Enzymatic in-situ transesterification of neutral lipids from simulated wastewater cultured Chlorella emersonii and Pseudokirchneriella subcapitata to sustainably produce fatty acid methyl esters

Alternative, more sustainable and environmentally positive, sources of energy are one of the current global challenges. One approach to achieving more sustainable sources of energy is to use waste from one system as a raw material for energy production, following the circular


Introduction
Fossil fuels, such as petrol, coal, and natural gas, contribute 88% of global energy consumption; with the transport sector subsequently being a major contributor of CO2 global emissions, which, by 2050 is estimated to increase to 2 billion vehicles (Balat and Balat, 2010). More urgently, by 2030, the emission of total anthropogenic greenhouse gases is projected to increase by 23 percent (Ullah et al., 2015). This environmental concern, coupled with the growing global energy demand, has resulted in increased interest in the production of environmentally friendly and sustainable liquid fuels, such as biofuels, as a suitable alternative source of energy (Milano et al., 2016). Biofuels, comprising Fatty Acid Methyl Esters (FAMEs), are typically produced from renewable biological oils and fats, such as vegetable oil, animal fats or nonedible plant feed stocks and can sometimes be used directly in existing engines with little or no modification (Scott et al., 2010). Currently, alternative sources of lipids, such as microalgae, are currently being bioprospected.
Microalgae; due to a short harvesting cycle and high growth rates allow rapid accumulation of significant amounts of lipids, with appropriate culturing conditions. Microalgae are highly adaptable towards their surrounding environment; they do not require fertile land and can therefore be grown almost anywhere, even on treated sewage or saltwater. Since microalgae require CO2 to grow and can efficiently remove phosphates and nitrates from wastewater, they are an appropriate biomass for bioremediation and biofixation (Lam and Lee, 2012). The lipid content in microalgae varies from 20-40% biomass dry weight, however, lipid content as high as 85% biomass dry weight has also been reported in certain microalgal strains (Mairet et al., 2011). Optimizing the metabolic pathways of microalgae cells can increase the lipid content (Lowrey, Brooks and McGinn, 2015). Therefore, utilising a short harvesting cycle, coupled to metabolic pathway optimisation to produce higher lipid content and with continuous harvesting suggests that microalgae may be very good candidates for sustainable fuel production (Miao and Wu, 2006). In the current study this hypothesis has been explored where two microalgae strains, Chlorella emersonii and Pseudokirchneriella subcapitata, were cultured and metabolically engineered in simulated wastewater for neutral lipids.
Nile Red dye was used for quantification of neutral lipids in the current study as a high correlation between neutral lipid content (as per the gravimetric method) and Nile Red fluorescence has been established and the assay is used as the quantitative analysis method for neutral lipid quantification in a variety of microalgae, such as Chlorella sp.  species also dependent on the composition and structure of the microalgal cell wall. Robust and thick cell walls (particularly in green algae and nutrient starved microalgae) act as a barrier; preventing efficient penetration of Nile Red in cells and neutral lipid staining (Doan and Obbard, 2011;Guzman et al., 2011;Pick and Rachutin-Zalogin, 2012). Hence, for the estimation of neutral lipids in Chlorella emersonii and Pseudokirchneriella subcapitata in the present study, a Nile Red assay was developed.
Microalgae lipid extraction usually follows two steps: cell disruption and solvent extraction.
Disruption of the microalgal cell wall for the extraction of oil in an extraction solvent increases the energy requirement and cost due to the additional steps of dewatering and drying of microalgal cells (Ronald, Micheal and Paul, 2012). Several mechanical, chemical or combinational cell (ultrasound, high-pressure homogenization, bead-beating, osmotic shocks) methods have been suggested to facilitate oil extraction from algal cells (Halim et al., 2012;Ronald, Micheal and Paul, 2012). However, due to increased processing costs and time, other transesterification methods are required. In the current study, FAMEs, as a precursor for biodiesel, were synthesized via in-situ transesterification using novel lipases and the neutral lipids from the biomass of Chlorella emersonii and Pseudokirchneriella subcapitata.
Lipases can catalyse both hydrolysis and synthesis of long-chain acylglycerols and most importantly can catalyse biodiesel production since they are the only enzymes that catalyse the synthesis of esters (i.e. transesterification (Jaeger and Reetzb, 1998). Traditionally, the majority of industrial scale transesterification reactions are carried out in organic solvents due to the ease of solubility of the non-polar lipid substrates. However, most lipases are denatured in organic solvents and, therefore, lose their catalytic activity. In an alternative approach, this study employed novel, solvent stable lipases from P. reinekei (H1) and P. brenneri (H3) for the in-situ transesterification of neutral lipids from Chlorella emersonii and Pseudokirchneriella subcapitata to produce sustainable, economical and environmentally positive FAMEs that were chemically comparable to commercially available biodiesel FAMEs mix.

Biomass generation and neutral lipid production:
Microalgae cultures were grown in simulated wastewater media under three different conditions.
Growth in autotrophic condition involved photosynthesis (presence of light and CO2); in mixotrophic condition involved photosynthesis along with 1% (w/v) glucose as carbon source; while heterotrophic condition eliminated photosynthesis (cultured in the dark) but utilised only 1% (w/v) glucose as a carbon source. The culturing conditions resulting in highest biomass were subsequently used for the generation of microalgal biomass as the first step of neutral lipids production. The microalgal biomass generated in the relevant growth media was harvested by centrifugation at 5,000*g for 5 mins at 18°C and was washed twice with sterile autoclaved distilled water before further experimentation. The washed cell pellet was transferred to simulated nitrogen deficient wastewater with and without 1% (w/v) glucose. The culture was mixed thoroughly by shaking the conical flask in a rotatory motion for uniform cell distribution.

Nile Red assay:
The Nile Red assay, for the estimation of neutral lipids in Chlorella emersonii and Pseudokirchneriella subcapitata was carried out using fixed number of cells (0.6 OD@590nm; with 1 O.D@590nm=10 7 cells/ml) from the simulated wastewater growth media. The cells were transferred to nitrogen deficient wastewater media and after one week of culturing (18°C with 16h:8h (light: dark) cycle, 120rpm) in nitrogen deficient wastewater; 10mls of culture were aseptically removed into a universal tube. The culture was centrifuged at 5,000*g for 5mins and the culture media was discarded. The microalgal cell pellet was then mixed thoroughly in 10mls of double distilled water and, subsequently, 50µl of this was used for assay. The assay was carried out in quintuplets utilising a flat bottom transparent 96-well microtiter plate. Into each well 100µl of double distilled water, initially, and 50µl of microalgae culture, subsequently, were added followed by 100µl of Nile Red dye. Neutral lipid estimation in Chlorella emersonii and Pseudokirchneriella subcapitata was carried out using Nile Red dye with 5µg/ml and 10µg/ml For quantification of neutral lipids, a working range of triolein standard (between 1µg/ml to 15µg/ml) was prepared in neat chloroform. For the quantification, in individual well of a 96-well plate, 100µl of ddH2O, 100µl of 10µg/ml nile red (in 20% v/v DMSO) and 50µl of the respective triolein concentration was mixed, in quintuplet. Fluorescence intensity was measured at 530nm excitation and 580nm emission wavelength after 5mins of incubation. The fluorescence intensity from various concentrations of triolein was used for standard curve preparation and for neutral lipids quantification in microalgae biomass.

In-Situ Transesterification
In-situ transesterification of microalgae biomass was executed as per Tran and co-workers' protocol (Tran et al., 2012) with minor deviations. In brief, 0.5g of freeze-dried microalgae biomass was mixed with 20ml of neat methanol and sonicated at 70 amplitude for 20mins, using QSonica Q55 sonicator. After sonication, methanol was evaporated by placing the container in a fume hood for 60mins to obtain an oil-containing slurry. This slurry was later mixed, by vortexing for 5mins with 5ml of n-hexane. 500IU of lipase/g of microalgae oil was subsequently added to the mixture along with 5ml of methanol to initiate the transesterification reaction at 40°C for 72h. To overcome the issue of solvent evaporation; each reaction was carried out in a sealed glass container. After transesterification, the sample was centrifuged at 5,000*g for 10mins and the upper solvent layer containing FAMEs was pipetted carefully to a clean sealed glass container. The FAMEs generated were analysed by Thin Layer Chromatography (TLC) and Gas Chromatography (GC). The yield of biodiesel from microalgae biomass was calculated as per (Cao et al., 2013); 3 100

Thin Layer Chromatography and Gas Chromatography
Post transesterification TLC FAMEs detection was carried out as per Kim and colleagues (Kim et al., 2014) with no deviations. In brief, a 90:10 (v/v) n-hexane: diethyl ether solvent mix was used as the mobile phase and after full development of TLC plate, the FAME spots were visualized using a 10% (v/v) ethanoic phosphomolybdic acid spray, followed by drying at 105°C for 5mins.
The GC method of (David, Sandra and Vickers, 2005) was used for FAME analysis. In brief, a Bruka GC column (0.25mm Internal Diameter and 30m Length) operating as part of a 0.25µm particle size Scion-436GC machine with FID detector, was used for FAME analysis. FAMEs identification was carried out by comparing their Retention Time (RT) of the product with RT of a 37-component FAME standard mix (Sigma).

Experimental Scheme
3. Results and Discussion:

Chlorella emersonii and Pseudokirchneriella subcapitata biomass generation
Mixotrophic cultivation improves the efficient use of light or eliminates its requirement by cells; CO2 and organic carbon are used in photosynthetic and respiratory metabolism simultaneously, resulting in a synergistic effect of autotrophic (carbon dioxide fixation by Calvin-Bensen cycle) and heterotrophic processes (Yeesang and Cheirsilp, 2014

In-situ transesterification
Furthermore, glucose is one of the final photosynthesis products; and would suggest that any photosynthetic microorganism must be able to incorporate it in its metabolism (Yeesang and Cheirsilp, 2014). Therefore, in the current study glucose was used as a source of organic carbon for mixotrophic and heterotrophic growth conditions for the microalgae strains. Figures 1 and 2 depict the growth curves, in different wastewater culturing conditions, for Chlorella emersonii and Pseudokirchneriella subcapitata respectively.  (Lee et al., 1996). Likewise, maximum biomass of 2.46g/L was also obtained for Botryococcus braunii cultured in mixotrophic conditions, where 5g/L of glucose was used as an organic carbon source (Yeesang and Cheirsilp, 2014). In the current study, to obtain a higher biomass of Chlorella emersonii and Pseudokirchneriella subcapitata, a 5-day and 8-day mixotrophic culturing mode respectively was found to be optimum. Furthermore, a difference in physiological appearance was observed for both the cultures for different growth conditions (see Appendices 1 and 2), indicating a reduced production of chlorophyll in mixotrophic and heterotrophic mode.

3.2.Neutral lipid production in nitrogen deficient conditions
The ability of microalgae to survive or proliferate over a wide range of environmental conditions can be due to unusual cellular lipids, as well as an ability of microalgae to modify lipid metabolism, in response to altered environmental conditions (Thompson, 1996;Wada and Murata, 1998; Gouveia and Oliveira, 2009). Shen and colleagues (Shen et al., 2009) have demonstrated that limiting certain nutrients (e.g. nitrogen) can result in higher lipid production and storage, as the microalgal cells respond to the stress conditions imposed. Spoehr and Milner (Spoehr and Milner, 1949), first demonstrated that lipid content could be increased (from 5% to 85%) in nitrogen starved Chlorella pyrenoidosa culture, and since then nutrient deficiency/depletion (particularly nitrogen) has been regarded as the most efficient approach to increase lipid content in algae (Rodolfi et al., 2009).
In the current study, the depletion of nitrogen sources from simulated wastewater media in cultivation of Chlorella emersonii ( Figure 3) and Pseudokirchneriella subcapitata (Figure 4) resulted in the production of maximum neutral lipids after 6 and 12 days of incubation respectively. The increase in lipid production under nitrogen limited conditions is typically due to the disorganization of lipid synthesizing enzymes being less than compared to carbohydrate synthesizing enzymes; thus, the major proportion of carbon is fixed in lipids (Becker, 1994). A high concentration of the appropriate nutrients is necessary for a high growth rate and maximum biomass; but accumulation of a large concentration of lipids normally cannot take place during such growth. Culturing under a nitrogen deficient conditions leads to higher lipid accumulation, but with lower biomass, and vice-versa. In order to balance the challenges associated with simultaneous production of biomass and neutral lipids; a two-stage cultivation process can be followed, as executed in this study. The first step of cultivation focuses on microalgal biomass generation by providing optimum nutrients. In the second stage, the biomass from the first stage is harvested, washed with deionized water to remove any traces of nitrogen or other nutrients from stage one and then the biomass is grown in nitrogen deficient media.
The absence of nutrients (nitrogen or phosphorous) in stage two of the cultivation process reduces the biomass however; incorporation of an organic carbon source like glucose or sodium acetate is known to enhance the lipid production along with biomass. A similar two-stage approach was also carried out to enhance neutral lipid content in Nannochloropsis gaditana, Tetraselmis chuii, Tetraselmis suecica and Phaeodactylum tricornutum (Pedro et al., 2013) and in Chlorella vulgaris (Cui et al., 2017). Heterotrophic lipid generation conditions in Chlorella protothecoides have demonstrated an increase in lipid content from 15% to 55% (Miao and Wu, 2006). Likewise, the addition of 4g/L sodium acetate in nitrogen and phosphorous deficient media increased total lipid content by 93% in Chlamydomonas reinhardtii (Yang et al., 2018).
This mirrors the observations in the current study; following the two-step cultivation for concentration of triacylglycerols in Chlorella emersonii and Pseudokirchneriella subcapitata after following the two step cultivation; 13 and 21 days of culturing in nitrogen deficient media with 1% (w/v) glucose was calculated to be 0.61±0.017mg/mg and 0.31±0.006mg/mg of biomass respectively.

3.3.In-situ transesterification of microalgae lipids to produce Fatty Acid Methyl Esters
In-situ transesterification of Chlorella emersonii and Pseudokirchneriella subcapitata