1 Introduction

The need for fossil fuels, including oil, coal, and natural gases, has increased dramatically due to the rapid growth of the world population, economic growth, and rapid industrialization. Consequently, fossil fuels are running out more quickly. It is estimated that natural gas and oil reserves will be exhausted in between 40 and 64 years. By 2030, the global energy demand will have roughly a 60% increase [1]. Moreover, the risk of global warming due to anthropogenic CO2 emissions is related to coal, natural gas, and petrol combustion.

New renewable energy sources need to be developed to overcome all these problems. One of the most promising biofuels that has received much interest in recent years is biodiesel which considers a sustainable, eco-friendly, and CO2- emission-free alternative fuel. Microalgae’s quick growth, great photosynthetic efficiency, and lack of dependence on arable land make them a promising feedstock for manufacturing biodiesel. Also, microalgae can utilize CO2, nitrogen, and phosphate during their cultivation, leading to the greenhouse effect and water pollution mitigation. To produce 1 kg of biomass, approximately 1.8 kg of CO2 is required [2]. According to Liu et al. [3], up to 80% of the biomass that microalgae produce can be stored as lipids, and triacylglycerols (TAGs), which make up most of these lipids, can be used as a feedstock for the transesterification of lipid to create biodiesel. Since transportation expenses are generally low, microalgal oil is more affordable than biomass from trees and crops [4]. The fundamental properties of biodiesel made from microalgal oil are similar to those of conventional biodiesel, such as their high degree of saturation, cetane number, and viscosity [5].

There are other methods than large-scale algae production for increasing biodiesel generation. The metabolic process changes when the cultivation conditions are improved, which increases the generation of relative and mesh biodiesel. Numerous growth factors such as the availability of nutrients, light intensity, salinity, the presence of heavy metals, the pH scale, and the temperature significantly impact microalgal growth and metabolism, including lipid accumulation [6,7,8]. Under optimal growth conditions, microalgae produce fatty acids primarily for esterification into glycerol-based membrane lipids, while under unfavorable stressed conditions, many microalgae alter their fat biosynthesis pathways to synthesize and accumulate neutral lipids, primarily in the form of triacylglycerides, which make up about 5 to 20% of the cell dry weight (20–50% DW) [9, 10]. Organic carbon sources can serve as enhancer compounds and provide a mixotrophic condition preferable for algal growth and lipid production [11,12,13]. However, there is not much research addressing how combined stressors affect lipid productivity and fatty acid profile in biodiesel quality, whether in phototrophic or mixotrophic environments.

This study aimed to determine the ideal treatment scenario for M. salina in terms of biomass and lipid production as well as biodiesel quality by examining the effects of various nitrogen sources, salt stress, and carbon enrichment, both separately and in combination.

2 Materials and methods

2.1 Cultivation and growth condition

The strain Microchloropsis salina (formerly Nannochloropsis salina) was provided by the National Institute of Oceanography and Fisheries in Alexandria, Egypt. In 1-L Erlenmeyer flasks (flasks were stopped with cotton plugs), 800 mL of artificial seawater (ASW) fortified with F/2 nutrients had its NaNO3 concentration changed from 0.075 to 0.75 g L−1 [14]. The flasks were sterilized in an autoclave at 121 °C and 1.5 atom for 20 min. After cooling, the Erlenmeyer flasks were inoculated with a certain volume (10 mL) of microalgal strain pre-culture in 250-mL Erlenmeyer flasks. Aeration was given to the culture through silicon tubing, with one end inserted into the culture flask and the other into the bacterial filter connected to the aerator. Algal culture flasks were incubated under the continuous fluorescent light of 45 µmole m−2 s−1 and at a temperature of 25 ± 1 °C.

2.2 Optimization of media

2.2.1 Different nitrogen sources

In order to examine the effects of various nitrogen sources on the cell growth and lipid production of M. salina, potassium nitrate (KNO3, 0.892 g L−1), glycine (NH2CH2COOH, 0.662 g L−1), urea (CO (NH2)2, 0.265 g L−1), ammonium chloride (NH4Cl, 0.472 g L−1), and yeast extract (YE) (initial nitrogen concentration was the same, at 8.82 mM (123.5 mg N L−1) were used in place of sodium nitrate (NaNO3, 0.750 g L−1) in F/2 medium according to Kim et al.[15].

2.2.2 Salt stress

Four different sodium chloride (NaCl) concentrations (100, 200, 300, and 400 mM) were used to test the effect of salt stress.

2.2.3 Carbon enrichment

Different concentrations of three different carbon sources, glucose, glycerol, and sodium acetate (1, 2, and 3 g L−1), were used.

2.3 Measurement of microalgal growth and biomass

2.3.1 Optical density

Every 2 days, optical density at 680 nm was measured using a spectrophotometer to evaluate the growth of each algal culture. The resulting absorbance was used to plot the growth curve of respective algae cultures.

2.3.2 Dry weight measurement

A specific amount (30 mL) of the algal suspension was centrifuged at 4000 rpm for 15 min, followed by numerous thorough washes with distilled water. The precipitated biomass was dried overnight at 60 °C in an oven in pre-weighed Petri plates until constant weight after removing all salts from the medium. Data were given by g L−1 [16].

2.3.3 Estimation of biomass productivity

According to Eq. (1), which is discussed by Abomohra et al. (2013) [17], biomass productivity was calculated.

$$\mathrm{Biomass}\;\mathrm{productivity}\;\mathrm{BP};\left(\mathrm gL^{-1}\mathrm{day}^{-1}\right)={\mathrm{CDW}}_{\mathrm L}-{\mathrm{CDW}}_{\mathrm E}/\left(t_L-t_E\right)$$
(1)

where CDWL (g L−1) is the cellular dry weight at the late exponential phase (tL) and CDWE (g L−1) is the cellular dry weight at the early exponential phase (tE).

2.4 Estimation of total lipids and productivity

This work adapted the Park procedure to use the SPV reaction to detect lipid content directly [18]. Initial preparation of the phospho-vanillin reagent involved constantly stirring until 0.06 g of vanillin was dissolved in 10 mL of deionized water, followed by the addition of 40 mL of concentrated phosphoric acid to the mixture. The biomass was extracted by centrifugation at 4000 RPM for 5 min using a known amount of biomass. The sample was treated with 2 mL of concentrated sulfuric acid (98%) and heated in a water bath at 100 °C for 10 min. The reaction mixture was then added to 5 mL of freshly made phospho-vanillin reagent once it had cooled, and it was all incubated for 15 min at 37 °C in the incubator. After that, a pink color emerged, and its absorbance was measured at 530 nm with a spectrophotometer to determine how much lipid was present in the sample. The standard curve was used to estimate the amount of lipids. The content of lipids was expressed as μg mL−1. According to Eq. (2), lipid productivity was calculated as discussed earlier by Abomohra et al. (2013) [17].

$$\mathrm{Lipid}\;\mathrm{productivity}\;\mathrm{LP};\left(\mathrm\mu\mathrm g\mathrm L^{-1}\mathrm{day}^{-1}\right)={\mathrm{LC}}_{\mathrm L}-{\mathrm{LC}}_{\mathrm E}/\left(t_L-t_E\right)$$
(2)

where LCL (μg mL−1) is the lipid content at the late exponential phase (tL) and LCE is the lipid content (μg mL−1) at the early exponential phase (tE).

2.5 Extraction of total lipids

Following certain changes, the modified Folch technique [19] was used to extract the total lipid content. After centrifuging 50 mL of algal culture for 15 min at 4500 rpm, 40 mL of chloroform/methanol (2:1, v/v) solution was added to the dried algal cells. To separate the liquid phase, the mixture was filtered through filter paper (Whatman No. 1) after being shaken at 120 rpm for 48 h at room temperature. The liquid phase was transferred to a fresh flask, rinsed with 0.9% NaCl (w/v) solution, and the two phases were separated using a separatory funnel. In order to evaporate the solvent, the lower phase containing the lipids was taken in a pre-weighed glass vial and dried at 40 °C for 2 days. The difference between W2 (weight of glass vial containing lipids) and W1 (weight of glass vial) indicates the quantity of total lipid content.

2.6 Cultivation of Microchloropsis salina in combined conditions

The most effective growth and total lipid content–affecting elements from the earlier trials were chosen, merged, and approved as the ideal conditions for additional experimentation and analysis.

2.7 Fatty acid profiles determination

For the preparation of fatty acid methyl esters (FAMEs), 1.0 mL of n-hexane was added to 15 mg of oil, followed by 1.0 mL of sodium methoxide (0.4 mol), according to the modified method [20]. After being vortexed for 30 s, the mixtures were left to settle for 15 min. The upper phase that contained the FAMEs was recovered and analyzed using gas chromatography (Perkin Elmer model: flame ionization detector). A Perkin Elmer Auto System XL with a flame ionization detector (FID) was used for the GC analysis. The capillary column was a DB-Wax made of fused silica (60 m × 0.32 mm i.d.). The oven’s temperature was initially kept at 150 °C and was designed to rise by 3 °C/min to 220 °C. 1.1 ml/min of helium was used as the carrier gas. Temperatures for the injector and detector were 230 and 250 °C, respectively.

2.8 Evaluation of some biodiesel properties

The biodiesel properties were calculated using the FAME composition of the microalgae, including the Average Degree of Unsaturation (ADU percent), Iodine Value (IV, g I2.100 g−1 oil), Cetane Number (CN), Kinematic viscosity (ʋi, mm2 s−1), Cloud point (CP, °C), density (ρ), Higher Heating Value (HHV), Saponification Value (SV, mg KOH g−1), Long Chain Saturation Factor (LCSF, wt %), and Cold Filter Plugging Point (CFPP, °C) using the following Eqs. (3–12) discussed by [21,22,23,24,25].

$$\mathrm{ADU}=\sum N\times \mathrm{Mf}$$
(3)

where N is the number of carbon–carbon double bonds in fatty acid and Mf is the mass fraction of each fatty acid.

$$\mathrm{CN}=-6.6684\;\mathrm{ADU}+62.876$$
(4)
$$\rho=0.0055\;\mathrm{ADU}+0.8726$$
(5)
$$v_i=-0.6313\;\mathrm{ADU}+5.2065$$
(6)
$$\mathrm{CP}=3.356\;\mathrm{ADU}+19.994$$
(7)
$$\mathrm{HHV}=1.7601\;\mathrm{ADU}+38.534$$
(8)
$$\mathrm{IV}=74.373\;\mathrm{ADU}+12.71$$
(9)
$$\mathrm{SV}=\sum \left(560\times {N}_{i}\right)/{M}_{i}$$
(10)

where Ni is the percentage of each FAME, and Mi is the molecular weight of each FAME.

$$\mathrm{LCSF}\;\left(\mathrm w\mathrm t\%\right)=\left(0.1\times\mathrm C16:0\right)+\left(0.5\times\mathrm C18:0\right)+\left(1\times\mathrm C20:0\right)+\left(1.5\times\mathrm C22:0\right)+\left(2\times\mathrm C24:0\right)$$
(11)
$$\mathrm{CFPP}\left(\mathrm{^\circ{\rm C} }\right)=\left(3.1417\times \mathrm{LCSF}\right)-16.477$$
(12)

where C16:0, C18:0, C20:0, C22:0, and C24:0 are the weight percentage of the corresponding fatty acids.

2.9 Statistical analysis

The mean and standard deviation (SD) of three replicates were used to express the results. Using the SPSS 23.0 program, the collected data were statistically evaluated by one-way analysis of variance (ANOVA), followed by Duncan’s multiple range testing for data with a significant difference, at p < 0.05.

3 Results and discussion

3.1 Growth and lipids production under different nitrogen sources of the microalga

It was found that M. salina efficiently utilizes organic nitrogen (particularly glycine and YE and, to a lesser degree, urea) and nitrate (NaNO3 and KNO3) for growth, but not ammonium. As shown in Fig. 1, the highest growth of M. salina was observed in Glycine (OD680 value was 2.40) and YE (OD680 value was 2.25) on the 12th day of cultivation followed by potassium nitrate (OD680 value was 1.95), sodium nitrate (OD680 value was 1.85), and urea (OD680 value was 1.39). Cell growth inhibition was observed in culture with ammonium chloride (OD680 value was 0.31).

Fig. 1
figure 1

Growth curves of M. salina grown on different sources of nitrogen sources determined by measuring optical density at 680 nm

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The results in Table 1 reveal that YE resulted in the highest biomass and lipid productivity among all six nitrogen sources. This improvement might be caused by the fact that YE contains a variety of other substances in addition to nitrogen, such as amino acids, peptides, vitamins, and carbs. Because M. salina appears to use both nitrogen and organic molecules in YE through mixotrophic metabolism, cell growth and lipid production were enhanced. Glycine, which had the second highest cell concentration and lipids content, can provide the dissolved free amino acids in a culture medium that microalgae can easily utilize [26]. It was reported that Tetraselmis sp. had the maximum cell concentration and lipid productivity under YE, then comes glycine [15]. Although M. salina continued to grow effectively in the presence of urea supplementation, the final cell concentration and lipid output were somewhat lower than those seen when nitrate was present.

Table 1 Effect of different nitrogen sources on biomass and lipid production by Microchloropsis salina
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On the other hand, it was reported that Isochrysis galbana be able to create the highest cell concentration in urea rather than nitrate or nitrite [27]. Cell growth was inhibited when ammonium chloride was supplemented, showing that the 8.82 mM concentration was toxic to cell growth. The excessive transfer of ammonium to the cells might interfere with the creation of ATP in the chloroplast, inhibiting photosynthesis, which is the cause of the inhibitory effect on cell growth [28]. Additionally, in some cases, excessive ammonium in the medium can cause a significant pH drop by releasing H+ ions, which prevents cell development and may even lead to cell lysis [29, 30].

3.2 Microalgal growth and lipids production under salt stress

As shown in Fig. 2, the highest growth of M. salina was observed at a concentration of 300 mM (OD 680 value was 2.41), followed by a concentration of 200 mM (OD 680 value was 2.19), control (OD 680 value was 1.85), the concentration of 100 mM (OD 680 value was 1.76), and concentration 400 Mm (OD 680 value was 1.66).

Fig. 2
figure 2

Growth curves of M. salina grown on different concentrations of sodium chloride determined by measuring optical density at 680 nm

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Table 2 reveals that the best concentration of NaCl for growth and lipids accumulation was 300 Mm. These results are in agreement with that reported by Zulkifli et al. [31], on which 400 mM of NaCl showed the highest cell density of N. oculata as compared to other salinity concentration among other salinity concentrations.

Table 2 Biomass and lipid production of Microchloropsis salina under different concentrations of sodium chloride
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These enhancements are due to some Nannochloropsis species being halophilic, and salinity strongly impacts their cellular physiology. Oxidative stress may be sodium chloride’s primary function in the algal medium, leading to the increment of the concentration of triacylglycerol in the cytoplasm [32, 33] and/or changes in lipid amount and composition [34]. The high lipid content at 300 mM NaCl may be due to adaptation under stress conditions, which help in the accumulation of lipid content in cells. In axenic conditions, salt stress was also used to maintain the culture leading to increased lipid production [35].

According to reports, M. salina’s biomass was maximum between salinities of 22 and 34%, whereas its highest lipid content was found at a salinity of 34% [36]. N. oculata can tolerate salinities between 10 and 35%, and the optimal salinity for their growth was 25% in nutrient-rich environments [37].

3.3 Microalgal growth and lipids production under carbon enrichment

It was found that M. salina effectively took up and metabolized both glucose and glycerol but not acetate. The best concentration of glycerol was 2 g L−1 which showed the highest growth of M. salina (OD 680 value was 2.50) (Fig. 3a), while the best concentration of glucose was 3 g L−1 (OD 680 value was 2.27) (Fig. 3b). Cell growth inhibition was observed in culture using all three concentrations of acetate (Fig. 3c).

Fig. 3
figure 3

Growth curves of M. salina grown on different organic carbon sources determined by measuring optical density at 680 nm. a Different concentrations of glycerol. b Different concentrations of glucose. c Different concentrations of sodium acetate

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Obtained results in Table 3 reveal that the final concentration of biomass and lipids accumulations of M. salina were higher than photoautotrophic control when grown mixotrophy on glycerol (2 g L−1) and glucose (3 g L−1). This was not the case when acetate was used. Although many microalgae are known to metabolize acetate [38, 39] easily, M. salina cannot. The results show that M. salina can utilize glycerol more effectively than glucose. Regarding lipid production, glycerol was a better organic carbon source than glucose at its best concentration of 3 g L−1 in the concentration range of (1:3) g L−1. It was found that the maximum biomass productivity in mixotrophic batch cultures using glucose or glycerol was identical at 170 mg L−1 day−1, being 30% higher than in control cultures. The optimal concentrations of the organic carbon sources for the growth of N. gaditana were 5 g L−1 for glucose and 1 g L−1 for glycerol. [40].

Table 3 Effect of different carbon sources on biomass and lipid production by Microchloropsis salina
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3.4 Cultivation of Microchloropsis salina under combined

The optimal level of each effect was used (optimized media) in accordance with the earlier findings, as M. salina was grown in F/2 medium with YE (123.5 mg N L−1), sodium chloride (300 mM), and glycerol (2 g L−1). The obtained results in Fig. 4 show that while the examined microalga’s total lipid content and lipid productivity significantly increased, their dry weight and biomass productivity decreased by 18 and 16%, respectively. M. salina’s lipid content and lipid productivity of M. salina increased by 72 and 80%, respectively.

Fig. 4
figure 4

Cellular dry weight (CDW) and lipids productivity (LP) of M. salina grown under control and combined conditions. a CDW. b LP. Error bar show the SD of three measurements

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3.5 Gas chromatography (GC) analysis of fatty acids

The given data in Table 4 and Fig. 5 reveal that the fatty acid profiles differ significantly (quantity and quality level). It was found that SFAs in all conditions are higher than the control as SFAs of culture cultivated in optimized conditions are more than onefold than control. Also, SFAs dramatically increased when the culture was grown on NaCl at a concentration of 300 mM and glycerol at a concentration (2 g L−1) by 45% and 42%, respectively. SFAs moderately increased by 27% when the culture was grown on a nitrogen source (YE). Regardless of the culture’s condition, the main fatty acids were C16:0, C18:1, C18:2, and C18:3 (palmitic, oleic, linoleic, and linolenic acids).

Table 4 Fatty acids profiles analysis of M. salina cultivated under control, YE, glycerol (2 g L−1), NaCL (300 mM), and combined culture (data are expressed as relative percentage)
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Fig. 5
figure 5

Fatty acids percentage (%) of different treatments of M. salina (SFAs, saturated fatty acids; MUFAs, monounsaturated fatty acids; PUFAs, polyunsaturated fatty acids)

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Oleic acid (C18:1) and palmitic acid (C16:0) were higher in the culture with YE, whereas the concentrations of linoleic acid (C18:2) and linolenic acid (C18:3) were lower than in the control culture. It was found that Tetraselmis sp. had linoleic acid (C18:2) and linolenic acid (C18:3) with higher concentrations than the culture with F/2 medium, and stearic acid (C18:0) content was significantly lower than those in the culture with YE [15].

Other fatty acids, such as lauric acid (C12:0), myristic acid (C14:0), pentadecanoic acid (C15:0), cis-10-heptadecanoic acid (C17:1), elaidic acid (C18:1), and linolelaidic acid (C18:2), are also found in addition to the major fatty acids under Nacl concentration 300 Mm. When Tetraselmis suecica cells are cultivated in 0.6 M NaCl, many additional fatty acids, including pentadecanoic acid (C15:0), cis-10-heptadecanoic acid (C17:1), elaidic acid (C18:1), and linolelaidic acid (C18:2), are discovered in addition to the two primary saturated fatty acids [41].

The fatty acid methyl ester profile (FAME) of M. salina grown mixotrophically using glycerol constitutes 86.5% of total fatty acids ranging between 16 and 18 carbon chain lengths which are the most abundant fatty acids needed for biodiesel application. FAME profile shows palmitic acid (C16:0) with a high percentage of 30.3%, followed by oleic acid (18:1) with 25.8%. These findings corroborated those of [42], who discovered that the FAME profile of E. gracilis under mixotrophic cultivation mode using glycerol contributes 93.46% of total fatty acids ranging between 16 and 18 carbon chain lengths and shows the presence of 24.17 and 25.8% of palmitic acid (C16:0) and oleic acid (18:1), respectively. Additionally, according to the C. vulgaris fatty acid profiles, the saturated fatty acid content increased from 16.91% in autotrophic conditions to 34.94% in the medium containing 5 g L−1 glycerol [43].

Under combined conditions, the content of SFAs greatly increased from 23.5 to 48.4%; and palmitic acid (C16:0) and stearic acid are the two main fatty acids (C18:0). It was reported that total fatty acids in N. oceanica increased under the combination of high salinity and high light [44]. In comparison to a single stress condition or a combination of more than three stress elements, lipid synthesis was significantly improved when two stress conditions were present. Although various research [45,46,47] have shown the positive effects of coupled stress factors on fat accumulation, the underlying mechanisms are hardly examined.

3.6 Biodiesel properties

Since saturated fatty acids positively impact transesterification and their biodiesel quality [7], the selected treatments showed high saturated and low unsaturated fatty acid content, which were recommended for good biodiesel quality. By comparing the obtained data with that of American standardization (ASTM D-6751) and European standardization (EN 14,214), cetane number (CN), iodine value (IV), density (ρ), and viscosity (V) which are considered limiting factors for the quality of biodiesel; they fall within the acceptable range (Table 5). The cetane number (CN) is a measure of how long it takes for fuel to ignite in diesel cycle engines. The ignition time decreases with increasing CN. The increase in CN is connected with a longer unbranched carbon chain in the FAME component parts [46].

Table 5 Evaluation of theoretical biodiesel properties of detected fatty acids of M. salina cultivated under control, YE, glycerol (2 g L−1), NaCL (300 mM), and combined conditions
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On the other hand, a lower IV value is advantageous for the manufacture of Biodiesel [47], as this quality is influenced by the number and location of double bonds in the alkyl esters’ carbon chains. There is a greater chance of oxidation, deposit development, and lubricity degradation of the biodiesel with the IV. The density and viscosity of the biodiesel increase with the length of the carbon chain in the methyl esters, and these characteristics will drop as the number of double bonds rises [46]. SV value can reflect the highest fatty acid contents. Saturated fatty acids (SFAs) may also lessen the CFPP properties of biodiesel because they have substantially higher melting points than unsaturated fatty acids (USFAs) [25].

4 Conclusion

Organic nitrogen sources, salt stress, and mixotrophic nutrition by glycerol, either individually or in combination, have significantly improved the productivities of biomass and lipid in M. salina. The best treatment condition included YE as a nitrogen source, 300 mM sodium chloride, and glycerol (g L−1). Individually, YE enhanced lipid content by more than on fold compared to control with only 24% lipids. Also, 300 mM NaCl improved lipids productivity by 92%, and glycerol (g L−1) promoted lipid productivity by 83%. Under the combined condition, lipid productivity increased by 80%, while biomass productivity reduced by 16%.

In all treatments, the fatty acids profile revealed a notable increase in saturated fatty acids, a discernible drop in polyunsaturated fatty acids, and no notable changes in monounsaturated fatty acids. This study provides experimental results on increasing biomass and lipid productivities and improving the fatty acids profile of M. salina for viable biodiesel production. However, further research efforts should be encouraged to achieve the necessary developments to implement this approach on a large scale.