁻¹ intensity.

A cylindrical tank with conical bottom was used for 300-L open-pond cultivation. The tank was made from translucent 1-mm fiberglass sheet, and covered with a fiberglass cap pierced with one hole for the aeration inlet. The tanks were shaded with green sheets 4 m above to prevent overheating during long periods of strongest sunlight irradiation. Tanks were washed and filled with chlorinated hot water overnight, and thoroughly rinsed before use. The tanks were then filled with microfiltered seawater and disinfected by chlorination. Chlorox was used as the disinfectant at 1–2 ppm concentration and stirred for even distribution. The oxidizing agent is often applied to situations where autoclaving is not practical, as in the case of large-volume cultures. After 24 h, the chlorox was neutralized by the addition of sodium thiosulfate. Media enrichment was made at the time of inoculation and 300-L tanks were inoculated with 8- to 11-day-old stock cultures from 30-L tanks. Cultures were grown for 16 days and samples were taken from the top, bottom and middle of the tanks to ensure representative samples for analyses.

Determination of cell dry weight (dw)

Cells were harvested in triplicates by removing 10- or 100-mL samples and later centrifuged at 25,000 rpm (Avanti J-251 Centrifuge). Large-scale culture samples were flocculated by using an appropriate quantity of alum. Algal suspension was filtered through a pre-dried and pre-weighed glass-fiber filter (Whatman GF/C). The biomass was washed with de-mineralized water, dried at 80 °C in an oven overnight, cooled in a desiccator, and weighed.

Growth kinetics

Cell concentration (optical density) was determined at 620 nm with a spectrophotometer and nanodrop technique. The cell number and density were measured by a hemocytometer (Hirschmann, Germany).

Maximum specific growth rate (μ _max ) and doubling time (t _d ) were calculated based on the following equation :

$$ \frac{dX }{dt }=\mu X $$

(1)

where X is the cell dry weight (g L⁻¹) and t is the time (day).

Measurement of lipid content

Lipid measurements were calculated based on a modified method adapted from Bligh and Dyer (1959). This method extracts lipids from the algal cells by using a mixture of methanol, chloroform and water.

Fatty acids analysis

The extracted lipid was first transesterified into fatty acid methyl esters (FAME) (Mbatia et al. 2010), where 20 mg of the lipid sample was mixed with 2 mL of toluene, followed by addition (2 mL) of 1.5 % of sulphuric acid in dry methanol. After mixing well, the mixture was incubated at 55 °C overnight. Four mL of saturated NaCl solution was added, vortexed and 2 mL of hexane (HPLC grade) added, followed by 3 mL of sodium hydrogen carbonate (2 % NaHCO₃). The mixture was vortexed and 180 μL of the upper phase was taken for gas chromatography analysis.

FAME were separated and quantified using gas chromatography (Varian Chrompack CP 3800 GC) system equipped with a flame ionization detector (FID), and separation achieved by Supelcowax TM 10 fused silica capillary column (60 m × 0.32 mm × 0.25 μm film thickness; Supelco, Bellefonte, PA, USA). The carrier gas was helium at 550 kpa. The temperature programme was as follows: initial column oven temperature of 35 °C held for 3 min, and increased to 240 °C at 10 °C/min and held further for 35 min. The detector temperature was kept constant at 300 °C.

Results and discussion

Shake flask cultivation

Cell growth and lipid content

Figure 2 shows the effects of photoperiod, salinity and pH on cell density and dry weight of P. lutheri. The highest cell density of 13.3–14.1 × 10⁶ cells mL⁻¹ and dry weight of 0.45 g L⁻¹ was obtained under 24 h illumination, at 35 ppt NaCl and pH 8–9. As shown in Table 1, this corresponds to the doubling time of 5.33–5.77 days, μ_max of 0.12–0.13 day⁻¹ and lipid content of 35–36 %. These values were comparable to Spirulina platensis biomass reported at 0.82 g L⁻¹, μ_max at 0.1357 day⁻¹ and t _d at 5.11 days (Costa et al. 2003); and those reported for Scenedesmus dimorphus and Scenedesmus quadricauda at biomass of 0.141 g L⁻¹ and 0.358 L⁻¹, μ_max of 0.202 day⁻¹ and 0.516 day⁻¹; t _d of 4.91 and 1.93; and lipid content of 34 % and 31 %, respectively (Goswami and Kalita 2011). Several strains such as Nannochloropsis, Nannochloris, Pavlova lutheri and Phaeodactylum tricornutum have reported lipid contents similarly around 26–36 % (Griffiths and Harrison 2009).

Fig. 2

Effects of a photoperiod, b salinity, c pH, on cell density and dry weight of P. lutheri cultivated in a 250-mL flask

Table 1 Kinetics of cell growth and lipid production of P. lutheri cultivated in a 250-mL flask

Effects of light and photoperiod

The lowest cell density of 2.5 × 10⁶ cells mL⁻¹, dry weight of 0.12 g L⁻¹ and lipid of 15 % obtained in dark condition, suggest the importance of efficient light utilization. Previous studies on Rhodomonas sp., Cryptomonas sp., and Isochrysis sp. also indicate that lipid content increase with increasing photoperiod by 15.5, 12.7, and 21.7 %, respectively (Sharma et al. 2012). Light is the driving force of photosynthesis, as well as for cell photo-acclimatization. Physiological properties of phytoplankton and photosynthetic organisms can be changed upon exposure to photoperiod and light intensity. The effects of light and photoperiod may also actually influence the cultivation temperature. Increased microalgal lipid productions to 26–36 % at temperatures of 25–30 °C for several microalgal species have been reported. However, reduced lipid production to 15–20 % at extremes of low (15 °C) and high temperature (30 °C) has been observed in Isochrysis galbana and Nanochloropsis species (Sayegh and Montagnes 2011). Low temperatures reduce enzyme activity in glycolysis and the Krebs cycle and consequently the metabolism of carbon sources. Metabolic engineering of pathways could assist to evaluate the activities of enzymes in the metabolic network that can be upregulated or downregulated for enhanced lipid productivity.

Effects of salinity and pH

Optimum salinity for P. lutheri at 30–40 ppt and optimal pH 7.5–9 agrees well with that reported for Navicula acceptata, N. pelliculosa and N. saprophila (Griffiths and Harrison 2009), Chlorella vulgaris at pH 7–8 (Wang et al. 2010) and pH 8.2–8.7 for cyanobacteria (Kim et al. 2007). The salinity level for growth retardation and low lipid content was observed at 15–25 ppt NaCl and pH 5–7 and at pH 10. Culture pH and salinity influence influx and efflux of anions and cations into the cellular system. The effects of low salinity level on growth retardation has also been observed in N. gregari (Morin et al. 2008); and on lower lipid contents (18–19 % dw) in nutrient replete conditions in Chlamydomonas applanata and C. reinhardtii (Griffiths and Harrison 2009). In our study, P. lutheri has been isolated from marine environment that thrives at high salinity. Different geographical strains have different preferences towards salinity. Optimal salinity may be a function of immediate conditions from which the strain is initially isolated. Species isolated at higher salinities will grow better at higher salinities and may not be as good at lower salinities (Cucchiari et al. 2008). Higher lipid (60.6 % dw) has been reported for the genus Dunaliella in salt concentrations as high as 0.5 M NaCl (equivalent to seawater, or 35 psu) (Takagi et al. 2006) and increases as NaCl increases. In a study with S. obliquus, the highest lipid content of 50 % is reported after 10 days stress period at NaCl of 1.0 M (Dujjanutat and Kaewkannetra 2011).

Fatty acids composition

In a typical process of biofuels production, trans-esterification (alcoholysis) produces esters of fatty acids and glycerol. From Table 2, trans-esterification of lipid yielded high long carbon-chain fatty acids. The major components in P. lutheri were hexadecanoate C 16:0 (38.9 %), palmitoloate C 16:1 (26.4 %) and eicosapentaenoate C 20:5 (12.1 %). The total saturated fatty acids of 42 %, monounsaturated fatty acids of 38.51 % and PUFA of 19.49 % were obtained. These are comparable to the lipid classes of P. lutheri, cultivated in semicontinuous mode, where neutral lipids and glycolipids, as the major constituents, accounted for 57 and 24 % of the total fatty acids residues (TFA), respectively, with emphasis on eicosapentaenoic (C20:5n-3, EPA) and docosahexaenoic (C22:6n-3, DHA) acids (Meireles et al. 2003). Another study in which P. lutheri undergoes UV-R treatment has reported total saturated fatty acids of 33.8 %, monounsaturated fatty acids of 18.7 % and PUFA of 46.1 % (Guihéneuf et al. 2010).

Table 2 Fatty acids composition of lipid recovered from P. lutheri cultivated in a 250-mL flask

Makri et al. (2011) cultivate Tetraselmis sp. and Chlorella in industrial-scale bioreactors, which produce 2.33 and 2.44 % w/w lipid (calculated as the sum of fatty acid methyl esters) in dry biomass, respectively. These lipids contain higher amounts of neutral lipids and glycolipids plus sphingolipids, than phospholipids. Lipids of Tetraselmis sp. are characterized by the presence of eicosapentaenoic acid (that is located mainly in phospholipids), and octadecatetraenoic acid (that is equally distributed among lipid fractions), while these fatty acids are completely absent in Chlorella lipids. Additionally, lipids produced by 16 newly isolated strains from Greek aquatic environments (cultivated in flask reactors) have reported the highest percentage of lipids in Prorocentrum triestinum (3.69 % w/w) while the lowest in Prymnesium parvum (0.47 % w/w). Several strains produce lipids rich in eicosapentaenoic and docosahexaenoic acids. For instance, docosahexaenoic acid is found in high percentage in lipids of Amphidinium sp. S1 and Prorocentrum minimum, while lipids produced by Asterionella sp. S2 contain eicosapentaenoic acid in high concentration. These lipids, containing ω-3-long-chain polyunsaturated fatty acids, have important applications in the food and pharmaceutical industries and in aquaculture (Makri et al. 2011).

Large-scale cultivation

The cell density of P. lutheri under 250 mL, and 1, 5, 30 and 300 L is shown in Fig. 3. The highest μ _max of 0.14 day⁻¹ and the doubling time of 4.95 days were achieved in 30-L tank cultivation (Table 3) although the highest cell density of 13.5 × 10⁶ cells mL⁻¹ and dry weight of 0.45 g L⁻¹ were obtained in 250-mL flasks. The goal of reproducing P. lutheri culture at 5–300 L was successfully achieved with cell density of 9–11 × 10⁶ cells mL⁻¹ and cell dry weight of 0.35–0.41 g L⁻¹ which were only slightly lower than 12–14 × 10⁶ cells mL⁻¹ and 0.43–0.45 g L⁻¹ at 250 mL to 1 L. The lowest cell density of 9.65 × 10⁶ cells mL⁻¹ and dry weight of 0.35 g L⁻¹ attained in a 300-L tank suggest the need to address the scale-up issues in an open-pond tank with regards to cultivation conditions, mixing, sampling and harvesting. The biochemical compositions of microalgae can change with their growth rates and environmental conditions and with the phase of their life cycle. Unlike laboratory conditions, outdoor natural climates are more complex and rigorous. It may be possible that variations of outdoor natural conditions influence microalgal growth rate and biochemical constituents. Cultivation of microalgae in open-pond tanks faces challenges such as overheating, fouling, accumulation of oxygen to toxic levels (Hu et al. 2008) and contamination with bacterial species.

Fig. 3

Comparison of growth curve of P. lutheri cultivated at 250 mL, and 1, 5, 30 and 300 L

Table 3 Comparison of kinetics between 250-mL and large-scale batch cultures of P. lutheri at optimized conditions (pH 8, salinity of 35 ppt, photoperiod of 24 h and illumination of 130 μmol photons m⁻² s⁻¹ intensity; just the 300-L tank was under sunlight illumination)

Most commercial, large-scale outdoor microalgae cultivation is in artificial open ponds as they are cheap to build and easy to operate and scale-up (Brennan and Owende 2010). The raceway pond has relatively low capital and maintenance costs while circular ponds are less attractive because of expensive concrete construction, high energy consumption of stirring, the mechanical complexity of supplying CO₂ and inefficient land use (Chen et al. 2009). Apart from disadvantages such as low productivity and biomass yield, high harvesting cost, water loss through evaporation, and the limited number of species that can be grown in ponds due to contamination, there may be low carbon dioxide consumption efficiency and temperature fluctuations due to diurnal variations, as they are difficult to control in open ponds (Chen et al. 2009).

Estimations of the production of algal oil as being able to reach over 100,000 L per hectare per year (compared with just 450 L for soybean oil and 6,000 L for palm oil) seems attractive (Burns 2010). However, what is feasible in the laboratory may not be achievable outdoors on a very large scale. With current technology, microalgae are too expensive to be viable alternatives to the major commodity plant oils. The challenge to make algal oils more economical will be in the form of open-pond cultivation with robust, fast-growing algae that can withstand adventitious predatory protozoa or contaminating bacteria, whilst attaining an oil content of at least 40 % of the biomass (Ratledge and Cohen 2008). If the prices of the major plant oils and crude oil continue to rise in the future, algal lipids may become a realistic alternative with better prospects focusing on algae as sources of polyunsaturated fatty acids, and a major genetic redesign of algal metabolic processes (Ratledge and Cohen 2008; Ratledge 2011). Cost-effective and energy-efficient harvesting and lipid extraction technologies need to be developed. Further research needs to focus on exploring integrated bioprocesses and genetic engineering aspects for the improvement and enhancement of microalgae as an optimum biofuel source.

Conclusions

The naturally occurring microalgal strain Pavlova lutheri was a good candidate for biodiesel production, because of high lipid content. The optimum growth and lipid production was obtained under 24 h illumination, salinity ranges of 30–40 ppt and pH ranges of 7.5–9 with maximum specific growth rate μ _max of 0.13 day⁻¹ and lipid content of 36 %. Cultures grown at 30 L had highest μ _max of 0.14 day⁻¹, while X _max was highest at 0.43–0.45 g L⁻¹ at 250-mL and 1-L scales. Scaling-up of P. lutheri and the effects of environmental conditions have been successfully established for further optimization.