Green integrated process for mitigation of municipal and industrial liquid and solid waste mixes for enhanced microalgal biomass and lipid synthesis for biodiesel

Abstract

Wastewater management has become one of the critical environmental challenges, as waste effluents from municipalities and industries including livestock farms are released into the environment resulting in the deterioration of water ecology, if left untreated. Through this study, we have developed a microalgae mediated green integrated process for wastewater remediation, while simultaneously producing biomass as potentially sustainable feedstock for biofuels. Chlorococcum sp. was cultivated in different types of wastewater and tested for nutrient remediation efficacy as well as biomass production. The results with respect to nutrient removal capacity in wastewater were encouraging as nitrogen was almost completely remediated, whereas removal percentages for carbon and phosphorous were between 67 and 89%. The results obtained using waste mixtures of wastewater effluents and poultry litter waste indicates substantial gain in microalgal biomass yield, an increase of more than 1.5-fold was observed as opposed to unsupplemented use. The obtained experimental microalgal growth data were fitted with a logistic growth model and found to provide a good fit between predicted values and investigated values. Furthermore, the lipid accumulation was enhanced by 2-fold in waste mixtures when compared to standard medium, and the lipids were mainly composed of C16 and C18 fatty acids, which are considered favourable for good-quality biodiesel. The strategy adopted in this study served the dual purpose of waste management and microalgal biomass production for biofuel application, thus making the entire process cost-effective and green.

---

Introduction

The world is confronted with multiple environmental challenges, such as climate change, energy security, a deteriorating ecosystem, waste disposal, pollution of air, soil and water, etc. These are contributed to by the growing human population and the consequent manifold increase in anthropogenic activities.^1–3 Besides, consumption of water has quadrupled because of many water intensive domestic and industrial applications, which consume freshwater and generate huge amount of wastewater effluents of varying characteristics.³ This generated wastewater not only puts huge pressure on treatment facilities but also poses serious hazards to our ecosystem if left untreated or disposed of improperly. Furthermore, there are fast-growing agro-based industries, particularly livestock farming for meat, milk and egg production. Since 1960, the number of animals has increased by 200% for pork and 280% for poultry, whereas in terms of meat production, one fourth accounts for poultry, largely concentrated in the USA, China and Brazil.⁴ Water contamination due to nutrient pollution by these industrial effluent and solid discharges (manure and litter) is generally caused by the leaching and runoff of minerals into the soil or by direct disposal of waste into waterbodies.^4,5 Wastewater effluents comprise of pollutants (organic and inorganic substances) and toxic metals which are not only harmful to the aquatic eco-system, but also have substantial negative impacts on agricultural development, ecosystem and people's lives.^5,6 Treatment technologies, typically based on chemical and physical processes, can play a role in the wastewater management. However the challenge persists that such technologies are not economical as well as not implementable at a large-scale.⁴ Microalgae have emerged as one of the most promising biological alternative for the removal of pollutants in wastewater.⁷ Studies on algal cultivation using industrial, municipal and agricultural wastewaters have been carried out for simultaneous wastewater remediation and algal biomass production.⁸ Additionally, algal biomass has been considered one of the most promising feedstock for the development of third generation biofuels due to the numerous advantages over terrestrial crops.^1,9 The use of biofuels not only mitigate greenhouse gas (GHG) emissions but can also substitute rapidly depleting fossil fuel sustainably. Moreover, microalgae have a diverse array of applications in various fields from low-value to high-value products, including most researched in the field of biofuels. Nevertheless, the biodiesel feedstock cost accounts for more than 70% of the overall cost of biodiesel production.¹⁰ Likewise, cultivation of microalga incurs high cost to the entire production process mainly due to huge requirement of nutrients and water resources.¹⁰ The utilization of waste discharges for algal biomass production can offset the cost of commercial nutrients or fertilizers.^11,12 The microalgal biomass generated can be utilized as a cheap biomass feedstock for the production of biofuels and biochemicals. The U.S. Department of Energy has recognized the potential synergy of incorporating wastewater treatment with algal biofuel production.¹³ Various microalgae like Scenedesmus acutus, Chlorella zofingiensis, Nannochloropsis salina, etc., have been efficiently used for the treatment of wastewaters and biofuels production.^5,14,15 Microalga, Chlorococcum sp. is one of the versatile species that has been used in various applications including, dairy wastewater treatment,¹⁶ carotenoids production¹⁷ and has emerged as a potential biofuel feedstock for the production of biodiesel¹⁸ and bioethanol.^19,20

However, microalgal cultivation in wastewater effluent suffers from lower biomass yields (0.1–0.5 g L⁻¹). In addition, commercial production of microalgae has not become viable and is still limited by the lower yield and high cost.^9,10 Therefore, development of an economically feasible process for large scale microalgal cultivation for biorefinery and biofuel applications demands the usage of cultivation medium involving low-cost substrates. A possible judicious approach can be the effective use of wastes from industries and municipal discharges as an alternative cultivation medium. It would be an added advantage if microalgal cultivation can achieve dual tasks of generating energy in the form of biodiesel and reducing pollutant load of the used wastewater. In this study, we have compared the cultivation of Chlorococcum sp. in different types of liquid and solid waste discharges such as municipal wastewater, industrial effluents and poultry litter waste (PW). Subsequently, these liquid and solid waste sources were judiciously mixed to examine its effect on enhancing microalgal biomass and lipid yields for biodiesel application. The strategy adopted was to establish a green integrated process for effective use of different waste discharges as an inexpensive and alternative source of water and nutrients for concomitant biofuels production and waste management for sustainable future development.

Materials and methods

Microalgae culture and growth medium

The green microalga culture, Chlorococcum infusionum, used in this study was maintained with frequent sub-culturing into bold basal medium (BBM), pH 6.8–7.²⁰ The experiments were carried out in temperature controlled shaking incubator at 27 ± 2 °C, 120 rpm, equipped with cool white fluorescent lamps, light intensity of 50 μmol m⁻² s⁻¹, with photoperiod of 14 : 10 h L/D (light/dark) cycle.

Collection of waste samples, characterization and experimental procedure

Three types of waste discharges were collected from various point sources such as (1) poultry litter waste (PW) assorted from poultry farms, Kharagpur (solid waste), (2) industrial effluent from a common effluent treatment plant (CETP), Butibori, Nagpur, and (3) municipal effluent from a secondary settling tank (MSS), Titagarh, Kolkata. Solid waste, PW, was powdered and mixed with distilled water as well as with liquid CETP and MSS effluents in various quantities (1–30 g L⁻¹) (overnight at 4 °C). The experiments were carried out by utilizing liquid and solid waste discharges independently, i.e. only CETP, MSS and PW in distilled water and in combination that is PW mixed with CETP and MSS. Pretreatment for all the waste discharges mentioned above were carried out by sedimentation and filtration using Whatman no. 1 filter paper to remove large and non-soluble particulate solids. All the filtrates were then sterilized, separately by autoclaving (121 °C for 20 min) and used as growth media for the cultivation of Chlorococcum sp. The experiments were carried out in 150 mL Erlenmeyer flask with working volume of 50 mL. The inoculums size of 10% of a logarithmically grown Chlorococcum infusionum culture (i.e. 7^th day) was used as seed culture. The culture was grown in a temperature controlled shaking incubator (120 rpm) at 27 ± 2 °C and illuminated using cool white fluorescent lamps at 50 μmol m⁻² s⁻¹ with L/D cycle of 14 : 10 h. The experiments were conducted over a period of 27 days in batch mode and the samples were collected every three days. For final biomass concentration and lipid content measurement, the algal biomass was harvested on 21^st day, as the maximum values were achieved at the end of log phase. The microalga was tested for nutrient removal percentage, biomass growth and lipid content in all wastewater medium. All experiments were performed in triplicates and expressed as mean with standard deviation (SD).

Methodology

Sampling and nutrients analysis

During the batch experiment, the biomass and nutrient concentrations were analyzed for the samples collected every three day intervals over a period of 27 days. The microalga, Chlorococcum sp. grown in BBM under the conditions described above was referred as the control. The waste effluents were characterised with respect to biological oxygen demand (BOD), chemical oxygen demand (COD), and total organic carbon (TOC), PO₄, NH₄, NO₃, NO₂ content and pH following the standard protocols of American Public Health Association (APHA).²¹ The samples collected were centrifuged and the supernatant was then filtered using a 0.22 μm membrane filter (Millipore). The filtrates obtained were appropriately diluted and analyzed for TOC, BOD, COD, PO₄, NH₄, NO₃ and NO₂. The characteristics of untreated wastewater samples were estimated for elemental/ions composition and concentration using ion chromatography (IC) (Dionex, USA) and an inductively coupled plasma mass spectrometer (ICP-MS) (Varian 810 ICP-MS System, California).

The percentage removal (R, %) of nutrients was calculated using the eqn (1), where C₀ and C_x were defined as the mean values of nutrient concentration at initial time t₀ and time t_x, respectively.

R% = 100% × (C₀ − C_x)/C₀ (1)

Biomass and lipid analysis

Biomass concentration (g L⁻¹) as dry cell weight (dcw) of algal cells was estimation from the known volume of harvested cells by centrifugation at 6500 g for 10 min and the pellets were washed twice with deionised water. The cell pellets were then dried at 60 °C for 24 h and the final concentration was noted when the constant weight was observed.²⁰ The specific growth rate (μ, d⁻¹) was calculated from the eqn (2), where X1 and X2 were the concentration of algal biomass at the beginning (t1) and at the end (t2) of the exponential growth phase, respectively.

μ = (ln X2 − ln X1)/(t2 − t1) (2)

The Chlorococcum sp. biomass was harvested by centrifugation (6500 g for 10 min and the pellets were washed twice with deionised water) and dried under vacuum conditions before being analyzed for lipid content. To determine the amount of total lipid, lipid extraction was performed following the modified Folch method.²⁰ The extracted total lipids was then filtered through filter paper and measured gravimetrically and lipid content was expressed as a % of dry weight. The quantified lipid content (%, dcw) corresponds to the neutral lipid fraction extracted with hexane from the total lipid. Thereafter, the lipid was transesterified to fatty acid methyl esters (FAME), and the fatty acids composition was analysed using a GC with FID detector.²⁰ All the experiments were carried out at least in triplicates and the average value with standard deviation were reported.

Determination of growth kinetic parameters

The logistic kinetic model was used to model the growth of the experimental biomass concentration. The model is a substrate-independent equation and can accurately describe the growth profile in the different culture conditions which occur in batch bioreactors.²² According to the model, the microbial growth can be expressed as a sinusoidal curve, as described by eqn (3).

Integrating this equation, we get eqn (4),

Where, X is the biomass concentration (g L⁻¹), t is the time, X_M is the maximum biomass concentration, X₀ is the initial biomass concentration and K_C is the apparent specific growth rate (d⁻¹).

Result and discussion

Characterization of different wastes and nutrient removal capability of Chlorococcum sp.

The characteristic of untreated CETP and MSS effluents with respect to BOD, COD, TOC, PO₄, NH₄, NO₃ and NO₂ content and trace elements are summarised in Table 1. The concentration values of COD, NH₄, NO₃ and PO₄ present in CETP were 850 mg L⁻¹, 153 mg L⁻¹, 52 mg L⁻¹ and 11 mg L⁻¹, respectively, while those in MSS were 152 mg L⁻¹, 101 mg L⁻¹, 60 mg L⁻¹ and 13.6 mg L⁻¹, respectively. CETP showed higher value of COD as compared to MSS effluent, while no considerable difference was observed in the case of NH₄, NO₃ and PO₄. The lower values of wastewater constituents present in MSS effluent indicated its weak strength, which is a typical characteristic of municipal wastewaters.²³ The C/N ratio of CETP and MSS was found to be 4.14 and 0.94 respectively, whereas N/P ratio was observed to be 18.68 and 11.91, respectively. Thus, these ratios indicate that the CETP effluent used in this study has relatively higher carbon and nitrogen concentration. The suitable range of inorganic N/P ratio for algal growth is reported to be in the range of 6.8–10.²⁴

Table 1 Characterization of CETP and MSS wastewater effluent^a

Parameter	Units	Concentration
		CETP	MSS
a Values represent averages ± standard errors of data based on three independent determinations.
BOD	mg L^−1	460 ± 19.25	81.5 ± 9.61
COD	mg L^−1	850 ± 27.15	152 ± 6.81
TOC	mg L^−1	72 ± 3.61	15.5 ± 1.42
NH4–N	mg L^−1	153.1 ± 5.72	101.3 ± 5.83
NO3–N	mg L^−1	52 ± 2.96	60.2 ± 4.19
NO2–N	mg L^−1	0.4 ± 0.03	0.5 ± 0.02
PO4–P	mg L^−1	11 ± 1.62	13.6 ± 1.37
C/N	Ratio	4.14	0.94
N/P	Ratio	18.68	11.91
Alkalinity	mg L^−1	586 ± 21.95	242 ± 9.27
Hardness	mg L^−1	380 ± 15.37	220 ± 12.85
Turbidity	NTU	111.2 ± 4.16	30.6 ± 2.87
Phenol	mg L^−1	1.83 ± 0.05	0.13 ± 0.02
DO	mg L^−1	0.8 ± 0.02	1 ± 0.03
pH	—	9 ± 0.35	6.7 ± 0.21
Elements
Al	μg L^−1	53 ± 4.81	68.5 ± 13.9
Fe	μg L^−1	74.2 ± 4.27	61.48 ± 13.2
Cd	μg L^−1	1.63 ± 0.01	1.45 ± 0.43
Cr	μg L^−1	77.08 ± 8.52	64.55 ± 12.1
Co	μg L^−1	45.8 ± 10.71	119.7 ± 15.3
Ni	μg L^−1	50.6 ± 2.59	69.2 ± 7.03
Zn	μg L^−1	215.9 ± 21.02	334.95 ± 32.3
Sc	μg L^−1	4.1 ± 2.05	3.2 ± 1.3
V	μg L^−1	5.57 ± 1.58	2.62 ± 0.64
As	μg L^−1	0.45 ± 0.2	0.43 ± 0.31
Pb	μg L^−1	60.42 ± 3.54	69.03 ± 17.77
W	μg L^−1	14.23 ± 1.89	10.83 ± 2.11
Ag	μg L^−1	1.45 ± 0.15	1.68 ± 0.3
Ba	μg L^−1	12.5 ± 0.25	17.35 ± 5.64
Sr	μg L^−1	41.17 ± 2.88	86.27 ± 14.15
Cl	mg L^−1	408.4 ± 52.81	130.4 ± 5.07
Na	mg L^−1	35.72 ± 3.88	92 ± 8.0
K	mg L^−1	55.78 ± 4.23	43.53 ± 2.99
Ca	mg L^−1	34.9 ± 2.72	62.14 ± 3.51
Mn	mg L^−1	46.13 ± 3.11	12.02 ± 1.57
Mg	mg L^−1	13.22 ± 1.26	8.16 ± 0.97

---

The growth of Chlorococcum sp. in CETP and MSS over the period of 27 days resulted in significant reduction of overall nutrient concentrations. The treated effluent of CETP and MSS were analysed for nutrient removal capability with respect to BOD, COD, TOC, PO₄, NH₄, NO₃ and NO₂. The profile of nutrient removal concentration for all the major nutrient parameters by Chlorococcum sp. is shown in Fig. 1. In high carbon and nitrogen containing CETP effluent, the nutrient removal percentage by Chlorococcum sp. for BOD, COD and TOC were found to be 76.1%, 66.5% and 72.9% values, respectively. This indicates that Chlorococcum sp., in addition to its autotrophic nutritional mode, was also able to utilize organic matter in a mixotrophic mode, as also reported with Chlorella vulgaris.²⁵ The nutrient removal percentage for nitrogen substrates, such as NH₄, NO₃, and NO₂ were 92.9%, 97.21% and 100%, respectively, which was significantly higher than that of phosphate (PO₄) (76.7%). In MSS, almost complete removal of nitrogen sources to nearly 96.6%, 99.1% and 100% were noticed for NH₄, NO₃, and NO₂, respectively, while other factors such as COD, BOD, TOC and PO₄ were considerably reduced to 84.1%, 86.3%, 85.5% and 70%, respectively. The complete utilization of different forms of nitrogen sources not only reduced the pollutant load, but also made the media favourable (nitrogen limiting conditions) for higher lipid accumulation in microalgal cells. The nutrient removal in MSS was better compared to CETP owing to the N/P ratio of 11.91 and 18.68 for MSS and CETP, respectively. Besides algal growth, N/P ratio has a substantial effect on nutrient removal efficiency, and the ratio of 14 : 1 has been reported to be an optimum value.²⁶ The rise in pH and DO values were attributed to uptake of carbon dioxide and release of oxygen in the process of microalgal photosynthesis.²⁷

Fig. 1 Bioremediation capacity of various parameters by Chlorococcum sp. in CETP and MSS effluent. Change in residual concentration of parameters: COD, BOD, TOC, NH₄, NO₃, NO₂ and PO₄ are shown in from (a)–(g), respectively, where as the change in pH and DO is shown in (h) and (i). Values represent averages values with standard errors of data based on three independent determinations.

Poultry farming is one of the fastest growing agro-based industries due to the increasing demand for poultry meat and egg products. Nevertheless, the major environmental challenges associated with manure and litter wastes results in soil and water pollution through nutrients build-up, runoff and leaching.²⁸ This study was carried out to evaluate the bioremediation capacity of Chlorococcum sp. in different quantities of PW (1 g L⁻¹, 5 g L⁻¹, 10 g L⁻¹, 20 g L⁻¹ and 30 g L⁻¹) mixed with distilled water. Table 2 represents the composition of 1 g L⁻¹ of PW mixed in distilled water. The C/N and N/P ratio of 7.5 and 1.8 indicates the presence of high C, N and P content. However, the composition of poultry dropping waste could vary according to diet and age of the birds.²⁹

Table 2 Characteristic of PW wastewater with respect to 1 g L⁻¹ of PW concentration dissolved in water^a

Parameters units	TOC	COD	NH4–N	NO3–N	NO2–N	PO4–P
a Values represent averages ± standard errors of data based on three independent determinations.
mg L^−1	16.3 ± 0.5	179.1 ± 6.2	20.47 ± 1.2	3.26 ± 0.2	0.3 ± 0.01	12.9 ± 0.3

---

Nutrient removal concentrations of various PW concentrations (1–30 g L⁻¹) are elaborately shown in Fig. 2. The main nutrients analysed over the period of 27 days were PO₄, NH₄, NO₃, NO₂, COD and TOC. The complete nitrogen removal was achieved for NO₂ and NO₃ at all the PW concentrations. The nitrogen removal of NH₄ was significantly higher in 1, 5 and 10 g L⁻¹ PW concentrations to almost 100%, while the removal percentages declined to 65.9% and 51.4% for 20 and 30 g L⁻¹ PW concentration. This residual NH₄ could be responsible for the decreased algal photosynthesis and thus the growth rate (Fig. 4).²³ The results also indicated that the culture with 5 and 10 g L⁻¹ PW were under nitrogen starvation conditions at the end of trials, which is beneficial for higher lipid accumulation in algal cells.²⁰ PO₄ removal percentages were found to be 96.8%, 79.5%, 54.6%, 42.7% and 28.2% for 1, 5, 10, 20 and 30 g L⁻¹ PW concentration, respectively. The removal percentages of COD and TOC declined linearly with increase in PW concentration and the efficacy further reduces to below 50% for 20 to 30 g L⁻¹ PW concentration.

Fig. 2 Bioremediation capacity of various parameters by Chlorococcum sp. in various concentration of PW (1–30 g L⁻¹). Change in residual concentration of parameters: COD, TOC, NH₄, NO₃, PO₄ and NO₂ are shown in from (a)–(f), respectively. Values represent averages values with standard errors of data based on three independent determinations.

Fig. 3 Time-course profile of Chlorococcum sp. cultivated in the CETP and MSS wastewaters. Values represent average values with standard errors of data based on three independent determinations.

Fig. 4 Growth rate, biomass and lipid yield of Chlorococcum sp. in different concentration of PW ranging from 1 to 30 g L⁻¹ (PW-1, PW-5, PW-10, PW-20 and PW-30 correspond to PW concentration of 1 g L⁻¹, 5 g L⁻¹, 10 g L⁻¹, 20 g L⁻¹ and 30 g L⁻¹, respectively, mixed in distilled water). Values represent averages values with standard errors of data based on three independent determinations.

Influence of different types of wastes on biomass production of Chlorococcum sp.

The performance and time-course profile of Chlorococcum sp. in wastewater effluents as growth medium was evaluated under axenic condition and results are shown in Table 3 and Fig. 3. The biomass concentration of 1.45 g L⁻¹ and 1.16 g L⁻¹ was achieved in CETP and MSS, respectively, the change in biomass concentration was found to be proportional to the change in wastewater strength. This result demonstrates the ability of Chlorococcum sp. to utilize wastewater as a potential growth medium.

Table 3 Summary of growth and product parameters for Chlorococcum sp. cultivated in CETP and MSS effluents^a

WW effluent	Growth rate (d^−1)	Biomass yield (g L^−1)	Lipid content (%)	Lipid yield (mg L^−1)
a Values represent averages ± standard errors of data based on three independent determinations.
CETP	0.168 ± 0.02	1.45 ± 0.05	27.2 ± 1.8	394.4 ± 29.4
MSS	0.148 ± 0.01	1.16 ± 0.06	21.0 ± 1.5	243.6 ± 21.5

---

The microalgal culture with CETP yielded maximum biomass concentration of 1.45 g L⁻¹ (Table 2) as against 1.2 g L⁻¹ in the BBM as a control (data not shown).²⁰ However, the typical biomass concentration in domestic wastewater level ranges from 0.1–0.5 g L⁻¹.¹⁰ Biomass concentration of 0.4–1.5 g L⁻¹ was obtained when Chlorella vulgaris was grown in various wastewater streams.³⁰ Further, growth of microalga in MSS resulted in lower biomass yield of 1.16 g L⁻¹. The relative higher biomass yield in CETP might be due to the presence of essential nutrients, mainly carbon and nitrogen in large quantity compared to MSS. Overall, 21% increase in biomass concentration was achieved in CETP when compared with control, whereas cultivation in MSS effluents showed lower biomass concentration.

Growth rate, biomass and lipid yield of Chlorococcum sp. in various PW concentrations ranging from 1–30 g L⁻¹ is illustrated in Fig. 4. The maximum biomass concentration of 1.62 g L⁻¹ was obtained at 10 g L⁻¹ PW concentration. However, the biomass concentration decreased to 1.31 g L⁻¹ and 1.12 g L⁻¹ with the increase in PW concentration to 20 and 30 g L⁻¹ PW, respectively. This decline in growth observed was because of substrate inhibition mainly due to the presence of excess nutrients.²³ Thus, the preliminary experiments for determining suitable concentration of wastes or wastewaters are important, as algal photosynthesis can be inhibited by high nutrient concentrations, e.g., presence of excess of nitrogen species like ammonium ion and nitrite.^23,31

The overall microalgal cultivation in PW showed higher growth rate in the initial phase of growth compared to WW (CETP and MSS), corresponding to shorter lag time. The highest growth rate of 0.187 d⁻¹ was obtained with 10 g L⁻¹ PW concentration, where as with CETP and MSS the growth rate of 0.168 d⁻¹ and 0.148 d⁻¹ was observed, respectively (Table 3 and Fig. 4). This could be due to the high content of essential nutrients such as N, P, K, trace elements, etc. present in PW,²⁸ a similar observation was made in our previous work where high concentration of N, P, K and Fe had a positive effect on Chlorococcum sp. growth.²⁰ Biomass concentration was enhanced by 35% in PW with 10 g L⁻¹ concentration compared to control, whereas extreme of concentrations of 1 and 30 g L⁻¹ of PW showed negative trend (Fig. 3).

Influence of mixing of solid and liquid wastes on biomass production of Chlorococcum sp.

In the previous section, it was found that the algal growth was considerably enhanced in PW because of its much higher levels of essential nutrients (C, N, P, K, etc.) than the CETP and MSS wastewater. In addition, PW contains many necessary macro- and micro-nutrients in good amounts.²⁸ Therefore, the results showed that the PW was the most appropriate waste medium for algal growth. In this experiment, the strategy involving mixing of different liquid and solid wastes to further maximize microalgal biomass production was adopted. The effect of supplementation of various concentrations of PW such as 1, 5 and 10 g L⁻¹ with wastewaters (CETP and MSS) on microalgal biomass yield was evaluated. Interestingly, the strategy of mixing 5 g L⁻¹ PW with CETP and MSS resulted in most enhanced biomass yield of 2.22 g L⁻¹ and 1.95 g L⁻¹ (Table 4), respectively. The literature reports suggest that the use of wastewater solely results in relatively lower biomass yields ranging from a minimum of 0.02–0.11 g L⁻¹ to maximum value of 0.4–1.5 g L⁻¹.^10,30 The strategy involving supplementation of PW in WW provided essential nutrients, including nitrogen, phosphate and potassium, for the cultivation of microalgae thereby significantly increasing biomass yield.²⁸ The biomass yield was enhanced by 60–85% when compared to control (BBM).

Table 4 Growth rate, biomass yield, lipid content and yield of Chlorococcum sp. in CETP and MSS discharges with or without PL supplementation^a,b

PW concentration (g L^−1)	CETP				MSS
	Growth rate (d^−1)	Biomass yield (g L^−1)	Lipid content (%)	Lipid yield (mg L^−1)	Growth rate (d^−1)	Biomass yield (g L^−1)	Lipid content (%)	Lipid yield (mg L^−1)
a PW-0, 1, 5 10 correspond to supplementation of; zero PW (only WW), 1 g L^−1 of PW, 5 g L^−1 of PW and 10 g L^−1 of PW concentration, respectively, in CETP and MSS wastewater effluent.b Values represent averages ± standard errors of data based on three independent determinations.
PW-0	0.168 ± 0.02	1.45 ± 0.05	27.2 ± 1.8	394.4 ± 29.4	0.148 ± 0.01	1.16 ± 0.06	21.0 ± 1.5	243.6 ± 21.5
PW-1	0.173 ± 0.01	1.75 ± 0.07	26.3 ± 2.1	460.3 ± 41.1	0.165 ± 0.01	1.53 ± 0.09	23.7 ± 1.6	362.6 ± 32.5
PW-5	0.185 ± 0.01	2.22 ± 0.12	23.8 ± 1.3	528.4 ± 40.6	0.186 ± 0.02	1.95 ± 0.17	25.6 ± 2.5	499.2 ± 65.3
PW-10	0.170 ± 0.02	2.01 ± 0.10	13.5 ± 1.0	271.4 ± 24.2	0.175 ± 0.02	1.74 ± 0.14	14.8 ± 1.2	257.5 ± 29.4

---

Hence, the most optimal microalgal growth data corresponding to mixing of 5 g L⁻¹ PW with CETP were fitted onto the logistic growth model to validate the experimental data. The model showed a good fit between predicted values and experimental data (Fig. 5). Model fitting resulted in the regression coefficient (r²) higher than 0.98. The predicted value of maximum biomass concentration (X_M) obtained by the model fitting was 2.37 g L⁻¹ that was close to the experimental value of 2.2 g L⁻¹. The predicted initial biomass concentration X₀ by this model was 0.15 g L⁻¹. The apparent specific growth rate (K_C) for this experiment was found to be 0.21 d⁻¹. The predicted X_M and K_C values were 8% and 14% higher than the experimental values, respectively. Overall, the mixing of various wastes resulted in nearly 2-fold increase in biomass concentration when compared with standard medium (BBM) and by 1.5 fold when compared with CETP only.

Fig. 5 Experimental data and model prediction for growth of Chlorococcum sp. in mixes of waste (CETP + PW-5) using the logistic equation. Solid lines indicate the predicted values and the experimental values are shown by the symbols (closed circle). Data values represent averages values.

Lipid production potential of Chlorococcum sp. indifferent waste discharges

Lipid content

Change in lipid accumulation was affected by change in compositions and concentration of different waste discharges. The maximum lipid content of 27.2% (dcw, w/w) was obtained in CETP effluent (Table 2), whereas BBM (control) was reported to contain only 12% (w/w) lipid (data not shown). With varying effluent types, initial COD (carbon source) concentration also showed variation ranging from 152–5590 mg L⁻¹, nevertheless, COD values close to 850 mg L⁻¹ was found to be optimal for lipid accumulation. Similar trend was reported where COD concentration of 400–800 mg L⁻¹ resulted in highest lipid accumulation.⁵ In addition, the complete removal of different nitrogen sources from the wastewater to nearly 100% resulted in nitrogen limited or starvation condition which is considered favourable for inducing lipid accumulation.^1,20 Previous study also suggests that under nitrogen limiting conditions, presence of excess carbon in the medium is channelized into lipid biosynthesis (and/or starch) as storage products.²³ However, several authors have also suggested that in autotrophic or heterotrophic cultures, lipid accumulation might be due to excess carbon in the culture medium, which get utilized at a higher rate than cell growth rate and hence promote conversion of carbon to lipids.³² Change in PW concentration significantly affected the lipid accumulation. Maximum lipid content of 25.1% in PW-5 g L⁻¹ and lowest of 13% in PW-30 g L⁻¹ was observed (Table 4). As the nutrient concentration in PW increased the lipid content of Chlorococcum sp. decreased from 27.2% to 13% of the dry weight. PW-5 mixed with CETP and MSS resulted in lipid content of 23.8% and 25.6%, respectively, while significantly higher lipid yield of 528.4 mg L⁻¹ and 499.2 mg L⁻¹ was observed. This could be due to the higher biomass concentration obtained at the end of trials as well as the presence of lower nitrogen and higher carbon reserves in the cultivation medium.²³ The use of anaerobically digested poultry litter effluent has been reported as a suitable medium for microalgal biomass production, containing high amount of protein, carbohydrate and lipid.³³ Thus, the approach of judicious mixing of different wastes could be effectively used as an alternative culture medium to produce increased biomass and lipid yields for biodiesel application.

Fatty acid composition

The lipid was transesterified and converted into fatty acid methyl esters (FAME), the fatty acid compositions of Chlorococcum sp. was determined and the presence of C14:0, C15:1, C16:0, C16:1, C17:1, C18:0, C18:1, C18:2, C18:3, C20:0,and C20:2 were detected. The predominant fatty acids present in the microalgae were mainly C16:0, C18:0, C18:1, C18:2, and C18:3 and their total fatty acid percentages (%) were in the range of 19.5–33.8%, 4.2–27.5%, 9.3–33.4%, 10.1–25.1%, and 6.7–18.2%, respectively. On the other hand, the remaining fatty acids such as C14:0, C15:1, C16:1, C17:1, C18:0, C20:0 and C20:2 were less common and ranged from 0.7–7.5%. When the Chlorococcum sp. was cultivated in waste discharges, the algal lipid largely comprised of C16 and C18 fatty acids in the range of 78.7–90.03% of the total fatty acids, whereas control cultivated in BBM resulted in only 67.35% as shown in Fig. 6. Moreover, the proportion of C16:0 (palmitic acid) and C18:1 (oleic acid) fatty acids accounted nearly 50% of the total fatty acids. The biodiesel fuel properties could be directly determined by the composition of FAME components.^2,34 Generally lipids containing C16 and C18 fatty acids are considered as essential component for biodiesel, which imparts good fuel properties, including its ignition quality, combustion heat, cold filter plugging point (CFPP), oxidative stability, viscosity, and lubricity.⁵ The variation in FAME profiles was observed using different wastewaters, as lipid content and fatty acid composition are greatly affected by culture parameters, growth phase and other environmental conditions.⁵ Overall, unsaturated fatty acids (UFA) and saturated fatty acids (SFA) were in the range of 43.79–67.02% and 30.76–55%, respectively. The UFA were found to be predominant in the FAME profile in the cultures associated with CETP and MSS effluent, corresponding to reduction in pour point value of biodiesel,^5,35 thus making the use of algal biodiesel more feasible in countries with cold climates.

Fig. 6 Fatty acid methyl ester (FAME) composition of biodiesel produced from Chlorococcum sp. grown under various conditions. Values represent averages values with standard errors of data based on three independent determinations.

Conclusion

This study demonstrated the development of an integrated sustainable process that involved the effective utilization of different waste discharges as alternative media for microalgal cultivation towards achieving concomitant biofuel generation and water pollution management. Chlorococcum sp. was able to grow efficiently in various wastewater effluents and produced high biomass and lipid yields. Wastewater remediation was satisfactorily accomplished with near complete removal of nitrogen and partial removal of carbon and phosphorous. Biomass and lipid yields were much greater when solid and liquid waste mixes, instead of any single waste, was used. The highest biomass yield of 2.2 g L⁻¹ and lipid yield of 528.4 mg L⁻¹ was achieved when 5 g L⁻¹ PW was mixed with CETP effluent. Thus, the attempt of mixing of different types of liquid and solid wastes meaningfully served the dual purpose of waste management and microalgal biomass production for biofuel application in an integrated biorefinery model. This model is potentially cost-effective as it replaces the precious freshwater and expensive synthetic nutrients or fertilizers with cheap industrial and municipal wastes in cultivation media. In the end, the integrated microalgal feedstock based biorefinery for simultaneous wastewater treatment and biofuels production may pave the way for a greener environment and sustainable biobased economy.

Acknowledgements

AK gratefully acknowledges Department of Biotechnology (DBT), New Delhi, Govt. of India for the research fellowship as SRF. The authors also acknowledge Department of Science & Technology (DST), Govt. of India, for the financial support (Project No. DST/IS-STAC/CO2-SR-160/13(G); Date: 08.07.2013).

References

1. Y. Chisti, Biotechnol. Adv., 2007, 25, 294–306.
2. R. a I. Abou-Shanab, M. K. Ji, H. C. Kim, K. J. Paeng and B. H. Jeon, J. Environ. Manage., 2013, 115, 257–264.
3. Z. Arbib, J. Ruiz, P. Álvarez-Díaz, C. Garrido-Pérez and J. a. Perales, Water Res., 2014, 49, 465–474.
4. J. Martinez, P. Dabert, S. Barrington and C. Burton, Bioresour. Technol., 2009, 100, 5527–5536.
5. L. Zhu, Z. Wang, Q. Shu, J. Takala, E. Hiltunen, P. Feng and Z. Yuan, Water Res., 2013, 47, 4294–4302.
6. S. S. Wong, T. T. Teng, a. L. Ahmad, a. Zuhairi and G. Najafpour, J. Hazard. Mater., 2006, 135, 378–388.
7. A. F. Aravantinou, M. a. Theodorakopoulos and I. D. Manariotis, Bioresour. Technol., 2013, 147, 130–134.
8. T. Cai, S. Y. Park and Y. Li, Renewable Sustainable Energy Rev., 2013, 19, 360–369.
9. P. T. Pienkos and A. Darzins, Biofuels, Bioprod. Biorefin., 2009, 3, 431–440.
10. L. Xin, H. Hong-ying and Y. Jia, New Biotechnol., 2010, 27, 59–63.
11. P. J. McGinn, K. E. Dickinson, S. Bhatti, J. C. Frigon, S. R. Guiot and S. J. B. O'Leary, Photosynth. Res., 2011, 109, 231–247.
12. T. J. Lundquist, I. C. Woertz, N. W. T. Quinn and J. R. Benemann, Energy, 2010, 1.
13. L. Christenson and R. Sims, Biotechnol. Adv., 2011, 29, 686–702.
14. G. Markou, I. Chatzipavlidis and D. Georgakakis, Bioresour. Technol., 2012, 112, 234–241.
15. T. Cai, S. Y. Park, R. Racharaks and Y. Li, Appl. Energy, 2013, 108, 486–492.
16. S. B. Ummalyma and R. K. Sukumaran, Bioresour. Technol., 2014, 165, 295–301.
17. B. H. Liu and Y. K. Lee, Biotechnol. Lett., 1999, 21, 1007–1010.
18. P. Feng, Z. Deng, Z. Hu, Z. Wang and L. Fan, Bioresour. Technol., 2014, 162, 115–122.
19. R. Harun and M. K. Danquah, Appl. Energy, 2011, 46, 304–309.
20. A. Karemore, R. Pal and R. Sen, Algal Res., 2013, 2, 113–121.
21. Association American Public Health, Standard methods for the examination of water and wastewater, American Public Health Association, Washington, DC, 20th edn, 1998.
22. K. Kumar, D. Banerjee and D. Das, Bioresour. Technol., 2014, 152, 225–233.
23. I. Rawat, V. Bhola, R. R. Kumar and F. Bux, Environ. Technol., 2013, 34, 1765–1775.
24. L. Wang, M. Min, Y. Li, P. Chen, Y. Chen, Y. Liu, Y. Wang and R. Ruan, Appl. Biochem. Biotechnol., 2010, 162, 1174–1186.
25. A. P. Abreu, B. Fernandes, A. a. Vicente, J. Teixeira and G. Dragone, Bioresour. Technol., 2012, 118, 61–66.
26. S. H. Lee, C. Y. Ahn, B. H. Jo, S. A. Lee, J. Y. Park, K. G. An and H. M. Oh, J. Microbiol. Biotechnol., 2013, 23, 92–98.
27. S. Mandal and N. Mallick, Appl. Environ. Microbiol., 2011, 77, 374–377.
28. N. S. Bolan, A. a. Szogi, T. Chuasavathi, B. Seshadri, M. J. Rothrock and P. Panneerselvam, World's Poultry Science Association, 2010, 66, 673–698.
29. M. Mahadevaswamy and L. V. Venkataraman, Agric. Wastes, 1986, 18, 93–101.
30. I. T. D. Cabanelas, J. Ruiz, Z. Arbib, F. A. Chinalia, C. Garrido-Pérez, F. Rogalla, I. A. Nascimento and J. a. Perales, Bioresour. Technol., 2013, 131, 429–436.
31. M. K. Ji, H. C. Kim, V. R. Sapireddy, H. S. Yun, R. a I. Abou-Shanab, J. Choi, W. Lee, T. C. Timmes, Inamuddin and B. H. Jeon, Appl. Microbiol. Biotechnol., 2013, 97, 2701–2710.
32. B. Riaño, D. Hernández and M. C. García-González, Ecol. Chem. Eng. A, 2012, 49, 112–117.
33. M. Singh, D. L. Reynolds and K. C. Das, Bioresour. Technol., 2011, 102, 10841–10848.
34. R. a I. Abou-Shanab, J. H. Hwang, Y. Cho, B. Min and B. H. Jeon, Appl. Energy, 2011, 88, 3300–3306.
35. a. a. Abdelmalik, a. P. Abbott, J. C. Fothergill, S. Dodd and R. C. Harris, Ind. Crops Prod., 2011, 33, 532–536.

---