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Enhanced biodiesel production through phyco-myco co-cultivation of Chlorella minutissima and Aspergillus awamori: An integrated approach
Accepted Manuscript Enhanced biodiesel production through phyco-myco co-cultivation of Chlorella minutissima and Aspergillus awamori: an integrated approach Archana Dash, Rintu Banerjee PII: DOI: Reference:
S0960-8524(17)30529-1 http://dx.doi.org/10.1016/j.biortech.2017.04.039 BITE 17934
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Bioresource Technology
Received Date: Revised Date: Accepted Date:
27 January 2017 8 April 2017 10 April 2017
Please cite this article as: Dash, A., Banerjee, R., Enhanced biodiesel production through phyco-myco co-cultivation of Chlorella minutissima and Aspergillus awamori: an integrated approach, Bioresource Technology (2017), doi: http://dx.doi.org/10.1016/j.biortech.2017.04.039
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Enhanced biodiesel production through phyco-myco co-cultivation of Chlorella minutissima and Aspergillus awamori: an integrated approach Archana Dash and Rintu Banerjee* Agricultural and Food Engineering Department, Indian Institute of Technology Kharagpur, Kharagpur-721302, India
* Address of corresponding author Prof Rintu Banerjee Department of Agricultural and Food Engineering, Indian Institute of Technology Kharagpur- 721302, India Tel: +91-3222-283104(O); Fax: +91-3222- 282244 E-mail: [email protected]
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Abstract Algae-fungus co-culture was investigated as an alternative biodiesel feedstock. An oleaginous filamentous fungus Aspergillus awamori was co-cultured with Chlorella minutissima MCC 27 and Chlorella minutissima UTEX 2219, respectively in N11 medium furnished with different carbon and nitrogen sources. The biomass and lipid production potential of the two C. minutissima–A. awamori co-cultures was compared against the monocultures. A substantial enhancement in biomass and lipid accumulation was observed in both the co-cultures. When supplemented with different carbon and nitrogen sources, glycerol and potassium nitrate were found to be the most effective. In the presence of glycerol, a 2.6-3.9 fold increase of biomass and 3.4-5.1 fold increase of total lipid yields were observed in the co-cultures as compared to the axenic monocultures. Furthermore, C16:0 (31.26-35.02%) and C18:1 (21.14-24.21%) fatty acids were the major composites of the co-culture oils, which suggest co-culture as a promising strategy for biodiesel production.
Keywords: Biodiesel; Chlorella minutissima; Aspergillus awamori; Co-culture; FAME
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1. Introduction Green energy, an alternative to rapidly-decreasing fossil fuels, has received a lot of attention and popularity in recent times. It is clean, renewable, and sustainable which can provide a robust, massive, and enduring energy supply. In this context, microalgae share many explicit advantages as a third generation biofuel. The subdued food vs fuel issue, round-the-year farming, higher biomass turnover per unit area with high photon-to-biomass conversion, environment-friendly exhaust emission profile, CO2 bio-mitigation through photosynthetic-fixation, substantial TAG/lipid storage, independence of the water quality for cultivation, and valorization of defatted biomass by the production of high-value, lowvolume products have made microalgae more advantageous than their oleaginous terrestrial competitors (Sayre, 2010; Sharma et al., 2012; Rai et al., 2015). Despite these positive accounts, there are several serious constraints which are a hindrance in the algae-to-diesel reality. First and foremost, one of the major issues is the slow-growing nature of the oleaginous species of the microalgae (Sharma et al., 2012) and their mutually incongruous nature of biomass and lipid accumulation pattern (Li et al., 2011). In most of the microalgae, high lipid contents are usually at the expense of biomass reserve of the cells under nutrient-deprived conditions (Zhu et al., 2016). Consequently, there is often a significant loss of total lipid yields which is the immediate substrate for transesterification i.e. biodiesel production. Besides, issues such as high-cost harvesting techniques, poorly sustainable and renewable nutrient supply, inefficient and costly lipid extraction techniques coupled with poor lipid content and fatty acid (FA) composition (Molina Grima et al., 2003) are yet to be properly addressed.
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The technical challenges faced in harvesting microalgae are due to the tiny cell size (2-20 µm), negative charge (−7.5 to −40 mV) on its surface, low density (0.3–5 g L-1), and colloidal stability in suspension (Liu et al., 2013; Li et al., 2008). Due to the negative charge on its cell surface, oleaginous microalgae overcome the force of natural gravity and remain suspended in the aqueous medium. The dewatering of microalgal biomass often involves one or more solid-liquid separation stages, which account for 20-30% of the gross cultivation cost of microalgae (Molina Grima et al., 2003; Zhou et al., 2015). The two widely practiced techniques for large-scale harvesting of microalgae by the industries are filtration and centrifugation. Centrifugation was found to be a cost-intensive process for the production of a high-volume, low-value product like biodiesel. On the other hand, conventional filtration method is a simpler and cheaper technique. However, for harvesting extremely small-dimensioned microalgae such as C. minutissima and Nannochloropsis oculata, this method alone cannot be effective and efficient. It may lead to membrane clogging, fouling and frequent replacement of the filter causing a many-fold increase in the overall cost of the process obviating its large-scale application (Singh et al., 2013). In this regard, filamentous fungi can be a possible solution because they can be utilized for bioflocculating microalgal biomass (Zhou et al., 2015) and subsequently easily separated through conventional filtration method. The fungal-assisted bio-flocculation technique requires less energy input and being free of chemical contaminants facilitates proper downstream processing of the harvested biomass and recycling of the spent medium (Li et al., 2008; Zhou et al., 2015). It shows immense potential for harvesting the microalgal cells as a relatively easy and cheaper technology compared to other conventional harvesting techniques. Besides these advantages, another important aspect of the co-culture is the 4
mutual behavior of the microalgae and filamentous fungus, which could be of great advantage. Obligate mutualism has been seen between a few photobionts and fungi in nature (Wrede et al., 2014). However, under synthetic co-culture conditions, it would be interesting to see their corresponding growth and lipid accumulation behavior. The consortia diversity could induce significant changes in the nutrient flux affecting the growth and subsequent modulation of the metabolic fluxes can bring about shifts in lipid metabolism of the oleaginous entities during their co-culture. In this study, two oleaginous alga-fungus consortia were investigated for biodiesel production. The consortia employed in this study were the amalgamation of two different strains of green microalgae, C. minutissima (MCC 27 and UTEX 2219) and a filamentous fungus, A. awamori. These two microbial entities of the co-culture systems are from widely different phylogenetic origins and have multiple nutrient-selectivity. For example, green algae are photoautotrophs while fungi are obligate heterotrophs. When these microorganisms are grown in a single system under mixotrophic nutritive mode, relative selectivity of the nutrients and the trophic modes for both may vary. Hence, the selectivity and suitability of the different carbon and nitrogen sources for the co-cultures were investigated. Also, the growth of each component can be affected by each other’s growth tendencies under defined conditions. These changes can bring about significant difference in the cumulative biomass and lipid yield of the consortia. Moreover, the interaction between the algae and filamentous fungus may also modulate the fatty acid composition in synergy, and subsequently can affect the biodiesel properties as well. Hence, the subsequent effect on the collective biomass and lipid yields by the co-culture systems and the changes in the FAME profiles were subjected to investigation in the presented study. 5
2. Materials and methods 2.1. Microorganisms, inoculum preparation, and culture conditions C. minutissima MCC 27 and C. minutissima UTEX 2219 were procured from the Center for Conservation and Utilization of Blue Green Algae (CCUBGA), IARI, New Delhi, India and Culture collection of Algae, University of Texas, Austin, respectively. A. awamori was previously isolated from the native soil of Kharagpur, West Bengal, India. The axenic cultures of microalgae were maintained in N11 medium (Saxena et al., 2016) under 6.8 pH (pH meter, M/s. ELICO, India), 25 ± 2 °C, 75 µmol photon m-2 s-1 PAR light intensity and 14:10 h (L:D) photoperiod. The algal cells were harvested using a centriguage (eppendorf centrifuge 5810R, 5000 rpm, 15 min) and subsequently washed twice with distilled water. Further, the cell counting was carried out using a hemocytometer (FeinOptik, Bad Blankenburg, Germany) and maintained by dilution with distilled water. A. awamori was maintained on Potato Dextrose Agar (PDA) at pH 5 and 30 ± 1 °C. Fungal spores were extracted from the 6-day-old culture from the petri dishes with 10 mL distilled water. After washing twice with distilled water, the fungal spores were counted under the microscope. The initial spore count was maintained by diluting the resultant spore solution with distilled water. Equimolar concentrations of different carbon sources viz. glucose, glycerol, and acetate and nitrogen sources such as potassium nitrate, urea, and yeast extract were supplemented in the growth medium. Incubation temperature (25 °-30 °C) and initial pH (5-8) were varied to investigate for maximum biomass and lipid accumulation. Supply of exogenous air or sparging of CO2 was avoided in this study. Only manual gentle shaking
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for 10 minutes was done thrice a day to prevent sticking of the cells with the wall and for uniform growth. 2.2. Growth study The growth of C. minutissima MCC 27 and UTEX 2219 in monocultures and cocultures was measured spectrophotometrically (UV-1800, SHIMADZU, Japan) at 540 nm. The dry cell weight of monocultures of both C. minutissima MCC 27 and A. awamori as well as the co-cultures was determined gravimetrically. The entire biomass was harvested through centrifugation and oven-dried (60 °C) in a pre-weighed glass vial. The biomass was dried till it achieved a constant weight and finally the dry cell weight was determined from the difference between the final and empty vial weight. Biomass of microalgae in the co-cultures was determined indirectly by measuring the Chla concentration in the co-culture and subsequently the biomass was determined from the standard curve of Chla concentration and dry cell weight of the microalgae. For chlorophyll extraction, chilled methanol (99.8 %) was used followed by a sonication bath for 15 minutes and kept overnight. Chla concentration was measured spectrophotometrically at 652 and 665 nm (Porra et al., 1989). The Chla and biomass of microalgae were correlated as per the following equation: Microalgae dry biomass (mg) =22.944 × A652 [R2 = 0.99] (equation 1) After obtaining the biomass of microalgae from the equation 1, biomass of A. awamori in the co-culture was determined from the difference of total biomass and biomass of microalgae. 2.3. Lipid extraction
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Lipid was extracted from the oven-dried microbial biomass by following the Bligh and Dyer (1959) protocol. After the extraction, lipid was filtered via filter paper (Whatman No. 1) and stored in a pre-weighed vial (W1). The solvent was evaporated in a water bath followed by an oven-drying (104 °C) till the constant weight was recorded consecutively for three times. The final weight of the vial was measured (W2). The amount of the lipid was estimated from the difference of W1 and W2, and expressed as % dcw as per the following formula:
Lipid content (% dcw) =
Mass of lipid Mass of biomass
× 100
2.4. Biodiesel production 2.4.1. Lipase production, extraction, and assay Lipase was produced indigenously from Rhizopus oryzae NRRL 3562 under solid state fermentation (SSF). Czapek-dox medium (Sarkar et al., 1998) was used in 1:1 ratio to moisturize the substrate (wheat bran) which was used as the major carbon source. The fermentation medium was further fortified with coconut oil (Garlapati and Banerjee, 2010). After inoculation with R. oryzae, the medium was incubated for the completion of the required fermentation, thereafter extracellular lipase was extracted for subsequent studies. The crude enzyme was assayed for lipase activity, which was determined spectrophotometrically by using p-nitrophenylpalmitate (pNPP) as substrate (Pencreac’h and Baratti, 2001). Total protein content was determined through modified Lowry method by using bovine serum albumin (BSA) as standard (Lowry et al., 1951). 2.4.2. Enzymatic transesterification
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The indigenously produced lipase was processed for transesterification studies. Lipasemediated transesterification was carried out by taking oil to methanol molar ratio 1:3 at reaction conditions of 37 °C, 200 rpm and 24 h. 10 U immobilized lipase g-1 of oil was used for the reaction. Immobilization of lipase on activated celite (50 U g-1) and preparation of sample for GCMS analysis were done as per the protocol of Garlapati et al. (2013). 2.4.3. FAME analysis FAME profiling was done through GC-MS analysis (Agilent 6890 N gas chromatography paired with Agilent MS-5975 inert XL mass selective detector; Agilent Technologies, Little Fall, NY, USA). HP-5MS (5 % phenyl methylsiloxane) capillary column (Agilent Technologies, Palo Alto, CA, USA) with 30 m х 0.25 mm х 0.25 mm dimension was used. The initial temperature was maintained at 150 °C and raised to 280 °C at the rate of 4 °C min-1, and kept at 280 °C for 9 min. The flow rate was 0.8 mL min-1. The injector and detector temperatures were 250 °C and 260 °C, respectively. For FAME identification, retention times of isolated compounds were matched with that of FAME standards. In addition, by using the National Institute of Standards and Technology (NIST) mass spectral library, mass fragment pattern of each resolved peak was compared.
3. Results and discussion 3.1. Growth and lipid accumulation in monoculture and co-culture The biomass and lipid accumulation have been investigated for a period of 28 days. Maximum biomass was observed on the 21st day of incubation in both the strains. Maximum biomass accumulation was seen in C. minutissima MCC 27 (1.140 ± 0.087 g L-1) followed by C. minutissima UTEX 2219 (0.900 ± 0.050 g L-1). Substantial lipid 9
accumulation was observed in the late exponential phase of their growth. C. minutissima MCC 27 and C. minutissima UTEX 2219 were found to accumulate 22.12 ± 0.99 % and 20.00 ± 1.00 % dcw lipid, respectively. C. minutissima MCC 27 was observed to be better biomass producer and lipid accumulator as compared to C. minutissima UTEX 2219. Fig. 1A depicts the growth and lipid production profile of the two different microalgae monocultures i.e. C. minutissima MCC 27 and C. minutissima UTEX 2219. The growth of C. minutissima MCC 27 as well as UTEX 2219 was monitored for 9 days under autotrophic, mixotrophic (1% glucose) and co-culture mode (1% glucose) and compared. Interestingly, an approximately 12.4-fold rise in the growth of C. minutissima MCC 27 in co-culture system was observed as compared to autotrophic axenic culture, while it was 1.8-fold more than the mixotrophically grown axenic test algae on the 6th day of incubation (Fig. 1B). Similar trends were also obtained in case of C. minutissima UTEX 2219/A. awamori co-culture. Increase in the biomass accumulation of C. minutissima UTEX 2219 in the co-culture with A. awamori were up to approximately 13.9- and 2.8-fold as compared to autotrophic and mixotrophic axenic C. minutissima UTEX 2219, respectively (Fig. 1B). The significant increase in the growth of test microalgae and significant biomass accumulation in presence of A. awamori indicate that the diversity of the consortia selected for this study may have turned out to be the triggering factor for the improved growth of these species. The relative contribution towards the productivity in the co-culture systems between the two species were comparable. Biomass and lipid yields in the co-cultures were cumulatively as well as individually more than the monocultures. Consortia diversity in cocultures can function as an efficient system, which can help in better utilization of the 10
resources. When the utilization of resources is complementary, ecologically different species co-exist (Gause, 1936; Weis et al., 2008). This way co-cultures can harness a greater fraction of the limited resources compared to the monoculture of the most productive species and result in enhanced biomass of the latter (Weis et al., 2008). Furthermore, when diverse species have to compete for the limiting resources, they complement each other in the utilization of their resources (Loreau et al., 2001; Korhonen et al., 2011) which can turn out to have a positive effect on the biomass accumulation in the co-culture system. One such example of complementary behavior and better resource utilization in presence of co-culture is reported by Xue et al. (2010). According to their report, incorporation of Spirulina platensis to the yeast culture of Rhodotorula glutinis led to significant enhancement of O2 in the co-culture. Inoculum composition was varied by changing the ratio between fungal spores and algal cells. The inoculum cells count ranged from 1:110 to 1:300 (fungal spores: algal cells). An increasing trend in biomass of co-culture was observed with an increase in the ratio of spore:algal cells on the 6th day of incubation (Fig. 2). Maximum biomass yields were found to be 2.80 ± 0.1 g L-1 and 2.54 ± 0.04 g L-1 for C. minutissima MCC 27 and C. minutissima UTEX 2219 respectively for 1:300. The increase of algal cell numbers in the inoculum composition was observed to have a prominent effect on the total lipid yield (mg L-1) and content (% dcw). Within the range 1:215 to 1:300, a minor difference of lipid content was observed in case of the C. minutissima MCC 27/A. awamori co-culture, whereas it was found comparable in case of the C. minutissima UTEX 2219/A. awamori co-culture in the range 1:205 to 1:300. However, overall maximum total lipid yield was observed with 1:300, which was 510.4 mg L-1 and 437.13 mg L-1 for C. minutissima MCC 11
27 and C. minutissima UTEX 2219, respectively. Similarly, the effect of incubation temperature on the co-cultures of C. minutissima MCC 27/A. awamori and C. minutissima UTEX 2219/A. awamori has been investigated. It was found that 25 °C was observed to be optimum for the growth of test algae while 30 °C was most suitable for A. awamori. However, in case of co-cultures, 25 °C was found to be the most suitable for biomass production of the co-cultures. A minor decrease in biomass was observed in the case of C. minutissima UTEX 2219/A. awamori co-culture when the incubation temperature increased from 25 ° to 30 °C. Overall the biomass production and lipid accumulation patterns were comparable for C. minutissima MCC 27/A. awamori co-cultures, while a slight decrease in biomass production and increase in lipid content (% dcw) were observed with C. minutissima UTEX 2219/A. awamori co-culture. However, the overall effects of incubation temperature were insignificant for both the co-cultivation systems suggesting tolerance of both the co-cultures within the range (25°-30 °C). When the effect of initial pH on the growth and lipid accumulation potential of the co-cultures were investigated, it was seen that pH 5 was found to be the most suitable followed by pH 8 for the consortial biomass yield for both the co-culture systems while pH 5 was found to yield maximum lipid contents. Optimum pH for the growth of C. minutissima was found to be 7.0; on the contrary, pH 5.0 was found to be the most suitable for the growth of A. awamori. The pH 5.0, 5.5 and 8.0 showed comparable biomass yield in both the co-cultures. It suggests that slightly acidic and slightly alkaline growth medium is best suited for the growth of both the cultures. Many Ascomycota fungi were known to secret different organic acids such as acetic acid, citric acid, lactic acid, oxalic acid, malic acid, etc., into the growth medium (Liaud et al., 2014). These organic acids severely affect the growth of the liable fungus by 12
lowering the cytosolic pH and malfunctioning the permeability of the cell membrane causing its death (Hassan et al., 2015). Yeast releases CO2 as a byproduct of its respiratory metabolism, and it readily dissolves in the aqueous growth medium. As a result, bicarbonate (HCO3 ̶ ) is produced under pH ≥ 7 (Sayre, 2010), which causes a further decline in pH. Microalgae can biologically capture CO2 by concentrating HCO3 ̶ intracellularly, later again get converted into CO2 spontaneously or by carbonic anhydrase (Moroney and Somanchi, 1999). The intracellular CO2 is then fixed by the algal photosynthetic machinery to be reciprocated into algal biomass. The OH ̶ produced in this process helps in increasing the pH towards alkalinity (Richmond, 1986). Microalgae can metabolize different organic acids such as acetic acid, malic acid, pyruvic acid, lactic acids (fungal metabolites) for its growth (Bollman and Robinson, 1977). Hence, the co-culture of fungus with microalgae could have been helpful in assuaging the inhibitory effects of organic acids on the growth of the fungus. On the other hand, the optimum pH for the growth of C. minutissima was found to be 7.0. Rai et al. (2015) also reported maximum growth at pH 7.0 while Sankar et al. (2011) reported optimum pH for the growth of C. minutissima as 6.0. During microalgae cultivation, pH tends to increase and becomes alkaline as a result of OH ̶ production during CO2 assimilation (Bi and Zhou, 2016). Thus, microalgae, in co-culture, could help in buffering of the medium for mutual growth of both the co-culturants. 3.2. Selection of suitable carbon and nitrogen sources for co-culture In this study, an attempt has been made to investigate the effect of various carbon sources on biomass and lipid accumulation of co-cultures as well as monocultures of microalgae and fungus. The type of carbon source employed for the growth of oleaginous 13
species may play a crucial role in steering the flux of carbon towards lipid. In microalgae, intracellular accumulation of neutral lipids occurs from the bioconversion of starch/carbon to lipids through different mechanisms depending on the strains (Li et al., 2011). Application of different carbon sources can amend the core carbon metabolism by driving the carbon-partitioning towards the lipid storage in both microalgae (Minhas et al., 2016) and fungi (Papanikolaou et al., 2007). Induction of lipid storage in oleaginous microbes is often associated with the nutrient-starving strategy, which is detrimental to the growth as well as total lipid yield. Hence, instead of impeding starch synthesis in microalgae, it could be more useful to divert photosynthetic carbon flux en route for lipid accumulation than starch storage (Griffiths and Harrison, 2009). The co-cultures of C. minutissima MCC 27/A. awamori and C. minutissima UTEX 2219/ A. awamori co-cultures were grown mixotrophically owing to the several advantages of mixotrophy over heterotrophy. It benefits not only by facilitating the utilization of light but also through greater integration of organic carbon to cells ensuing increased growth rate, productivity and lipid manufacture (Li et al., 2007; Richmond, 1986). Mixotrophy has minimal energy dissipation and hence has high energetic efficiency (Lalucat et al., 1984). Reports also suggest mixotrophy resulted in decreased CO2 production in comparison to heterotrophic cultures. In mixotrophic outdoor conditions, light energy has been seen to play an important role in glucose metabolism or photosynthetic CO2-refixation in Chlorella (Lee et al., 1996). On the other hand, fungus can utilize a broad range of organic carbon supplemented in the growth medium. Glucose was found to be the best carbon source for C. minutissima MCC 27 followed by glycerol while C. minutissima UTEX 2219 showed comparable growth for 14
both glucose and glycerol. Choi and Yu (2015) found 39.42%, 60.00% and 57.82% higher biomass yield in Chlorella vulgaris, Scenedesmus sp. and Botryococcus braunii, respectively under glycerol-fed mixotrophic conditions in contrast to autotrophic conditions. A 3.5-fold higher lipid productivity was reported in Chlorella vulgaris when supplemented with glycerol than the control medium (Sharma et al., 2016). A. awamori showed the best growth in glycerol followed by glucose. Jia et al. (2009) reported that glycerol can induce morphological differentiation in fungi, Aspergillus terreus, which not only improved the biomass yield, but also found favorable for the secondary metabolite like lipid accumulation as compared to glucose. Oleaginous fungus, Yarrowia lipolytica, showed comparably more glycerol consumption than dextrose and 31% TAGs yield was obtained when supplemented with glycerol in the medium (Sestric et al., 2014). Glycerol, as a carbon source, resulted in the highest biomass accumulation (3.2 ± 0.12 g L-1) followed by glucose (2.8 ± 0.05 g L-1) in the C. minutissima MCC 27/A. awamori co-culture system (Fig. 3). Comparable growth was obtained for both glucose and glycerol in C. minutissima UTEX 2219/A. awamori co-culture system. The increase in total lipid yield and lipid content (% dcw) was observed in both the co-culture systems with glycerol as carbon source. The co-culture C. minutissima MCC 27/A. awamori resulted in a 3.4 times higher lipid yield against axenic C. minutissima MCC 27 while 5.1 times more lipid yield than axenic A. awamori. Similarly, a relatively 3.7 and 4.4 fold rise of lipid yield was observed against axenic C. minutissima UTEX 2219 and A. awamori in C. minutissima UTEX 2219/A. awamori co-culture. Cheirsilp et al. (2012) reported 5.7 times increase of biomass and 3.8 times increase of lipid yield in the co-culture of R. glutinis and C. vulgaris when supplemented with 3% pure glycerol and urea as carbon and nitrogen source. While looking 15
at the mechanism of glycerol uptake by the phyco-myco co-cultivation system, it has been seen that glycerol can be conveniently obtained by the microbes in its reduced form. Further, it can be easily metabolized by the microbes that convert it to a cascade of metabolites having similar productivity to that of any other sugars as substrates (Sharma et al., 2016). Glycerol can enter into cells through simple diffusion without expending any energy. Glycerol, before entering into glycolysis for complete oxidation, undergoes two consecutive enzymatic reactions forming glycerol α-phosphate and dihydroxyacetone phosphate, respectively. These two reactions exclusively produce one molecule of NADPH, which does not exist in general sugar metabolism (Jia et al., 2009). The cost of the carbon source is crucial in microbial culture. It was found to have accounted for 50% of the expenditure of the medium during microalgae culture (Cheng et al., 2009). The cost of glucose is higher compared to other nutrients required for the cultivation. Hence, glycerol could be a promising alternative economic carbon source for the cultivation of the co-culture. There are many substitutes for the pure glycerol that could further reduce the production cost. For example, glycerol produced as a byproduct of biodiesel production process and soap industries, which are of trivial commercial value (Tan et al., 2013) could be used for this process. However, these are in crude forms and require further research for its applicability for the cultivation of algae-fungus consortia. Nitrogen was identified as one of the most important nutrient having pronounced effects on the biomass and lipid accumulation of many microalgae (Griffiths and Harrison, 2009). Oleaginous microbes can utilize a variety of nitrogen sources. It is also a limiting nutrient and hence can inhibit the growth of cells depending on the species and cultivation environment. The effect of various nitrogen sources on the growth and lipid accumulation 16
of co-culture has also been investigated to identify the suitable nitrogen source for the coculture and compared with the axenic algal and fungal cultures. Potassium nitrate and yeast extract yielded comparable biomass in microalgae and fungus monocultures whereas regarding lipid content (% dcw), potassium nitrate was found to be more suitable for C. minutissima and yeast extract for A. awamori monocultures. Xiong et al. (2008) found yeast extract as the most suitable nitrogen source for high biomass and lipid accumulation in Chlorella. In C. minutissima MCC 27/A. awamori co-culture, yeast extract resulted in maximum biomass accumulation (3.3 ± 0.13 g L-1) followed by potassium nitrate (3.1 ± 0.05 g L-1) (Fig. 4). On the other hand, comparable results were obtained for yeast extract (3.0 ± 0.21 g L-1) and potassium nitrate (2.94 ± 0.05 g L-1) in C. minutissima UTEX 2219/A. awamori co-culture. In the presence of potassium nitrate, lipid concentrations were found to be the maximum in both the co-cultures. A significant increase of total lipid yield was observed in both the co-culture systems as compared to the respective monocultures. Total lipid yield in C. minutissima MCC 27/A. awamori co-culture systems was found to be 3.0 (C. minutissima MCC 27/A. awamori) and 3.3 (C. minutissima UTEX 2219/A. awamori) times more than the corresponding axenic algal cultures. Yeast extract is an organic nitrogen source, and in comparison to the other two inorganic nitrogen sources, it is overpriced. Hence, potassium nitrate can also be a potential and economic nitrogen source for the cultivation and lipid accumulation of algae-fungus co-cultures. 3.3. Analysis of fatty acid methyl esters of the oil produced from the co-culture When fatty acid composition of the enzymatically transesterified axenic microlgae and fungal oil as well as the co-culture oils was analyzed, the C16-C18 fatty acids were found to be predominating the profile (Table 1). In axenic C. minutissima MCC 27 and C. 17
minutissima UTEX 2219, palmitic acid, methyl esters (C16:0) were observed to be the prinipal fatty acid. Several researchers reported palmitic acid as the major fatty acid constituent in Chlorella spp. (Santhoshkumar et al., 2015; Yeh and Chang, 2012). On the other hand, in axenic A. awamori biodiesel, oleic acid, methyl ester (C18:1) was the primary fatty acid constituent. Several researchers have reported oleic acid as the dominant fatty acid in many fungal and yeast species (Li et al. 2007). Wrede et al. (2014) and El-batal et al. (2016) reported 30% and 46.5% of oleic acid in Aspergillus spp. However, in both co-culture systems, palmitic acid, methyl esters (C16:0) were found to be the principal component of the FAME composite followed by the oleic acid, methyl esters (C18:1). Microalgae appeared to be the principal contributor of the palmitic acid (C16), while both algae and fungus seem to be equally contributing oleic acid (C17) in both C. minutissima MCC 27/ A. Awamori and C. minutissima UTEX 2219/ A. Awamori consortia. Both strains of C. minutissima were found to have solely contributed C16:1 and C18:3 fatty acids in the co-culture oils as these fatty acids were absent in the unifungal oil fraction. On the other hand, A. awamori was found to have contributed more towards C18:0 and C18:2 fatty acid pool in the co-cultures. In this study, co-culture oils show similarities to plant oils with C16:0 and C18:1 as the principal fatty acid, methyl ester composites of the co-culture oils (Gui et al., 2008). This in turn suggests the co-culture used in this study could be employed as a potential biodiesel feedstock. However, the co-culture oils were more advantageous being more saturated than several plant oils such as soybean oil, jatropha oil, rapeseed oil and A. awamori oil (this study). It could help in enhancing several crucial combustion related fuel properties such as cetane number (CN), oxidative stability and reduced NOx emissions, etc 18
(Knothe, 2011). Conversely, the retainment of unsaturated fatty acids in the co-culture oils could aid in ameliorating the cold temperature performance and kinematic viscosity (Knothe, 2011) of the microbial oil.
4. Conclusions Enhanced biomass and lipid production at an early stage of growth compared to the monocultures were obtained in both the co-cultures. Co-cultures showed growth and lipid accumulation on alternative carbon and nitrogen sources. Glycerol and potassium nitrate respectively were found to be the most suitable carbon and nitrogen sources for the coculture. The fatty acid composition of the lipid produced from co-cultures was found to be similar to that of microalgal components of the co-culture; however, both algal and fungal counterparts made contributions to the co-culture FAME profiles, which made it pertinent for biodiesel production.
Acknowledgement The authors would like to thank Indian Institute of Technology Kharagpur India for providing financial assistance to carry out the research work.
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Figure captions Fig. 1. (A) Biomass and lipid accumulation profile of C. minutissima MCC 27 and C. minutissima UTEX 2219. (B) Comparision of growth of C. minutissima MCC 27 and C. minutissima UTEX 2219 under autotrophy (control), mixotrophy, and co-culture with A. awamori in N11 medium supplemented with 1% glucose. All values are mean ± SE, n = 3. Data analyzed by one-way ANOVA with post-hoc t-tests. p<0.05 was considered statistically significant.* P<0.05, ** P<0.01, *** P<0.001. Fig. 2. Variation of biomass production and lipid accumulation with respect to different inoculum composition of co-culture C. minutissima MCC 27/A. awamori (A) and C. minutissima UTEX 2219/ A. awamori (B). All values are mean ± SE, n = 3. Data analyzed by one-way ANOVA with post-hoc t-tests. p<0.05 was considered statistically significant.* P<0.05, ** P<0.01, *** P<0.001. Fig. 3. Effect of different carbon sources on Biomass concentration (g L-1; A, B), Lipid concentration (mg L-1 ; C, D) and Lipid content (% dcw; E, F) of axenic cultures of C. minutissima MCC 27, C.minutissima UTEX 2219, A. awamori, and their co-cultures. Carbon content per liter of growth medium was kept constant. Nitrogen source was 1gL-1 potassium nitrate. All values are mean ± SE, n = 3. Data analyzed by one-way ANOVA with post-hoc t-tests. p<0.05 was considered statistically significant.* P<0.05, ** P<0.01, *** P<0.001. Fig. 4. Effect of different nitrogen sources on Biomass concentration (g L-1; A, B), Lipid concentration (mg L-1 ; C, D) and Lipid content (% dcw; E, F) of axenic cultures of C. minutissima MCC 27, C. minutissima UTEX 2219, A. awamori, and their co-cultures.
26
Nitrogen content per liter of growth medium was kept constant. Carbon source was 1% glycerol. All values are mean ± SE, n = 3. Data analyzed by one-way ANOVA with posthoc t-tests. p<0.05 was considered statistically significant.* P<0.05, ** P<0.01, *** P<0.001.
27
Figures
***
B
A
*** *** *** *** **
Fig. 1.
28
A
**
* ***
A
B B **
* *
** ***
** ***
Fig. 2.
29
*
**
* ***
*** *
**
* * **
**
***
**
*
*
**
**
*
** *
*
* **
** **
** *
*
Fig. 3.
30
*** ***
**
*** A
***
**
**
** **
**
B
**
*
*
*
**
C
**
D
**
**
*
** ***
**
**
**
**
**
*
* E
Fig. 4.
31
F
Tables
Table 1. Distribution of major fatty acids in monocultutre and co-culture oils
C. minutissima C. minutissima
Co-culture (C.
Co-culture (C.
minutissima
minutissima
MCC 27/ A.
UTEX 2219/ A.
awamori)
awamori)
35.02
31.26
4.73
5.11
A. awamori
FAME (%) MCC 27
UTEX 2219
Palmitic (C16:0)
40.50
38.20
7.50
Palmitoleic (C16:1)
8.25
12.43
Stearic (C18:0)
1.70
3.41
5.20
2.11
4.80
Oleic (C18:1)
20.32
17.52
37.00
24.21
21.14
Linoleic (C18:2)
7.89
2.53
10.21
7.80
4.32
Linolenic (C18:3)
10.31
10.48
4.77
6.26
32
Highlights
•
Co-culture of microalgae and fungus were investigated for biodiesel production.
•
Co-culture fostered synergistic effects on the cumulative biomass and lipid yields.
•
Both algae and fungus contributed to the FAME profile of the co-culture oil.
33
S0960-8524(17)30529-1 http://dx.doi.org/10.1016/j.biortech.2017.04.039 BITE 17934
To appear in:
Bioresource Technology
Received Date: Revised Date: Accepted Date:
27 January 2017 8 April 2017 10 April 2017
Please cite this article as: Dash, A., Banerjee, R., Enhanced biodiesel production through phyco-myco co-cultivation of Chlorella minutissima and Aspergillus awamori: an integrated approach, Bioresource Technology (2017), doi: http://dx.doi.org/10.1016/j.biortech.2017.04.039
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Enhanced biodiesel production through phyco-myco co-cultivation of Chlorella minutissima and Aspergillus awamori: an integrated approach Archana Dash and Rintu Banerjee* Agricultural and Food Engineering Department, Indian Institute of Technology Kharagpur, Kharagpur-721302, India
* Address of corresponding author Prof Rintu Banerjee Department of Agricultural and Food Engineering, Indian Institute of Technology Kharagpur- 721302, India Tel: +91-3222-283104(O); Fax: +91-3222- 282244 E-mail: [email protected]
1
Abstract Algae-fungus co-culture was investigated as an alternative biodiesel feedstock. An oleaginous filamentous fungus Aspergillus awamori was co-cultured with Chlorella minutissima MCC 27 and Chlorella minutissima UTEX 2219, respectively in N11 medium furnished with different carbon and nitrogen sources. The biomass and lipid production potential of the two C. minutissima–A. awamori co-cultures was compared against the monocultures. A substantial enhancement in biomass and lipid accumulation was observed in both the co-cultures. When supplemented with different carbon and nitrogen sources, glycerol and potassium nitrate were found to be the most effective. In the presence of glycerol, a 2.6-3.9 fold increase of biomass and 3.4-5.1 fold increase of total lipid yields were observed in the co-cultures as compared to the axenic monocultures. Furthermore, C16:0 (31.26-35.02%) and C18:1 (21.14-24.21%) fatty acids were the major composites of the co-culture oils, which suggest co-culture as a promising strategy for biodiesel production.
Keywords: Biodiesel; Chlorella minutissima; Aspergillus awamori; Co-culture; FAME
2
1. Introduction Green energy, an alternative to rapidly-decreasing fossil fuels, has received a lot of attention and popularity in recent times. It is clean, renewable, and sustainable which can provide a robust, massive, and enduring energy supply. In this context, microalgae share many explicit advantages as a third generation biofuel. The subdued food vs fuel issue, round-the-year farming, higher biomass turnover per unit area with high photon-to-biomass conversion, environment-friendly exhaust emission profile, CO2 bio-mitigation through photosynthetic-fixation, substantial TAG/lipid storage, independence of the water quality for cultivation, and valorization of defatted biomass by the production of high-value, lowvolume products have made microalgae more advantageous than their oleaginous terrestrial competitors (Sayre, 2010; Sharma et al., 2012; Rai et al., 2015). Despite these positive accounts, there are several serious constraints which are a hindrance in the algae-to-diesel reality. First and foremost, one of the major issues is the slow-growing nature of the oleaginous species of the microalgae (Sharma et al., 2012) and their mutually incongruous nature of biomass and lipid accumulation pattern (Li et al., 2011). In most of the microalgae, high lipid contents are usually at the expense of biomass reserve of the cells under nutrient-deprived conditions (Zhu et al., 2016). Consequently, there is often a significant loss of total lipid yields which is the immediate substrate for transesterification i.e. biodiesel production. Besides, issues such as high-cost harvesting techniques, poorly sustainable and renewable nutrient supply, inefficient and costly lipid extraction techniques coupled with poor lipid content and fatty acid (FA) composition (Molina Grima et al., 2003) are yet to be properly addressed.
3
The technical challenges faced in harvesting microalgae are due to the tiny cell size (2-20 µm), negative charge (−7.5 to −40 mV) on its surface, low density (0.3–5 g L-1), and colloidal stability in suspension (Liu et al., 2013; Li et al., 2008). Due to the negative charge on its cell surface, oleaginous microalgae overcome the force of natural gravity and remain suspended in the aqueous medium. The dewatering of microalgal biomass often involves one or more solid-liquid separation stages, which account for 20-30% of the gross cultivation cost of microalgae (Molina Grima et al., 2003; Zhou et al., 2015). The two widely practiced techniques for large-scale harvesting of microalgae by the industries are filtration and centrifugation. Centrifugation was found to be a cost-intensive process for the production of a high-volume, low-value product like biodiesel. On the other hand, conventional filtration method is a simpler and cheaper technique. However, for harvesting extremely small-dimensioned microalgae such as C. minutissima and Nannochloropsis oculata, this method alone cannot be effective and efficient. It may lead to membrane clogging, fouling and frequent replacement of the filter causing a many-fold increase in the overall cost of the process obviating its large-scale application (Singh et al., 2013). In this regard, filamentous fungi can be a possible solution because they can be utilized for bioflocculating microalgal biomass (Zhou et al., 2015) and subsequently easily separated through conventional filtration method. The fungal-assisted bio-flocculation technique requires less energy input and being free of chemical contaminants facilitates proper downstream processing of the harvested biomass and recycling of the spent medium (Li et al., 2008; Zhou et al., 2015). It shows immense potential for harvesting the microalgal cells as a relatively easy and cheaper technology compared to other conventional harvesting techniques. Besides these advantages, another important aspect of the co-culture is the 4
mutual behavior of the microalgae and filamentous fungus, which could be of great advantage. Obligate mutualism has been seen between a few photobionts and fungi in nature (Wrede et al., 2014). However, under synthetic co-culture conditions, it would be interesting to see their corresponding growth and lipid accumulation behavior. The consortia diversity could induce significant changes in the nutrient flux affecting the growth and subsequent modulation of the metabolic fluxes can bring about shifts in lipid metabolism of the oleaginous entities during their co-culture. In this study, two oleaginous alga-fungus consortia were investigated for biodiesel production. The consortia employed in this study were the amalgamation of two different strains of green microalgae, C. minutissima (MCC 27 and UTEX 2219) and a filamentous fungus, A. awamori. These two microbial entities of the co-culture systems are from widely different phylogenetic origins and have multiple nutrient-selectivity. For example, green algae are photoautotrophs while fungi are obligate heterotrophs. When these microorganisms are grown in a single system under mixotrophic nutritive mode, relative selectivity of the nutrients and the trophic modes for both may vary. Hence, the selectivity and suitability of the different carbon and nitrogen sources for the co-cultures were investigated. Also, the growth of each component can be affected by each other’s growth tendencies under defined conditions. These changes can bring about significant difference in the cumulative biomass and lipid yield of the consortia. Moreover, the interaction between the algae and filamentous fungus may also modulate the fatty acid composition in synergy, and subsequently can affect the biodiesel properties as well. Hence, the subsequent effect on the collective biomass and lipid yields by the co-culture systems and the changes in the FAME profiles were subjected to investigation in the presented study. 5
2. Materials and methods 2.1. Microorganisms, inoculum preparation, and culture conditions C. minutissima MCC 27 and C. minutissima UTEX 2219 were procured from the Center for Conservation and Utilization of Blue Green Algae (CCUBGA), IARI, New Delhi, India and Culture collection of Algae, University of Texas, Austin, respectively. A. awamori was previously isolated from the native soil of Kharagpur, West Bengal, India. The axenic cultures of microalgae were maintained in N11 medium (Saxena et al., 2016) under 6.8 pH (pH meter, M/s. ELICO, India), 25 ± 2 °C, 75 µmol photon m-2 s-1 PAR light intensity and 14:10 h (L:D) photoperiod. The algal cells were harvested using a centriguage (eppendorf centrifuge 5810R, 5000 rpm, 15 min) and subsequently washed twice with distilled water. Further, the cell counting was carried out using a hemocytometer (FeinOptik, Bad Blankenburg, Germany) and maintained by dilution with distilled water. A. awamori was maintained on Potato Dextrose Agar (PDA) at pH 5 and 30 ± 1 °C. Fungal spores were extracted from the 6-day-old culture from the petri dishes with 10 mL distilled water. After washing twice with distilled water, the fungal spores were counted under the microscope. The initial spore count was maintained by diluting the resultant spore solution with distilled water. Equimolar concentrations of different carbon sources viz. glucose, glycerol, and acetate and nitrogen sources such as potassium nitrate, urea, and yeast extract were supplemented in the growth medium. Incubation temperature (25 °-30 °C) and initial pH (5-8) were varied to investigate for maximum biomass and lipid accumulation. Supply of exogenous air or sparging of CO2 was avoided in this study. Only manual gentle shaking
6
for 10 minutes was done thrice a day to prevent sticking of the cells with the wall and for uniform growth. 2.2. Growth study The growth of C. minutissima MCC 27 and UTEX 2219 in monocultures and cocultures was measured spectrophotometrically (UV-1800, SHIMADZU, Japan) at 540 nm. The dry cell weight of monocultures of both C. minutissima MCC 27 and A. awamori as well as the co-cultures was determined gravimetrically. The entire biomass was harvested through centrifugation and oven-dried (60 °C) in a pre-weighed glass vial. The biomass was dried till it achieved a constant weight and finally the dry cell weight was determined from the difference between the final and empty vial weight. Biomass of microalgae in the co-cultures was determined indirectly by measuring the Chla concentration in the co-culture and subsequently the biomass was determined from the standard curve of Chla concentration and dry cell weight of the microalgae. For chlorophyll extraction, chilled methanol (99.8 %) was used followed by a sonication bath for 15 minutes and kept overnight. Chla concentration was measured spectrophotometrically at 652 and 665 nm (Porra et al., 1989). The Chla and biomass of microalgae were correlated as per the following equation: Microalgae dry biomass (mg) =22.944 × A652 [R2 = 0.99] (equation 1) After obtaining the biomass of microalgae from the equation 1, biomass of A. awamori in the co-culture was determined from the difference of total biomass and biomass of microalgae. 2.3. Lipid extraction
7
Lipid was extracted from the oven-dried microbial biomass by following the Bligh and Dyer (1959) protocol. After the extraction, lipid was filtered via filter paper (Whatman No. 1) and stored in a pre-weighed vial (W1). The solvent was evaporated in a water bath followed by an oven-drying (104 °C) till the constant weight was recorded consecutively for three times. The final weight of the vial was measured (W2). The amount of the lipid was estimated from the difference of W1 and W2, and expressed as % dcw as per the following formula:
Lipid content (% dcw) =
Mass of lipid Mass of biomass
× 100
2.4. Biodiesel production 2.4.1. Lipase production, extraction, and assay Lipase was produced indigenously from Rhizopus oryzae NRRL 3562 under solid state fermentation (SSF). Czapek-dox medium (Sarkar et al., 1998) was used in 1:1 ratio to moisturize the substrate (wheat bran) which was used as the major carbon source. The fermentation medium was further fortified with coconut oil (Garlapati and Banerjee, 2010). After inoculation with R. oryzae, the medium was incubated for the completion of the required fermentation, thereafter extracellular lipase was extracted for subsequent studies. The crude enzyme was assayed for lipase activity, which was determined spectrophotometrically by using p-nitrophenylpalmitate (pNPP) as substrate (Pencreac’h and Baratti, 2001). Total protein content was determined through modified Lowry method by using bovine serum albumin (BSA) as standard (Lowry et al., 1951). 2.4.2. Enzymatic transesterification
8
The indigenously produced lipase was processed for transesterification studies. Lipasemediated transesterification was carried out by taking oil to methanol molar ratio 1:3 at reaction conditions of 37 °C, 200 rpm and 24 h. 10 U immobilized lipase g-1 of oil was used for the reaction. Immobilization of lipase on activated celite (50 U g-1) and preparation of sample for GCMS analysis were done as per the protocol of Garlapati et al. (2013). 2.4.3. FAME analysis FAME profiling was done through GC-MS analysis (Agilent 6890 N gas chromatography paired with Agilent MS-5975 inert XL mass selective detector; Agilent Technologies, Little Fall, NY, USA). HP-5MS (5 % phenyl methylsiloxane) capillary column (Agilent Technologies, Palo Alto, CA, USA) with 30 m х 0.25 mm х 0.25 mm dimension was used. The initial temperature was maintained at 150 °C and raised to 280 °C at the rate of 4 °C min-1, and kept at 280 °C for 9 min. The flow rate was 0.8 mL min-1. The injector and detector temperatures were 250 °C and 260 °C, respectively. For FAME identification, retention times of isolated compounds were matched with that of FAME standards. In addition, by using the National Institute of Standards and Technology (NIST) mass spectral library, mass fragment pattern of each resolved peak was compared.
3. Results and discussion 3.1. Growth and lipid accumulation in monoculture and co-culture The biomass and lipid accumulation have been investigated for a period of 28 days. Maximum biomass was observed on the 21st day of incubation in both the strains. Maximum biomass accumulation was seen in C. minutissima MCC 27 (1.140 ± 0.087 g L-1) followed by C. minutissima UTEX 2219 (0.900 ± 0.050 g L-1). Substantial lipid 9
accumulation was observed in the late exponential phase of their growth. C. minutissima MCC 27 and C. minutissima UTEX 2219 were found to accumulate 22.12 ± 0.99 % and 20.00 ± 1.00 % dcw lipid, respectively. C. minutissima MCC 27 was observed to be better biomass producer and lipid accumulator as compared to C. minutissima UTEX 2219. Fig. 1A depicts the growth and lipid production profile of the two different microalgae monocultures i.e. C. minutissima MCC 27 and C. minutissima UTEX 2219. The growth of C. minutissima MCC 27 as well as UTEX 2219 was monitored for 9 days under autotrophic, mixotrophic (1% glucose) and co-culture mode (1% glucose) and compared. Interestingly, an approximately 12.4-fold rise in the growth of C. minutissima MCC 27 in co-culture system was observed as compared to autotrophic axenic culture, while it was 1.8-fold more than the mixotrophically grown axenic test algae on the 6th day of incubation (Fig. 1B). Similar trends were also obtained in case of C. minutissima UTEX 2219/A. awamori co-culture. Increase in the biomass accumulation of C. minutissima UTEX 2219 in the co-culture with A. awamori were up to approximately 13.9- and 2.8-fold as compared to autotrophic and mixotrophic axenic C. minutissima UTEX 2219, respectively (Fig. 1B). The significant increase in the growth of test microalgae and significant biomass accumulation in presence of A. awamori indicate that the diversity of the consortia selected for this study may have turned out to be the triggering factor for the improved growth of these species. The relative contribution towards the productivity in the co-culture systems between the two species were comparable. Biomass and lipid yields in the co-cultures were cumulatively as well as individually more than the monocultures. Consortia diversity in cocultures can function as an efficient system, which can help in better utilization of the 10
resources. When the utilization of resources is complementary, ecologically different species co-exist (Gause, 1936; Weis et al., 2008). This way co-cultures can harness a greater fraction of the limited resources compared to the monoculture of the most productive species and result in enhanced biomass of the latter (Weis et al., 2008). Furthermore, when diverse species have to compete for the limiting resources, they complement each other in the utilization of their resources (Loreau et al., 2001; Korhonen et al., 2011) which can turn out to have a positive effect on the biomass accumulation in the co-culture system. One such example of complementary behavior and better resource utilization in presence of co-culture is reported by Xue et al. (2010). According to their report, incorporation of Spirulina platensis to the yeast culture of Rhodotorula glutinis led to significant enhancement of O2 in the co-culture. Inoculum composition was varied by changing the ratio between fungal spores and algal cells. The inoculum cells count ranged from 1:110 to 1:300 (fungal spores: algal cells). An increasing trend in biomass of co-culture was observed with an increase in the ratio of spore:algal cells on the 6th day of incubation (Fig. 2). Maximum biomass yields were found to be 2.80 ± 0.1 g L-1 and 2.54 ± 0.04 g L-1 for C. minutissima MCC 27 and C. minutissima UTEX 2219 respectively for 1:300. The increase of algal cell numbers in the inoculum composition was observed to have a prominent effect on the total lipid yield (mg L-1) and content (% dcw). Within the range 1:215 to 1:300, a minor difference of lipid content was observed in case of the C. minutissima MCC 27/A. awamori co-culture, whereas it was found comparable in case of the C. minutissima UTEX 2219/A. awamori co-culture in the range 1:205 to 1:300. However, overall maximum total lipid yield was observed with 1:300, which was 510.4 mg L-1 and 437.13 mg L-1 for C. minutissima MCC 11
27 and C. minutissima UTEX 2219, respectively. Similarly, the effect of incubation temperature on the co-cultures of C. minutissima MCC 27/A. awamori and C. minutissima UTEX 2219/A. awamori has been investigated. It was found that 25 °C was observed to be optimum for the growth of test algae while 30 °C was most suitable for A. awamori. However, in case of co-cultures, 25 °C was found to be the most suitable for biomass production of the co-cultures. A minor decrease in biomass was observed in the case of C. minutissima UTEX 2219/A. awamori co-culture when the incubation temperature increased from 25 ° to 30 °C. Overall the biomass production and lipid accumulation patterns were comparable for C. minutissima MCC 27/A. awamori co-cultures, while a slight decrease in biomass production and increase in lipid content (% dcw) were observed with C. minutissima UTEX 2219/A. awamori co-culture. However, the overall effects of incubation temperature were insignificant for both the co-cultivation systems suggesting tolerance of both the co-cultures within the range (25°-30 °C). When the effect of initial pH on the growth and lipid accumulation potential of the co-cultures were investigated, it was seen that pH 5 was found to be the most suitable followed by pH 8 for the consortial biomass yield for both the co-culture systems while pH 5 was found to yield maximum lipid contents. Optimum pH for the growth of C. minutissima was found to be 7.0; on the contrary, pH 5.0 was found to be the most suitable for the growth of A. awamori. The pH 5.0, 5.5 and 8.0 showed comparable biomass yield in both the co-cultures. It suggests that slightly acidic and slightly alkaline growth medium is best suited for the growth of both the cultures. Many Ascomycota fungi were known to secret different organic acids such as acetic acid, citric acid, lactic acid, oxalic acid, malic acid, etc., into the growth medium (Liaud et al., 2014). These organic acids severely affect the growth of the liable fungus by 12
lowering the cytosolic pH and malfunctioning the permeability of the cell membrane causing its death (Hassan et al., 2015). Yeast releases CO2 as a byproduct of its respiratory metabolism, and it readily dissolves in the aqueous growth medium. As a result, bicarbonate (HCO3 ̶ ) is produced under pH ≥ 7 (Sayre, 2010), which causes a further decline in pH. Microalgae can biologically capture CO2 by concentrating HCO3 ̶ intracellularly, later again get converted into CO2 spontaneously or by carbonic anhydrase (Moroney and Somanchi, 1999). The intracellular CO2 is then fixed by the algal photosynthetic machinery to be reciprocated into algal biomass. The OH ̶ produced in this process helps in increasing the pH towards alkalinity (Richmond, 1986). Microalgae can metabolize different organic acids such as acetic acid, malic acid, pyruvic acid, lactic acids (fungal metabolites) for its growth (Bollman and Robinson, 1977). Hence, the co-culture of fungus with microalgae could have been helpful in assuaging the inhibitory effects of organic acids on the growth of the fungus. On the other hand, the optimum pH for the growth of C. minutissima was found to be 7.0. Rai et al. (2015) also reported maximum growth at pH 7.0 while Sankar et al. (2011) reported optimum pH for the growth of C. minutissima as 6.0. During microalgae cultivation, pH tends to increase and becomes alkaline as a result of OH ̶ production during CO2 assimilation (Bi and Zhou, 2016). Thus, microalgae, in co-culture, could help in buffering of the medium for mutual growth of both the co-culturants. 3.2. Selection of suitable carbon and nitrogen sources for co-culture In this study, an attempt has been made to investigate the effect of various carbon sources on biomass and lipid accumulation of co-cultures as well as monocultures of microalgae and fungus. The type of carbon source employed for the growth of oleaginous 13
species may play a crucial role in steering the flux of carbon towards lipid. In microalgae, intracellular accumulation of neutral lipids occurs from the bioconversion of starch/carbon to lipids through different mechanisms depending on the strains (Li et al., 2011). Application of different carbon sources can amend the core carbon metabolism by driving the carbon-partitioning towards the lipid storage in both microalgae (Minhas et al., 2016) and fungi (Papanikolaou et al., 2007). Induction of lipid storage in oleaginous microbes is often associated with the nutrient-starving strategy, which is detrimental to the growth as well as total lipid yield. Hence, instead of impeding starch synthesis in microalgae, it could be more useful to divert photosynthetic carbon flux en route for lipid accumulation than starch storage (Griffiths and Harrison, 2009). The co-cultures of C. minutissima MCC 27/A. awamori and C. minutissima UTEX 2219/ A. awamori co-cultures were grown mixotrophically owing to the several advantages of mixotrophy over heterotrophy. It benefits not only by facilitating the utilization of light but also through greater integration of organic carbon to cells ensuing increased growth rate, productivity and lipid manufacture (Li et al., 2007; Richmond, 1986). Mixotrophy has minimal energy dissipation and hence has high energetic efficiency (Lalucat et al., 1984). Reports also suggest mixotrophy resulted in decreased CO2 production in comparison to heterotrophic cultures. In mixotrophic outdoor conditions, light energy has been seen to play an important role in glucose metabolism or photosynthetic CO2-refixation in Chlorella (Lee et al., 1996). On the other hand, fungus can utilize a broad range of organic carbon supplemented in the growth medium. Glucose was found to be the best carbon source for C. minutissima MCC 27 followed by glycerol while C. minutissima UTEX 2219 showed comparable growth for 14
both glucose and glycerol. Choi and Yu (2015) found 39.42%, 60.00% and 57.82% higher biomass yield in Chlorella vulgaris, Scenedesmus sp. and Botryococcus braunii, respectively under glycerol-fed mixotrophic conditions in contrast to autotrophic conditions. A 3.5-fold higher lipid productivity was reported in Chlorella vulgaris when supplemented with glycerol than the control medium (Sharma et al., 2016). A. awamori showed the best growth in glycerol followed by glucose. Jia et al. (2009) reported that glycerol can induce morphological differentiation in fungi, Aspergillus terreus, which not only improved the biomass yield, but also found favorable for the secondary metabolite like lipid accumulation as compared to glucose. Oleaginous fungus, Yarrowia lipolytica, showed comparably more glycerol consumption than dextrose and 31% TAGs yield was obtained when supplemented with glycerol in the medium (Sestric et al., 2014). Glycerol, as a carbon source, resulted in the highest biomass accumulation (3.2 ± 0.12 g L-1) followed by glucose (2.8 ± 0.05 g L-1) in the C. minutissima MCC 27/A. awamori co-culture system (Fig. 3). Comparable growth was obtained for both glucose and glycerol in C. minutissima UTEX 2219/A. awamori co-culture system. The increase in total lipid yield and lipid content (% dcw) was observed in both the co-culture systems with glycerol as carbon source. The co-culture C. minutissima MCC 27/A. awamori resulted in a 3.4 times higher lipid yield against axenic C. minutissima MCC 27 while 5.1 times more lipid yield than axenic A. awamori. Similarly, a relatively 3.7 and 4.4 fold rise of lipid yield was observed against axenic C. minutissima UTEX 2219 and A. awamori in C. minutissima UTEX 2219/A. awamori co-culture. Cheirsilp et al. (2012) reported 5.7 times increase of biomass and 3.8 times increase of lipid yield in the co-culture of R. glutinis and C. vulgaris when supplemented with 3% pure glycerol and urea as carbon and nitrogen source. While looking 15
at the mechanism of glycerol uptake by the phyco-myco co-cultivation system, it has been seen that glycerol can be conveniently obtained by the microbes in its reduced form. Further, it can be easily metabolized by the microbes that convert it to a cascade of metabolites having similar productivity to that of any other sugars as substrates (Sharma et al., 2016). Glycerol can enter into cells through simple diffusion without expending any energy. Glycerol, before entering into glycolysis for complete oxidation, undergoes two consecutive enzymatic reactions forming glycerol α-phosphate and dihydroxyacetone phosphate, respectively. These two reactions exclusively produce one molecule of NADPH, which does not exist in general sugar metabolism (Jia et al., 2009). The cost of the carbon source is crucial in microbial culture. It was found to have accounted for 50% of the expenditure of the medium during microalgae culture (Cheng et al., 2009). The cost of glucose is higher compared to other nutrients required for the cultivation. Hence, glycerol could be a promising alternative economic carbon source for the cultivation of the co-culture. There are many substitutes for the pure glycerol that could further reduce the production cost. For example, glycerol produced as a byproduct of biodiesel production process and soap industries, which are of trivial commercial value (Tan et al., 2013) could be used for this process. However, these are in crude forms and require further research for its applicability for the cultivation of algae-fungus consortia. Nitrogen was identified as one of the most important nutrient having pronounced effects on the biomass and lipid accumulation of many microalgae (Griffiths and Harrison, 2009). Oleaginous microbes can utilize a variety of nitrogen sources. It is also a limiting nutrient and hence can inhibit the growth of cells depending on the species and cultivation environment. The effect of various nitrogen sources on the growth and lipid accumulation 16
of co-culture has also been investigated to identify the suitable nitrogen source for the coculture and compared with the axenic algal and fungal cultures. Potassium nitrate and yeast extract yielded comparable biomass in microalgae and fungus monocultures whereas regarding lipid content (% dcw), potassium nitrate was found to be more suitable for C. minutissima and yeast extract for A. awamori monocultures. Xiong et al. (2008) found yeast extract as the most suitable nitrogen source for high biomass and lipid accumulation in Chlorella. In C. minutissima MCC 27/A. awamori co-culture, yeast extract resulted in maximum biomass accumulation (3.3 ± 0.13 g L-1) followed by potassium nitrate (3.1 ± 0.05 g L-1) (Fig. 4). On the other hand, comparable results were obtained for yeast extract (3.0 ± 0.21 g L-1) and potassium nitrate (2.94 ± 0.05 g L-1) in C. minutissima UTEX 2219/A. awamori co-culture. In the presence of potassium nitrate, lipid concentrations were found to be the maximum in both the co-cultures. A significant increase of total lipid yield was observed in both the co-culture systems as compared to the respective monocultures. Total lipid yield in C. minutissima MCC 27/A. awamori co-culture systems was found to be 3.0 (C. minutissima MCC 27/A. awamori) and 3.3 (C. minutissima UTEX 2219/A. awamori) times more than the corresponding axenic algal cultures. Yeast extract is an organic nitrogen source, and in comparison to the other two inorganic nitrogen sources, it is overpriced. Hence, potassium nitrate can also be a potential and economic nitrogen source for the cultivation and lipid accumulation of algae-fungus co-cultures. 3.3. Analysis of fatty acid methyl esters of the oil produced from the co-culture When fatty acid composition of the enzymatically transesterified axenic microlgae and fungal oil as well as the co-culture oils was analyzed, the C16-C18 fatty acids were found to be predominating the profile (Table 1). In axenic C. minutissima MCC 27 and C. 17
minutissima UTEX 2219, palmitic acid, methyl esters (C16:0) were observed to be the prinipal fatty acid. Several researchers reported palmitic acid as the major fatty acid constituent in Chlorella spp. (Santhoshkumar et al., 2015; Yeh and Chang, 2012). On the other hand, in axenic A. awamori biodiesel, oleic acid, methyl ester (C18:1) was the primary fatty acid constituent. Several researchers have reported oleic acid as the dominant fatty acid in many fungal and yeast species (Li et al. 2007). Wrede et al. (2014) and El-batal et al. (2016) reported 30% and 46.5% of oleic acid in Aspergillus spp. However, in both co-culture systems, palmitic acid, methyl esters (C16:0) were found to be the principal component of the FAME composite followed by the oleic acid, methyl esters (C18:1). Microalgae appeared to be the principal contributor of the palmitic acid (C16), while both algae and fungus seem to be equally contributing oleic acid (C17) in both C. minutissima MCC 27/ A. Awamori and C. minutissima UTEX 2219/ A. Awamori consortia. Both strains of C. minutissima were found to have solely contributed C16:1 and C18:3 fatty acids in the co-culture oils as these fatty acids were absent in the unifungal oil fraction. On the other hand, A. awamori was found to have contributed more towards C18:0 and C18:2 fatty acid pool in the co-cultures. In this study, co-culture oils show similarities to plant oils with C16:0 and C18:1 as the principal fatty acid, methyl ester composites of the co-culture oils (Gui et al., 2008). This in turn suggests the co-culture used in this study could be employed as a potential biodiesel feedstock. However, the co-culture oils were more advantageous being more saturated than several plant oils such as soybean oil, jatropha oil, rapeseed oil and A. awamori oil (this study). It could help in enhancing several crucial combustion related fuel properties such as cetane number (CN), oxidative stability and reduced NOx emissions, etc 18
(Knothe, 2011). Conversely, the retainment of unsaturated fatty acids in the co-culture oils could aid in ameliorating the cold temperature performance and kinematic viscosity (Knothe, 2011) of the microbial oil.
4. Conclusions Enhanced biomass and lipid production at an early stage of growth compared to the monocultures were obtained in both the co-cultures. Co-cultures showed growth and lipid accumulation on alternative carbon and nitrogen sources. Glycerol and potassium nitrate respectively were found to be the most suitable carbon and nitrogen sources for the coculture. The fatty acid composition of the lipid produced from co-cultures was found to be similar to that of microalgal components of the co-culture; however, both algal and fungal counterparts made contributions to the co-culture FAME profiles, which made it pertinent for biodiesel production.
Acknowledgement The authors would like to thank Indian Institute of Technology Kharagpur India for providing financial assistance to carry out the research work.
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Figure captions Fig. 1. (A) Biomass and lipid accumulation profile of C. minutissima MCC 27 and C. minutissima UTEX 2219. (B) Comparision of growth of C. minutissima MCC 27 and C. minutissima UTEX 2219 under autotrophy (control), mixotrophy, and co-culture with A. awamori in N11 medium supplemented with 1% glucose. All values are mean ± SE, n = 3. Data analyzed by one-way ANOVA with post-hoc t-tests. p<0.05 was considered statistically significant.* P<0.05, ** P<0.01, *** P<0.001. Fig. 2. Variation of biomass production and lipid accumulation with respect to different inoculum composition of co-culture C. minutissima MCC 27/A. awamori (A) and C. minutissima UTEX 2219/ A. awamori (B). All values are mean ± SE, n = 3. Data analyzed by one-way ANOVA with post-hoc t-tests. p<0.05 was considered statistically significant.* P<0.05, ** P<0.01, *** P<0.001. Fig. 3. Effect of different carbon sources on Biomass concentration (g L-1; A, B), Lipid concentration (mg L-1 ; C, D) and Lipid content (% dcw; E, F) of axenic cultures of C. minutissima MCC 27, C.minutissima UTEX 2219, A. awamori, and their co-cultures. Carbon content per liter of growth medium was kept constant. Nitrogen source was 1gL-1 potassium nitrate. All values are mean ± SE, n = 3. Data analyzed by one-way ANOVA with post-hoc t-tests. p<0.05 was considered statistically significant.* P<0.05, ** P<0.01, *** P<0.001. Fig. 4. Effect of different nitrogen sources on Biomass concentration (g L-1; A, B), Lipid concentration (mg L-1 ; C, D) and Lipid content (% dcw; E, F) of axenic cultures of C. minutissima MCC 27, C. minutissima UTEX 2219, A. awamori, and their co-cultures.
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Nitrogen content per liter of growth medium was kept constant. Carbon source was 1% glycerol. All values are mean ± SE, n = 3. Data analyzed by one-way ANOVA with posthoc t-tests. p<0.05 was considered statistically significant.* P<0.05, ** P<0.01, *** P<0.001.
27
Figures
***
B
A
*** *** *** *** **
Fig. 1.
28
A
**
* ***
A
B B **
* *
** ***
** ***
Fig. 2.
29
*
**
* ***
*** *
**
* * **
**
***
**
*
*
**
**
*
** *
*
* **
** **
** *
*
Fig. 3.
30
*** ***
**
*** A
***
**
**
** **
**
B
**
*
*
*
**
C
**
D
**
**
*
** ***
**
**
**
**
**
*
* E
Fig. 4.
31
F
Tables
Table 1. Distribution of major fatty acids in monocultutre and co-culture oils
C. minutissima C. minutissima
Co-culture (C.
Co-culture (C.
minutissima
minutissima
MCC 27/ A.
UTEX 2219/ A.
awamori)
awamori)
35.02
31.26
4.73
5.11
A. awamori
FAME (%) MCC 27
UTEX 2219
Palmitic (C16:0)
40.50
38.20
7.50
Palmitoleic (C16:1)
8.25
12.43
Stearic (C18:0)
1.70
3.41
5.20
2.11
4.80
Oleic (C18:1)
20.32
17.52
37.00
24.21
21.14
Linoleic (C18:2)
7.89
2.53
10.21
7.80
4.32
Linolenic (C18:3)
10.31
10.48
4.77
6.26
32
Highlights
•
Co-culture of microalgae and fungus were investigated for biodiesel production.
•
Co-culture fostered synergistic effects on the cumulative biomass and lipid yields.
•
Both algae and fungus contributed to the FAME profile of the co-culture oil.
33