Biofuels from Microalgae: A Review on Microalgae Cultivation, Biodiesel Production Techniques and Storage Stability

Abstract

Rising global energy demands, depleting fossil fuel reserves, and growing environmental concerns have led to an increasing demand for clean and renewable energy sources. Recently, microalgae biofuels have emerged as a promising and sustainable energy source due to their high biomass productivity, lipid content, and wastewater treatment capabilities. However, the viability of microalgae biofuels as a commercial-scale renewable fuel remains uncertain due to high production costs and storage stability issues. This review focuses on advanced technologies aimed at enhancing both the production of microalgae biodiesel and its storage stability. It explores the potential and challenges of recent developments in microalgae cultivation systems, particularly those factors that have contributed to increased lipid content in microalgae biomass. The study also examines the role of industrial wastewater in promoting microalgae growth and provides an overview of recent advances in biodiesel production. Additionally, it discusses various strategies to improve the storage stability of biodiesel, a critical consideration for the commercialization of microalgae biodiesel.

1. Introduction

Modern industry, urbanization, and modernization have all contributed to an increased consumption of fossil fuels. However, due to significant environmental issues, the depletion of reserves, and fluctuations in prices, this fuel has become increasingly unsustainable. Furthermore, the combustion of these fossil fuels generates greenhouse gas emissions, contributing to global warming and environmental degradation [1]. To address these issues, scientists have shifted their focus toward the development of renewable energy sources, which has become a primary research focus in recent years. Among various renewable energy sources, biofuels have shown significant potential to meet current energy demands while also reducing carbon emissions. According to India’s National Biofuel Policy 2018 (GAIN Report Number: IN9069, Date: 8 February 2019), the country aims to achieve a 20% ethanol (E20) and 5% biodiesel (B5) blend with gasoline by 2030. However, the competition for arable land between food crops and biofuel production has hindered the commercial viability of first- and second-generation biofuels. Currently, microalgae have emerged as the most promising feedstock for biodiesel production.

Microalgae are the most primitive types of plants that lack stems, roots, and leaves and belong to the thallophytes. Microalgae have a widespread presence in diverse aquatic environments, from fresh water to marine systems. For example, Chlorella, spirulina, and Scenedesmus are generally considered freshwater algae, while Dunaliella, Nannochloropsis, and Isochrysis are some microalgae species that grow well in marine water [2]. Furthermore, microalgae are considered photosynthesis-driven factories that use light to transform carbon dioxide into specific compounds such as carbohydrates, proteins, lipids, vitamins, and pigments [3,4]. However, the growth rate and biochemical composition of microalgae vary between species and are generally influenced by cultivation conditions such as pH, temperature, and nutrient availability [5,6]. Like terrestrial plants, microalgae also need N, P, K, and some micronutrients (like Fe, B, and Co) for their growth.

One of the key advantages of microalgae is its ability to grow in wastewater year-round in open ponds or photobioreactors and higher carbon sequestration compared to terrestrial plants. According to literature, 1 kg of algae can fix 1.8 kg of carbon dioxide [7]. Furthermore, microalgae can produce 10–30 times more biodiesel per hectare per year than terrestrial crops such as jatropha or soybean [8,9,10]. There are numerous industrial uses for microalgae. Certain microalgae, such as chlorella and spirulina, are high in protein and are used as dietary supplements [11]. On the other hand, microalgae Chlorella Dunaliella, Nannochloropsis, and Isochrysis can be utilized to produce a range of biofuels, including jet fuel, biodiesel, bioethanol, biogas, and biohydrogen depending upon biochemical composition [12,13,14,15] (Figure 1). High lipid content (between 20% and 80%), their ability to survive year-round even in wastewater, their high biomass productivity (doubling times of up to 24 h), and higher photosynthetic efficiency are some characteristics that make microalgae well-suited feedstock for biodiesel production [16,17,18,19]. Despite these advantages, several experiments aimed at commercializing microalgae-based biodiesel face significant challenges, primarily the high cultivation, harvesting, and lipid extraction costs [20]. Another significant challenge in microalgae biodiesel production is scaling up to meet market demands. The use of fresh water and nutrient availability during cultivation are some sustainability concerns. Hence, it is essential to address these challenges to enhance the sustainability and economic viability of microalgae biodiesel. Microalgae-derived biodiesel has a bright future ahead of it despite these obstacles. The full potential of microalgae biodiesel may be achieved using sustainable practices such as improved cultivation systems, genetic engineering, and integration with industrial waste streams [21].

Exploring resilient microalgae strains that can easily adapt to diverse conditions and utilize waste resources such as carbon dioxide and industrial wastewater for cultivation could greatly reduce costs and increase environmental benefits [21,22]. Hence, this manuscript reviews the strategies to optimize the higher lipid accumulation in microalgae biomass along with advances in cultivation, harvesting, and transesterification processes. Additionally, storage stability is a concern in microalgae-based biodiesel industries, as oxidation over time leads to increased viscosity due to gum formation. This paper also explores strategies to improve the storage stability of algae biodiesel.

2. Biochemical Composition and Lipid Synthesis Pathway in Microalgae

Microalgae are tiny photosynthetic organisms, varying in size from micrometers to millimeters, depending on the species. Microalgae can be classified as autotrophic, heterotrophic, or mixotrophic based on their carbon utilization strategies. Autotrophic microalgae use inorganic carbon sources, such as CO₂ and sunlight (Figure 2), for photosynthesis, while heterotrophic microalgae rely on organic carbon sources like glucose. Mixotrophic microalgae can utilize both inorganic and organic carbon sources [7].

Microalgae biomass can be either highly protein-rich or highly lipid-rich, or it can have a balanced composition of carbohydrates, proteins, and lipids. Using fundamental nutritional requirements as a basis, Chisty approximates the molecular formula for the microalgal biomass to be CO_0.48H_1.83N_0.11P_0.01 [23]. Microalgae are composed of proteins, lipids, and carbohydrates; the ratio of each component changes based on the type of algae. The broad cell content, elemental composition, and energy properties of these important components are listed in Table 1.

Carbohydrate: Carbohydrate-rich microalgae offer a promising source for producing bioethanol, biobutanol, and biohydrogen. Different classes of microalgae produce specific types of polysaccharides, which serve as energy reserves. For instance, green microalgae synthesize starch, composed of both amylose and amylopectin. The green alga Tetraselmis suecica can accumulate between 11% and 47% of its dry weight as starch, depending on nutrient availability [24]. Microalgae-derived carbohydrates are primarily starch and cellulose, without any lignin content. Certain species, such as Tetraselmis, Dunaliella, Scenedesmus, Chlorella, and Chlamydomonas, have been found to store significant amounts of carbohydrates, often exceeding 40% of their dry weight [25].

Protein: Proteins are highly valuable, particularly for use as animal feed, because the human body cannot synthesize certain essential amino acids [26]. Over the past few decades, more than 75% of the annual microalgal biomass production has been dedicated to the health food industry, where it is commonly processed into powders, pills, and capsules. Spirulina (a type of cyanobacterium) is widely cultivated worldwide as a food and dietary supplement. This filamentous blue-green alga is available in various forms, including pills, flakes, and powders [27].

Table 1. Elemental composition of microalgal biochemical components [28,29,30,31,32,33,34].

S. No.	Strain/Microalgae Biomass	Biochemical Composition			Elemental Composition
		Carbohydrate	Protein	Lipid	Carbon	Hydrogen g	Nitrogen	Sulfur	Oxygen
1	T. dimorphous	26.15	36.63	28.33	49.94	7.58	6.60	0.52	35.35
2	T. obliquus	27.94	46.37	18.33	49.31	7.46	7.01	0.55	35.67
3	Chlorella sp.	34.61	31.13	21.66	53.91	7.71	6.01	0.43	31.95
4	C. sorokiniana	36.56	39.38	15.83	52.80	7.74	5.32	0.17	33.79
5	P. cruentum	13.89	42.90	14.67	-	-	-	-	-
6	M. coccoides	40.65	20.65	17	35.55	6.01	5.67	-	-
7	Desmodesmus sp.	-	18.6	38.8	25.8	5.4	6.2	0.6	-
8	Nanofrustulum sp. 2	-	12..52	12.52	27.45	4.3	2.90	0.96	-
9	Scenedesmus sp.	-	-	50.6	43.8	5.7	8.1	-	-
10	Chroococcidiopsis sp.	45.40	36.72	5.6	-	-	-	-	-

Table 1. Elemental composition of microalgal biochemical components [28,29,30,31,32,33,34].

S. No.	Strain/Microalgae Biomass	Biochemical Composition			Elemental Composition
		Carbohydrate	Protein	Lipid	Carbon	Hydrogen g	Nitrogen	Sulfur	Oxygen
1	T. dimorphous	26.15	36.63	28.33	49.94	7.58	6.60	0.52	35.35
2	T. obliquus	27.94	46.37	18.33	49.31	7.46	7.01	0.55	35.67
3	Chlorella sp.	34.61	31.13	21.66	53.91	7.71	6.01	0.43	31.95
4	C. sorokiniana	36.56	39.38	15.83	52.80	7.74	5.32	0.17	33.79
5	P. cruentum	13.89	42.90	14.67	-	-	-	-	-
6	M. coccoides	40.65	20.65	17	35.55	6.01	5.67	-	-
7	Desmodesmus sp.	-	18.6	38.8	25.8	5.4	6.2	0.6	-
8	Nanofrustulum sp. 2	-	12..52	12.52	27.45	4.3	2.90	0.96	-
9	Scenedesmus sp.	-	-	50.6	43.8	5.7	8.1	-	-
10	Chroococcidiopsis sp.	45.40	36.72	5.6	-	-	-	-	-

Lipid: Lipids are diverse biomolecules that can be classified into various categories, including fatty acids, glycerophospholipids, glycerolipids, sterol lipids, sphingolipids, saccharolipids, prenol lipids, and polyketides [35]. These lipids encompass a range of molecules, such as fats, sterols, waxes, monoglycerides, diglycerides, triglycerides, phospholipids, and fat-soluble vitamins. Lipids, like carbohydrates, are structural elements of cell membranes as well as energy stores. Simple fatty acid triglycerides are important as energy reserves. Various microalgal species are known to accumulate significant amounts of lipids. For example, species like Botryococcus braunii, Dunaliella tertiolecta, Nannochloropsis sp., Chlorella emersonii, Porphyridium cruentum, and Neochloris oleoabundans have been found to have a lipid content exceeding 60% of their dry weight [36]. Given the focus of this study on microalgae biodiesel production, a detailed discussion on the biosynthesis of lipids in microalgae follows.

3. Factors Responsible for Enhancing Lipid Production in Microalgae

Numerous studies have demonstrated the strong influence of various environmental factors on lipid content and composition, including temperature, light intensity, cell culture density, pH, alkalinity, contamination by other microorganisms, and composition of nutrient media (concentration of nitrogen, phosphate, and iron). Some of them are discussed below.

3.1. Light

Light is the primary energy source for microalgae growth, making its availability and intensity crucial factors in the efficiency of microalgae biomass production. At very low light intensity, the average growth rate of microalgae is zero, a point known as the compensation point. As light intensity increases, the growth of microalgae accelerates until it reaches the saturation point, which represents the peak of photosynthetic activity) [37,38,39]. Beyond this point, further increases in light intensity can lead to a decline in growth rate and may cause photo-oxidation, damaging light receptors and reducing photosynthetic efficiency and productivity, a phenomenon known as photo-inhibition [40]. Additionally, the chemical composition, pigment concentration, and photosynthetic activity of microalgae can vary significantly under different light intensities. Generally, high light intensity leads to a reduction in polar lipid content while increasing the amount of neutral storage lipids, primarily triacylglycerols (TAGs). Conversely, low light intensity tends to promote the production of polar lipids, particularly those associated with the chloroplast membrane [39,41,42,43].

In addition to this, the photoperiod and light wavelength are crucial factors for algal growth and lipid accumulation. Some experimental results showed that microalgae cultivation could be performed under different dark–light periods, such as 12:12, 14:10, 10:14, 16:8, 0:24, and 24:0, to give rise to higher TGA accumulation. Under dark conditions, biomass productivity downfalls, and sometimes the losses might reach as high as 25%. Since there is no sunlight throughout the night, dark respiration causes a loss of biomass, which results in an overall drop in biomass [44]. Light from artificial light sources, solar light, or a combination of light sources can all be used to illuminate a microalgae growth system. LED, cool fluorescent, and light bulbs are examples of artificial lighting [37,45,46]. However, these electricity-consuming sources raise the cost of producing total algal biomass. On Earth, solar energy is the most economical light source available. Better technologies are required to complement the sunlight employed in most commercial algal production systems in order to accelerate the growth of microalgae outside.

3.2. Temperature

Temperature is one of the most significant environmental factors influencing the rate of growth and biochemical composition of microalgae. Although microalgae can be produced in a range of temperatures (generally 20–30 °C), only a few specific temperatures are optimal for each strain of algae. Thus, the ideal temperature for biomass generation depends on the genus and strain. Prolonged temperatures above the optimal range can damage algae cells, while temperatures below the ideal range result in slower development [47]. In addition, the temperature fluctuations, both daily and seasonal, affect the rate of algae growth. Moreover, a lower temperature seems to reduce the amount of biomass lost by respiration at night. Temperature has a major impact on the oxygen-evolving activity of Photosystem II (PSII) [48]. There is a substantial correlation between temperature, light, and photoinhibition. High light intensities cause photo-inhibition at low temperatures. Considering this, one of the key variables restricting outdoor cultivation in the winter is temperature. Nonetheless, increasing the temperature can greatly lessen photoinhibition [48,49].

Algal fatty acid composition is significantly influenced by temperature. Numerous algae and cyanobacteria have a general trend that increases fatty acid unsaturation with decreasing temperature and increases saturated fatty acids with rising temperature [47,48,49]. For example, Visentin et al. 2024 [50], focused on the generation of value-added bioproducts by cultivating Phormidium autumnale in sugarcane vinasse at several temperatures and assessing the removal of contaminants. At 35 °C or 127.20 mg/L/d, with a pH of 7.5 and a C/N ratio of 16, the maximum biomass production was attained. Organic carbon, nitrogen, and phosphorus were removed at rates of 49%, 47%, and 28%, respectively [50]. In another study, temperature-acclimated cultures of Nannochloropsis oceanica were obtained using the turbidostat method [51]. The ideal temperature range for growth was found to be 25–29 °C, while temperatures above 31 °C and below 9 °C entirely stopped growth. However, some microalgae species (e.g., Chlorella vulagaris) have shown good potential to survive even in a high-temperature range of 40 °C [52]. A notable metabolic shift occurred in response to temperature stress, with triacylglycerol concentrations increasing at 17 °C and decreasing at 9 °C. Despite these changes in lipid content, the levels of total and polar eicosapentaenoic acid (EPA) remained consistent at 3.5% and 2.4% w/w, respectively.

In addition, daily fluctuations and seasonal temperature changes can significantly affect the development of microalgae, often limiting their cultivation to specific times of the year, depending on the species’ thermotolerance. Even brief temperature fluctuations throughout the day can have a substantial impact on biomass productivity and lipid quality and quantity. The productivity of lipid and biomass has a similar trend, peaking between 21 and 25 °C [48,50,51].

3.3. Growth Nutrients

Microalgae need several nutrients to grow. Carbon, nitrogen, and phosphorous are considered essential elements, although calcium, sulfur, iron, potassium, sodium, hydrogen, manganese, magnesium, cobalt, and sulfur are used as trace nutrients [35,53]. In the next section, the effects of carbon, nitrogen, and phosphorus on microalgae development and lipid production are discussed:

(a) Carbon: Microalgae require carbon for their growth, which can be sourced either organically or inorganically. These organisms use fixed carbon for respiration, energy production, and cell growth. Autotrophic microalgae rely on inorganic carbon sources like carbonate and bicarbonate, whereas heterotrophic microalgae utilize organic carbon sources such as glucose, glycerol, and sodium acetate [54,55].

The ability to use bicarbonate ions (HCO₃⁻) and CO₂ as inorganic carbon sources is unique to microalgae. Bicarbonate, often stored intracellularly, is converted to CO₂ by the enzyme carbonic anhydrase (CA). The bicarbonate–carbonate buffer system is a weak acid/base buffer present in water with CO₂. The development of different inorganic carbon species is influenced by temperature and pH levels. Bicarbonate (HCO₃⁻) dominates at pH levels up to 10.5, while carbonate (CO₃⁻²) becomes more prevalent at higher pH values. This buffer system provides the necessary carbon for photosynthesis, with the pH, temperature, and nutrient content of the water influencing the available form of CO₂ [56].

CO₂ + H₂O ⇌ H₂CO₃ ⇌ H⁺ + HCO₃⁻ ⇌ 2H⁺ + CO₃ ²⁻

(1)

However, the dissolution of CO₂ in water can lead to acidification due to the formation of carbonic acid. To mitigate this issue, a high concentration of inoculum should be provided. At high inoculum rates, microalgal cells can efficiently utilize all available CO₂ for photosynthesis, preventing the reaction of CO₂ with water and thereby avoiding acidification [56].

CO₂ is a most important substrate for photosynthesis and plays a significant role in determining algal growth and fatty acid biosynthesis. Tetradesmus obliquus, Desmodesmus opoliensis, and Chlorella sp. have shown great promise as CO₂-to-fuel converters [57]. These species efficiently convert CO₂ into lipid-rich biomass suitable for biodiesel production. The best performance was observed at a CO₂ concentration of 20%, where all three strains demonstrated significant CO₂ capture capabilities, along with notable biomass production and lipid accumulation. The results showed that CO₂ fixation rates ranged from 0.020 to 0.072 g-CO₂/L/day, biomass productivity from 0.011 to 0.040 g/L/day, and lipid accumulation from 0.22 to 3.39 mg/L/day.

As shown in Table 2, certain microalgae can directly utilize organic carbon sources such as glucose, acetate, glycerol, fructose, sucrose, lactose, galactose, and mannose as substrates and energy sources [15,54,58,59]. These microalgae are referred to as heterotrophs. Research shows that species like Chlorella vulgaris, Scenedesmus acutus, C. protothecoides, Chlorella saccharophila, and Chlorella sorokiniana can be grown using heterotrophic methods [60]. However, the cost of these carbon sources is a significant challenge that could limit the scalability of this technology for industrial applications. Additionally, contamination presents another concern in heterotrophic cultivation systems.

(b) Nitrogen: Nitrogen is also a crucial nutrient for microalgae growth, as it controls protein synthesis and the production of growth metabolites. The nitrogen content in microalgae biomass can range from 1% to over 10%, depending on the availability, amount, and type of nitrogen source [61]. Microalgae can utilize nitrogen in the forms of nitrate, nitrite, ammonia, and urea. When ammonia is available, it is the preferred nitrogen source, and microalgae will not utilize other sources until the ammonia is depleted [62]. However, the high uptake of ammonia can lower the medium’s pH to below 6.0, a level at which microalgae cannot grow. The uptake of ammonia also leads to the production of hydrogen ions (H⁺), further contributing to the acidification of the medium (Shown in Equation (2)).

NH₄⁺ + 7.6 CO₂ + 17.7H₂O → C_7.6H_8.1O_2.5N_1.0 + 7.6 O₂ +15.2H₂O +H⁺

(2)

The intake of nitrogen causes pH variations that impact the growth rate of microalgae cells. Nitrate absorption raises pH because it produces OH⁻. Theoretically, digesting one mole of nitrate produces one mole of OH⁻, as indicated by Equation (Shown in Equation (3).

NO⁻₃ + 5.7CO₂ + 5.4H₂O → C_5.7H_9.8O_2:3N_1.0 + 8.25O₂ + OH⁻

(3)

During the cultivation phase, when microalgae experience nitrogen deficiency, it is shown that this promotes the biosynthesis and accumulation of lipids and triglycerides [39,49,62,63,64]. A study investigated the effects of nitrogen starvation on the fatty acid profile, biochemical composition, and growth responses of Nannochloropsis oculata, Phaeodactylum tricornutum, and Dunaliella tertiolecta [65]. All three microalgae species exhibited changes in their lipid, fatty acid, protein, and carbohydrate compositions in response to nitrogen deficiency. Under nitrogen stress, P. tricornutum showed a significant increase in lipid content by 139%, while D. tertiolecta experienced a 59% increase in carbohydrate content compared to the control groups. Hence, nitrogen deficiency is considered an effective method for enhancing lipid accumulation in microalgae. However, while nitrogen stress can significantly increase lipid production, it also leads to slower growth rates. This reduction in growth rate can ultimately impact overall biomass and lipid productivity.

3.4. Phosphorous

Phosphorus is another crucial macronutrient for the regular growth and development of microalgae, as it plays a key role in cellular metabolic activities by forming various structural and functional components [66]. The preferred method for providing phosphorus to microalgae is through orthophosphate. Despite constituting only about 1% of the microalgal biomass, phosphorus is a significant growth-limiting element in microalgae biotechnology. The availability of phosphorus also greatly affects biomass composition, particularly influencing the accumulation of fats and carbohydrates [66,67]. The ideal N/P ratio for microalgae is 20:1. In the stationary phase, microalgae require a lower N/P value, and in the exponential phase, a higher one. In the exponential phase, high N/P promotes the formation of more microalgae biomass. In the stationary phase, low N/P encourages the accumulation of additional lipids [68]. Optimizing N/P ratios during both the stationary and exponential growth phases can lead to a two- to three-fold increase in lipid production compared to using the same ratios throughout both phases. Additionally, the availability of CO₂ significantly affects the absorption of nutrients such as phosphorus and nitrogen. To maximize biomass and lipid production, it is crucial to optimize the ratios of carbon to nitrogen and nitrogen to phosphorus.

3.5. Trace Metals

Essential trace metals that microalgae need for various kinds of metabolic activities include copper (Cu), cobalt (Co), iron (Fe), manganese (Mn), nickel (Ni), and zinc (Zn) [69]. Several studies have been carried out to see the impact of different metals on lipid accumulation in algae [69,70,71]. While high metal concentrations are harmful to growth, deficiencies in trace metals slow down the rate of growth. In addition to serving as a redox catalyst during photosynthesis and nitrogen absorption and mediating electron transport pathways in photosynthetic organisms, iron is a crucial trace element for healthy growth.

3.6. Cell Density and Mixing

The cell density of a culture system significantly impacts the growth of microalgae. Maintaining optimal levels of light intensity and penetration is crucial and depends on the specific strain being cultivated. Both high and low cell concentrations can reduce biomass productivity. When cell density is low, insufficient light absorption occurs because unabsorbed light passes through the culture without being utilized by the cells. Conversely, if the cell concentration is too high, light limitation creates a ‘dark zone’ within the culture, reducing photosynthetic activity. At high cell densities, mutual shading occurs, leading to increased turbidity, which further reduces the light intensity, photosynthetic efficiency, and, ultimately, biomass yield [72].

The chosen mixing system plays a crucial role in both the construction and operational costs of photobioreactors, as well as the overall efficiency of microalgae culture systems. Optimal mixing or turbulence serves multiple purposes, such as preventing cell settling, uniformly distributing nutrients, CO₂, and temperature, removing oxygen produced during photosynthesis, and enhancing effective light utilization [73]. Insufficient mixing can create a dark, stagnant zone in the microalgae suspension, negatively impacting growth. On the other hand, excessive mixing can cause shear stress, which may damage the cells. Various mixing techniques are employed, including mechanical stirring, gas injection, and pumping. Among these, mechanical stirring is the most effective, but it also results in higher hydrodynamic stress compared to other methods [73].

3.7. Genetic and Metabolic Regulation

Environmental conditions can cause microalgal wild types to acquire lipids, but they often impair their biomass productivity. One potential answer to this problem is genetic alteration, which could allow for the cultivation of microalgae strains with higher lipid productivity without sacrificing biomass growth. The DNA of the microalgae, which supplies the innate properties needed to build the cell, is the central idea of this approach. Upon identifying a desired characteristic in a microalgae strain’s DNA, the feature is separated and combined with plasmid DNA containing an enzyme to create a new plasmid, which is then introduced into a specific microalgal cell [74]. At the molecular level, this genetic engineering procedure can be carried out via a few techniques referred to as genome editing systems. An illustration of this is the synthetic enzyme Transcription Activator-Like Effector Nuclease (TALEN), which cleaves the desired DNA sequence so that the new plasmid can use it [75]. In the research of Takahashi et al. [76], TALEN was utilized to produce mutants of Coccomyxa sp. with changed DNA sequences, resulting in starch-free microalgae cultures. Chang et al. [77] used this process to create two novel strains of Tetraselmis sp. with elevated lipid levels (from 7.8% to 21.1% and 24.1%, respectively) caused by a mutation in the strain’s ADP-glucose pyrophosphorylase protein [77]. CRISPR techniques have advanced to the point where Jeon et al. [78] classified the systems into two classes: Class I CRISPR systems refer to effectors made of multi-subunit proteins, while Class II CRISPR systems contain a single effector with multiple domains. The class II CRISPR system is defined by the method previously described using the Cas9 protein. RNA interference (RNAi) is another technique that has advanced recently. The cleaved segments of microRNA also referred to as short interfering RNA, are coupled to a molecule that contains an RNA-induced silencing complex. These segments are broken down by the enzyme DICER. By using genetic engineering to access new microalgal species and studying their unique traits, more process improvement can be accomplished.

Table 2. Biodiesel yield from different microalgae grown under various cultivation conditions.

Microalgae	Cultivation Mode	Cultivation Conditions	Bioreactor/Open Ponds	Biomass Productivity	Lipid Productivity	Biodiesel Yield	Ref.
Spirulina		Growth medium—ASN III Temp—25–27 °C, Time—14 days, Light intensity—94.5 μmol−2 s−1	PBR	-	-	99%	[79]
Golenkinia sp. SDEC-16s algae	Autotrophic	Growth medium—seawater supplemented with monosodium glutamate wastewater Time—10 days	PBR	0.26 g/L/d	98.99 mg/L/d	98%	[80]
Chlorella sp.	Heterotrophic	Growth medium—BBM Carbon source—Glycerol (4 g/L)	PBR	446.50 mg/L/d	165.15 mg/L/d	>20%	[81]
Chlorella vulgaris ESP	Autotrophic	Growth medium—Basal Carbon source −2% CO2, Artificial light	PBR	250 mg/L/d	56.2 mg/L/d
Scenedesmus obliquus	Heterotrophic	Growth medium—BG-11 Carbon source—acetate	EMF	92.3	56	60.56%	[82,83]
Chlorella vulgaris	Autotrophic	Growth medium—Commercial fertilizer Outdoor conditions, pH = 7	Open raceway ponds	77 mg/L/d	11.52 mg/L/d	84.01%	[6]
Chlorella minutissima	Autotrophic	Growth medium—Commercial fertilizer Outdoor conditions, pH = 7	Open raceway ponds	162.0 mg/L/day	31.43 mg/L/day	90.21%	[5]
Ettlia sp. YC001	Mixotrophic	Growth medium—BG-11 Carbon source—2% CO2–mixed air + Glucose; Artificial light	PBR	410 mg/L/day	-	15.5%	[84]
Chlorella protothecoides	Heterotrophic	Growth medium—BG-11 Carbon source—Glucose	EMF	1656.25 mg/L/day	687.5 mg/L/day	73.2%	[85]
Chlorella pyrenoidosa	Heterotrophic	glucose and B.Peptone concentrations—~13–24 g/L and 1.3 ± 0.1 g/L	stir-tank bioreactor	-	-	76%	[86]

Table 2. Biodiesel yield from different microalgae grown under various cultivation conditions.

Microalgae	Cultivation Mode	Cultivation Conditions	Bioreactor/Open Ponds	Biomass Productivity	Lipid Productivity	Biodiesel Yield	Ref.
Spirulina		Growth medium—ASN III Temp—25–27 °C, Time—14 days, Light intensity—94.5 μmol−2 s−1	PBR	-	-	99%	[79]
Golenkinia sp. SDEC-16s algae	Autotrophic	Growth medium—seawater supplemented with monosodium glutamate wastewater Time—10 days	PBR	0.26 g/L/d	98.99 mg/L/d	98%	[80]
Chlorella sp.	Heterotrophic	Growth medium—BBM Carbon source—Glycerol (4 g/L)	PBR	446.50 mg/L/d	165.15 mg/L/d	>20%	[81]
Chlorella vulgaris ESP	Autotrophic	Growth medium—Basal Carbon source −2% CO2, Artificial light	PBR	250 mg/L/d	56.2 mg/L/d
Scenedesmus obliquus	Heterotrophic	Growth medium—BG-11 Carbon source—acetate	EMF	92.3	56	60.56%	[82,83]
Chlorella vulgaris	Autotrophic	Growth medium—Commercial fertilizer Outdoor conditions, pH = 7	Open raceway ponds	77 mg/L/d	11.52 mg/L/d	84.01%	[6]
Chlorella minutissima	Autotrophic	Growth medium—Commercial fertilizer Outdoor conditions, pH = 7	Open raceway ponds	162.0 mg/L/day	31.43 mg/L/day	90.21%	[5]
Ettlia sp. YC001	Mixotrophic	Growth medium—BG-11 Carbon source—2% CO2–mixed air + Glucose; Artificial light	PBR	410 mg/L/day	-	15.5%	[84]
Chlorella protothecoides	Heterotrophic	Growth medium—BG-11 Carbon source—Glucose	EMF	1656.25 mg/L/day	687.5 mg/L/day	73.2%	[85]
Chlorella pyrenoidosa	Heterotrophic	glucose and B.Peptone concentrations—~13–24 g/L and 1.3 ± 0.1 g/L	stir-tank bioreactor	-	-	76%	[86]

4. Upstream Process for Biodiesel Production

The upstream process in microalgae-based biodiesel production includes cultivation and harvesting techniques.

4.1. Growth Techniques

Microalgae can be grown in photoautotrophic, heterotrophic, and mixotrophic environments since they can thrive on both organic and inorganic substrates. Two cultivation methods are typically the most widely used ones: closed photobioreactors and open pond systems.

4.1.1. Open Ponds System

Popular open-pond systems for microalgae cultivation include natural ponds, circular ponds, raceway ponds, and inclined systems (Figure 3). These ponds are typically constructed with clay or concrete and coated with materials such as porcelain tiles or polyvinyl chloride to minimize nutrient and media loss. To ensure optimal sunlight penetration, the pond depth is usually kept between 0.15 and 0.45 m [5,87]. Air pumps or paddlewheels are used for mixing and circulation. These systems are cost-effective and have simple designs, making them easier to manage, though they require careful monitoring to control water and nutrient loss.

Among these systems, raceway ponds are the most used for outdoor microalgae production. However, they are affected by weather patterns and lack precise control over parameters such as lighting and temperature. As a result, microalgae productivity can vary significantly with seasonal changes. The production of microalgal biomass in open ponds is heavily influenced by local climate conditions. Additionally, open-pond systems face several challenges: (1) poor mixing, (2) low mass transfer rates, (3) low biomass productivity, (4) significant water loss through evaporation, and (5) difficulties in maintaining microalgae monocultures due to contamination from bacteria and other algal strains [88].

Figure 3. Different types of ponds for microalgae culture: (a) open raceway ponds and (b) circular ponds [89,90] (license no: 5959200070726).

Figure 3. Different types of ponds for microalgae culture: (a) open raceway ponds and (b) circular ponds [89,90] (license no: 5959200070726).

4.1.2. Closed System

Photobioreactors (PBRs) are closed systems that allow light and energy exchange but not material interaction with the surrounding atmosphere. As a result, they allow the long-term, low-contamination culture of single species of microalgae [23]. As shown in Figure 4, various kinds of photobioreactors have been developed to achieve optimal productivity. Of them, tubular, flat plate, and column PBRs are the three most common types [91]. Mixing within these PBRs usually takes place by airlift or mechanically stirring/pumping. Mixing is also necessary for gas exchange. PBRs require less space than open ponds for the purpose of producing the same quantity of biomass. PBRs can utilize natural light, artificial light, or a variety of combinations of light sources, making them more adaptable than open ponds. This allows them to potentially boost photoperiod and enhance low light intensities caused by variations in sunshine [91].

Tubular PBRs are among the best options for large-scale outdoor production because they provide a vast surface area that is exposed to light and can be utilized for both indoor and outdoor growing [91]. Transparent glass or plastic solar collector tubes used in photobioreactors (PBRs) typically have a diameter of just over 0.1 m, as light penetration is limited by the density of the culture broth [23]. Due to their high culture densities and large surface area exposed to light, flat plate PBRs have garnered significant research interest in recent years [92,93]. These PBRs are particularly well-suited for large-scale cultivation, offering high photosynthetic efficiency and minimal accumulation of dissolved oxygen compared to tubular PBRs. Despite their advantages, scaling up these PBRs remains challenging. PBRs outperform open ponds primarily due to their increased biomass productivity and enhanced management capabilities. They offer the opportunity to optimize light exposure in terms of duration, intensity, and light wavelength. The capital and operating expenditures are the main barriers to using PBRs for large-scale microalgae culture [91,94]. Moreover, regular cleaning and maintenance of Photobioreactors (PBRs) are essential because biofilm formation on the walls can obstruct light penetration, thereby reducing photosynthetic efficiency. Consequently, the development of effective PBRs that minimize construction and operational costs is crucial for large-scale microalgae cultivation in biotechnologies.

Building-integrated photobioreactors are gaining popularity to encourage environmentally friendly construction. Buildings powered by algae have the potential to influence public perceptions regarding the value and potential of green building architecture. The BIQ building, the Algae Green Loop Tower, and the Process Zero Concept Building are the three microalgae-powered structures that exist today [95]. The aim behind these designs is to use building wastewater and atmospheric carbon dioxide to produce microalgae biomass while combining the idea of green buildings. However, before it is commercialized, this concept needs more research on cost-cutting measures because it is highly expensive [95].

C. Hybrid systems: In hybrid systems, both raceway ponds and photobioreactors are employed in a two-stage microalgae cultivation process to leverage the strengths of each method and mitigate their respective drawbacks [96]. The photobioreactor is used as the first stage, where it helps reduce contamination and achieve a high concentration of biomass. The second stage takes place in raceway ponds, where microalgae are exposed to nutritional stressors to enhance lipid production. Depending on local temperature conditions, hybrid systems can yield up to 20–30 tons of lipids per hectare annually. The benefits of hybrid systems include higher productivity, reduce energy consumption, and improved control over cultivation parameters. However, the primary drawback is the increased capital and maintenance costs compared to non-hybrid reactors [97].

4.2. Utilization of Wastewater for Microalgae Cultivation

The use of wastewater for microalgae cultivation has recently gained significant attention as a cost-effective and environmentally friendly approach for producing microalgae-based biorefineries [98]. Microalgae can utilize the nutrients, such as nitrogen (N) and phosphorus (P), present in wastewater for growth while simultaneously breaking down undesirable and hazardous substances [99]. Traditional algal cultivation systems often require the addition of specific nutrients like N, P, and micronutrients, which increases the overall cost of cultivation. In contrast, using wastewater, where these nutrients are already present, can significantly reduce the cost of producing microalgae biomass [98]. Bora et al. [100] used Chlorella sorokiniana KP to conduct experiments on the production of microalgal biodiesel, the generation of bioelectricity, and the treatment of wastewater. Rice mill wastewater (RMWW) and dairy wastewater (DWW) were used as substrates for optimization studies on the production of biodiesel and bioelectricity. Better biodiesel yield was obtained in DWW by Chlorella sorokiniana KP than in RMWW; hence, DWW was used as a substrate for more integrated bioelectricity investigations. In one study, the nitrogen removal rate was evaluated by growing Chlorella vulgaris in different water-to-urine dilution ratios [101]. The Bligh and Dyer method was employed to extract biodiesel from the Chlorella vulgaris algae (CAB), which was then trans esterified and analyzed for its elemental and functional groups. In another study, the microalga Tetradesmus obliquus (Turpin) Kützing was utilized to treat industrial wastewater from paper pulp, including both primary-treated (PTW) and secondary-treated (STW) wastewater [102]. A batch culture study revealed that adding 50% secondary-treated wastewater could enhance biomass yield by 1.8 times, achieving 2.51 g/L in 15 days while also producing 390.2 mg/L of lipids. When this approach was scaled up to 200 L open raceway ponds using 50% STW as the growth medium, the results showed lipid productivity of 26.80 g and areal biomass productivity of 3.90 g.

Integrating wastewater treatment with algae cultivation offers significant environmental and economic benefits. This approach allows for the organic treatment of wastewater without relying on chemicals, providing essential nutrients for algae growth [98]. The resulting biofuel and treated wastewater are environmentally friendly and pose no harm. Additionally, this integration reduces the workload for those managing these processes, making it a more efficient and sustainable solution [103].

4.3. Microalgae Harvesting

After cultivation, harvesting is the key process for removing microalgae biomass from the growing medium. Typically, 2–7% of the slurry contains microalgae cells post-harvest, and harvesting costs account for approximately 25–35% of the total microalgae biomass production expenses [104]. The ideal harvesting method depends on factors such as the type of microalgae, cell density, and growth conditions [17]. The four primary methods for harvesting microalgae include flotation, centrifugation, filtration, electrophoresis, and sedimentation. The choice of harvesting technique should be guided by the intended final product.

Gravity sedimentation can be employed for low-value products, with enhanced sedimentation through flocculation or the use of settling ponds, particularly for harvesting biomass grown in sewage [105]. For high-value products, such as those used in food or aquaculture applications, centrifugation is recommended. However, not all microalgae species are suitable for every harvesting method; for instance, centrifugation is versatile and can be used with any type of microalgae, while filtration is more appropriate for specific species like Scenedesmus platensis [17,106]. To optimize biomass recovery, harvesting can be combined with mechanical dewatering—using centrifugation for pre-concentration and screw centrifugation or heat drying for post-concentration. Additionally, economic considerations should play a key role in selecting the appropriate biomass collection method.

Centrifugation is a widely used technique for harvesting microalgae, providing efficient separation of microalgal cells. Typically, centrifugation can achieve a harvesting yield of approximately 80–90% within two to five minutes of operation [106]. It is particularly effective for large-scale harvesting, offering superior recovery and producing a concentrated slurry. However, the cost of centrifugation equipment is high, and the process can be economically unfeasible due to significant energy consumption and large capital investment. Additionally, the high shear forces and gravitational stress involved may damage the cell structure, impacting the quality of the harvested microalgae.

Flocculation is another technique that is mostly used for microalgae harvesting [107]. Generally, before using additional harvesting techniques like filtration floatation or gravity sedimentation, flocculation can be used independently or in conjunction with them as a pre-concentration step [108]. Cells combine during the first phase of flocculation to enlarge the size of the particles. The spontaneous aggregation of negatively charged cells in suspension is prevented by the negative charges on the surface of microalgal cells. This can be counteracted by adding multivalent cations and cationic polymers, such as ferric chloride, aluminum sulfate, etc. Consequently, allowing microalgae to self-aggregate has been explored as an alternative harvesting method. Inorganic flocculants used for this purpose include Fe₂(SO₄)₃, FeCl₃, Al₂(SO₄)₃, AlCl₃, ZnSO₄, ZnCl₂, CaSO₄, CaCl₂, MgSO₄, MgCl₂, (NH₄)₂SO₄, and NH₄Cl [104,107]. However, the high concentration of metal flocculants used in biomass harvesting can negatively impact the biodiesel production process and pose potential hazards. As a result, research is being directed towards organic polymers as potential substitutes for inorganic flocculants. Some of these organic alternatives include chitosan, cationic starch, and grafted starch, which offer higher harvesting efficiency and leave the biomass less contaminated.

5. Downstream Processes

The downstream process includes lipid extraction and transesterification process. Generally, this involves a two-step process: in the first step, the lipid is extracted from the microalgae, followed by transesterification in the second step to produce biodiesel. However, some recent studies showed that dry or wet microalgae can be employed directly for biodiesel production through a single-step process known as in situ transesterification. This approach combines lipid extraction and transesterification into a simultaneous process, streamlining production and improving the efficiency of the process. Some examples of microalgae-based biodiesel production are shown in

Table 3. Biodiesel production from different microalgae using different catalysts.

Microalgae	Cultivation Conditions	Reaction Conditions					Yield (%)	Ref.
		Catalyst Type	Catalyst	Alcohol: Oil (mol/mol)	Temperature (°C)	Time (h)
Nannochloropsis	Artificial growth medium -	Hydrotalcite catalyst	CaO	1:1 Hexane/ Methanol	60 °C	4 h	Olefine (61.49%) Gasoline (38.51%)	[109]
Tetraselmis indica	Wastewater growth medium 16:8 light/dark period beneath the light intensity of 94.5 μmol m−2 s−1 for 14 days	Nanocatalyst	Lithium-impregnated calcium oxide	-	-	-	87.25%	[110]
Spirulina sp.	Artificial Temperature: (25–27 °C) Time: 14 days Light intensity: 94.5 μ/mol/s 16:08 dark and light cycle	Nanocatalyst	Ca(OCH)2	30:1 M/O	80 °C	3 h	99%	[79]
Nannochloropsis sp.	-	Composite catalysts		2:1 Dichloromethane and methanol	80 °C	4 h	~80.0%	[111]
Chlorella sorokiniana	Artificial growth medium 25 ± = 2 °C (300 µmol m−2 s−1)	Magnetic nanoparticles	(Fe3O4)	-	350.0 °C	45 min	28.0%	[112]
Chlorella pyrenoidosa	Artificial, fertilizer nutrient media Outdoor conditions (20–252 °C)	Homogenous	KOH	4:1	65.0 °C	7 min	15.5% of dry wt.	[18]
Chlorella minutissima	commercial fertilizer Outdoor conditions (20–252 °C)	Homogenous	KOH	10.5:1	63.9 °C	6 min	90.2%	[5]
Neochloris sp. SK57 and Chlorella sp. SL7A	Slaughterhouse wastewater as growth media 25 ± 2 °C and a day-night cycle of 11:13	Homogenous	2% H2SO4	-	100.0 °C	6 h	-	[113]
C.vulgaris	Textile industry wastewater as growth media 4600 lux, 30 ± 5 °C	Heterogenous	[HMIM][HSO4]/ Bio-MOF	15:1	70.0 °C	30 min	92.0%	[114]
Scenedesmus sp.	-	Ni dopped MgAl layered double hydroxides	NixMg2Al-LDH	-	300.0 °C	1 h	31.4%	[115]

.

5.1. Conventional Processes of Microalgae Biomass

Traditional methods include the extraction of lipids from microalgae, followed by transesterification.

5.1.1. Lipid Extraction from Microalgae

Since lipids are intracellular materials, processing them requires first removing them from the cells. Lipid extraction from microalgae can be broadly classified into two categories: mechanical and chemical approaches. Mechanical techniques, in general, result in substantial biomass losses and limited lipid selectivity. Mechanical pressing is commonly used for oil extraction from both edible and non-edible seeds such as Jatropha, Karanja, and Mustard [116]. Certain mechanical methods, on the other hand, limit the use of hazardous solvents while also shortening the processing time. Microalgae are unicellular strains that have thick cell walls that prevent intracellular lipids from being released, making them unsuitable for mechanical pressing [117].

The most popular technique to extract lipids from microalgae is chemical extraction. The chemical principle of “like dissolving like” serves as the foundation for the process of extracting oil from microalgae. Non-polar solvents play a crucial role in disrupting the hydrophobic interactions between neutral and non-polar lipids within microalgae biomass [117,118]. The Folch technique, named after Folch et al. [119] utilizes a 2:1 volumetric ratio of methanol to chloroform to extract lipids from animal tissues. Similarly, the Bligh and Dyer method employs a 1:12 v/v mixture of methanol and chloroform for the extraction and purification of total lipids. However, in the batch process of extraction, the lipid content from biomass into the solvent reaches a state of equilibrium with the lipid concentration in the solvent, which restricts further lipid transfer to the solvent. A continuous organic solvent extraction method could overcome this restriction, but it is costly because it uses a lot of organic solvents. Soxhlet equipment can play a major role in addressing these issues by replacing biomass with new organic solvents throughout cycles of solvent evaporation and condensation, reducing solvent use. Soxhlet apparatus is primarily made up of three parts: a condenser for continuous cooling and a Soxhlet extractor to store microalgae biomass in a round bottom flask that is continuously heated [118]. Rashd et al. [120] carried out lipid extraction of Scenedesmus parvus using the Soxhlet apparatus with a variation of temperature, time, and algae-to-solvent ratio and obtained 24% lipid content under optimized conditions of 70 °C temperature and 8 h time. Aravind et al. [121] extracted 26.86% algae oil from Spirogyra using hexane as a solvent and Soxhlet apparatus. However, the robust and complex structure of the microalgal cell wall, generally composed of proteins and polysaccharides, hides the complete extraction of lipids through the Soxhlet/chemical method. The extraction process is challenging because breaking down the resilient cell wall is essential to maximize microalgae biodiesel yield. Therefore, an appropriate pretreatment tailored to the specific type of cell is necessary before lipid extraction.

Bead pounding is one of the popular techniques to pretreat microalgae cells where algal cells are disrupted using high-speed mechanism techniques [117]. In this approach, a vertical or horizontal cylindrical compartment is filled to the required level with microalgae biomass and many fine high-velocity steel or glass beads and spun at a specific speed. The suspended cells are disturbed by compaction or shear pressures with energy transfer from the beads inside the compartment colliding at a higher velocity [117,118]. The process is highly influenced by factors such as the size of the beads, the temperature time, the number of cycles, and the spinning speed.

Ultrasonication and microwave-assisted techniques are also employed to improve lipid recovery from algal biomass. The energy of high-frequency acoustic waves creates dense micro bubbles in a liquid medium, which expand and collapse rapidly to generate the cavitation phenomena in ultrasonication, resulting in high temperature [120]. Selecting the optimal extraction technology for microalgae largely depends on the specific strain being used, like the way extraction methods are tailored for many higher plants.

Supercritical fluid extraction is emerging as a promising green technology that could potentially replace traditional solvent extraction methods. This technique is highly efficient and capable of extracting nearly all the oils present in the biomass [118,122]. The underlying principle of supercritical fluid extraction involves increasing the temperature and pressure of carbon dioxide (CO₂) until it surpasses its critical point, achieving a liquid–gas state. In this state, CO₂ acts as a solvent when combined with microalgae biomass [123]. The temperature and pressure required to reach this critical point depend on the type of solvent used. However, this method has significant drawbacks, primarily due to the high infrastructure and operating costs. Lipid extraction from microalgae is particularly challenging due to the energy-intensive and expensive nature of the process, which often requires solvents for additional energy recovery [117]. Moreover, the extraction process must be rapid and efficient, ensuring that the quality of the recovered lipids is not compromised.

In addition to the above, Ionic Liquids (ILs) have emerged as promising solvents for lipid extraction from microalgae due to their ability to solubilize lignocellulose, a major structural component of algal cell walls [116]. This property enables ILs to disrupt the robust cell walls of microalgae, thereby facilitating lipid recovery. For instance, Mukund et al. investigated the use of ten different ILs for the pre-treatment of Nannochloropsis oculata and Chlorella salina to enhance lipid extraction [124]. Among the ILs tested, butyrolactam hexanoate demonstrated superior cell disruption efficacy, lysing approximately 84% of the cells. Compared to the conventional Bligh–Dyer extraction method, lipid recovery rates for Nannochloropsis oculata and Chlorella salina were significantly higher at 134.9% and 85.4%, respectively. While ILs are generally considered environmentally benign and non-toxic, it is important to note that certain ILs may generate hazardous byproducts during their synthesis, potentially posing environmental risks.

Furthermore, enzyme-assisted extraction, which targets the cell walls to improve lipid recovery, provides an economical and environmentally beneficial substitute [116,118]. This technique has demonstrated the ability to both facilitate lipid fractionation and improve the efficiency of lipid extraction. Lipid recovery is greatly enhanced by enzymatic hydrolysis; nonetheless, the procedure relies on the appropriate enzyme selection and reaction conditions to efficiently break down the microalgal cell walls. To effectively speed up the lipid extraction process, these variables must be well controlled. For instance, He et al. [125] carried out an experiment to rupture the cell wall of Nannochloropsis using four hydrolase enzymes: cellulase, hemicellulase, papain, and pectinase and found that cellulase was the most successful of them.

5.1.2. Transesterification of Extracted Lipids

Crude microalgae lipid is then used in the transesterification reaction to produce biodiesel. Because microalgae crude oil has a high viscosity, it must be transformed into fatty acid alkyl esters, which have a reduced molecular weight. The process of transesterification transforms free fatty acids and triacylglycerols, or raw microalgal lipids, into biodiesel that is biodegradable, renewable, and non-toxic [53]. The lipid-to-alcohol ratio, temperature, duration, catalyst type, and FFA of the lipid all have a significant impact on transesterification efficiency [126]. Figure 5 illustrates the different stages of the transesterification procedure.

Different homogenous and heterogenous catalysts are employed for biodiesel production, as shown in Table 3. Among them, homogenous alkaline catalysts like NaOH and KOH are widely available, inexpensive, and utilized in the production of biodiesel fuels. Atmospheric pressure and 60–70 °C are used to conduct the reaction [5,61,89,127,128]. The catalyst and alcohol are combined first, and the mixture is then put into the reactor vessel to combine with the algal lipid to initiate the reaction [18]. Base catalyst reactions have some limits because they are not appropriate for algal lipids with more than 2% free fatty acid (FFA) concentration. Reduced catalytic activity and higher FFA favor the saponification process over transesterification, which lowers the output of biodiesel [126].

Heterogeneous catalysts are an environmentally benign and economical solution to the usual challenges associated with homogeneous catalysts, including equipment corrosion, water pollution, and challenging product and catalyst mixture separation [129]. Heterogeneous catalysts exhibit biphasic reactions between the catalyst and the reaction substance, allowing for easier separation of the solid catalyst from the end product and subsequent recycling and reuse [130]. It was discovered that heterogeneous catalysts possess numerous active sites for reactant adsorption and subsequent transesterification catalysis. In the alkaline catalytic process, the catalyst provides a negative charge at the active site to facilitate methanol adsorption, while an acidic catalyst supplies a positive charge for the adsorption of fatty acids. Like acidic catalysis, nucleophiles can attack the tetrahedral intermediate, leading to the removal of glycerol, the formation of a new ester, and the regeneration of the catalyst. Initially, the alkoxy ion in the triglyceride (TAG) reacts with the alkyl group, which then interacts with alcohol to form a tetrahedral intermediate. This process leads to the generation of carbon cations. The formation of this intermediate contributes to higher biodiesel yields and reduced activation energy, making the process efficient while preserving the extracted lipids. Zinc oxide (ZnO), magnesium oxide (MgO), calcium oxide (CaO), and other metal oxides are examples of heterogeneous alkaline catalysts. CaO has been examined the most extensively among them. Furthermore, several nano-catalysts. High biodiesel yields (>98.0%) were also demonstrated by Mn–ZnO, Li/ZnO–Fe₃O₄, CaO/MgO, SO₄/Mg–Al–Fe₃O₄, nanocrystalline CaO, and nano-sulfated zirconia [131].

Traditional methods for producing microalgae biodiesel face several significant challenges, such as lengthy extraction and conversion times, low recovery rates, high solvent consumption, and difficulties in optimizing biodiesel yield. Emerging methods like microwave-assisted and ultrasonic-assisted transesterification have been developed to address these issues. These techniques enhance the reaction rate, resulting in significantly shorter reaction times while also minimizing energy consumption [126]. However, Enzymatic transesterification of microalgae lipids can be considered the most ecofriendly and energy-efficient method due to the reaction being at room temperature, easy to separate from biodiesel, and reusability [132]. When concentrated alcohols are present in the medium reaction, most enzymes become sensitive. Nonetheless, Lipozyme TL IM (Thermomyces lanugi nosus lipase immobilized on acrylic resin) and Novozym^® 435 (Candida antarctica lipase immobilized on microporous resin) exhibit a high tolerance against deactivation, irrespective of the presence of methanol and ethanol, respectively [7]. However, these enzymes are much more expensive, and further investigation is needed on cheap and novel enzymes.

5.2. In Situ Transesterification of Microalgae Biomass

Lipid extraction and conversion to fatty acid alkyl esters (FAAEs) can be efficiently achieved using an integrated method known as in situ transesterification (ISTE) [133]. This process utilizes oily raw materials and combines lipid extraction and transesterification into a single step. Methanol plays a dual role in ISTE, functioning both as a chemical solvent and as a reactant, which reduces the need for additional solvents. By eliminating the separate processes for lipid extraction and solvent recovery, this method significantly decreases energy consumption [134]. Additionally, ISTE avoids the need for costly and labor-intensive steps such as cell killing and drying, leading to lower overall costs. Oliva et al. successfully extracted lipids using a hexane–methanol mixture (19:1 v/v) with KOH in a Kumagawa extractor, achieving an extraction yield of over 12% [135]. These lipids were then converted into fatty acid methyl esters (FAMEs), showcasing the potential for bio-sourced fuels within a circular economy framework. Mathimani et al. [136] compared the effectiveness of one-step direct transesterification (OSDT) and dual-step extraction-transesterification (DSET) on Chlorococcum sp. and Nannochloropsis sp., evaluating the FAME yield in each case. The FAME yields from both OSDT and DSET were nearly identical, at approximately 21 g per 100 g of dry biomass. Among the various OSDT reaction conditions tested, a reaction temperature of 75 °C produced the highest FAME yield, ranging from 70% to 71% based on lipid weight. This technology has the potential to reduce both fuel conversion process costs and processing unit costs. Unlike traditional biodiesel production methods, it eliminates the need for solvent extraction processes. Although direct esterification has shown promising results in laboratory settings, there are still several challenges that need to be addressed before it can be effectively scaled up for large-scale production.

5.3. Fuel Properties and Storage Stability of Microalgae

The suitability of microalgae-based biodiesel as a replacement for fossil diesel fuel depends significantly on meeting established standards, such as ASTM D6751, EN 14214:2008, and IS: 15607 [137,138,139]. The fatty acid composition of biodiesel plays a crucial role in determining its fuel properties. Common fatty esters found in typical biodiesel include palmitic, stearic, oleic, linoleic, and linolenic acids, which vary depending on the feedstock used [5,18,61]. Biodiesel rich in saturated fatty acids tends to have a high cloud point and viscosity, increasing the risk of engine nozzle clogging. Conversely, biodiesel high in polyunsaturated fatty acids generally exhibits better cloud points but suffers from poor oxidation stability [1]. Table 4 displays the fuel properties of biodiesel fuels extracted from different microalgae.

A crucial aspect of the practical application and long-term viability of biodiesel is its storage stability [140]. When exposed to air, oxygen, and light, biodiesel undergoes oxidation, leading to the formation of deposits in the fuel and resulting in increased viscosity and density. During storage, the unsaturated methyl esters in biodiesel start to oxidize and produce peroxides and hydroperoxides. These are further degraded into ketones and aldehydes and finally polymerized, forming sludge or deposits and rendering them unsuitable for use as motor fuel because they clog fuel filters [141]. Oxidative stability is an indicator used to quantify the resistance of fats and oils to oxidation. The oxidation stability can be improved by adding antioxidants, both natural and synthetic. In addition to using antioxidants, structurally altering the fatty acid methyl esters by hydrogenating or transisomerizing the original double bonds in the cis configuration to reduce the number of double bonds could improve the oxidative stability of biodiesel. Because trans fatty acids are less reactive than cis fatty acids, changing the FAME structure from cis to trans improves the stability of biodiesel [142,143].

Microalgae biodiesel, with its higher content of unsaturated fatty acids, is particularly susceptible to oxidation during storage, posing challenges to its long-term stability [7]. For instance, the storage stability of biodiesel produced from Nannochloropsis sp. is influenced by factors such as temperature, light exposure, and oxygen availability. Research has shown that the oxidative stability of microalgae biodiesel can be significantly improved by incorporating antioxidants like butylated hydroxytoluene (BHT) or naturally occurring antioxidants derived from microalgae. Additionally, proper storage conditions—such as keeping the biodiesel in dark, sealed containers at controlled temperatures—can help extend its shelf life. For example, studies on biodiesel made from Chlorella sp. suggest that storing it below 25 °C and minimizing air exposure can slow oxidation and prolong its shelf life.

For instance, biodiesel derived from Chlorella minutissima sp. has demonstrated varying storage stability, influenced by factors such as temperature, light exposure, and oxygen levels, and increased by the addition of 1000 ppm propyl gallate antioxidants [5]. Studies have shown that the presence of antioxidants, such as butylated hydroxytoluene (BHT) or natural antioxidants, can significantly improve the oxidative stability of microalgae biodiesel [5]. Moreover, the use of appropriate storage conditions, such as sealed, dark containers and controlled temperatures, can further enhance the longevity of microalgae biodiesel. For example, research on Chlorella sp. biodiesel has indicated that maintaining storage temperatures below 25 °C and minimizing exposure to air can reduce the rate of oxidation and extend storage life. These measures are essential for ensuring that microalgae biodiesel remains a viable and reliable alternative to fossil fuels.

Table 4. Microalgae biodiesel fuel properties [5,6,144,145,146].

S. No.	Parameters	Method	Microalgae Biodiesel Chlorella vulgaris	Chlorella minutissima	Spirulina platensis	Chlorella emersonii
1.	Calorific value (MJ/kg)	D240	39.45	39.15	45.63	36.9
2.	Density at 15 °C (kg/m3)	D7777	889	0.877	0.8637	nd
3	Viscosity at 40 °C (mm2/s) or cSt	D445	5.72	4.75	12.4	3.96
	Flash point (°C)	D93	155	-	189	212
4	Pour point (°C)	D97	−12	−3.76	−9	7
5	Copper strip corrosion	D130	1	-	1a	nd
6	Cetane number		57.03	-	70	52.5
7	Acid number (mg KOH/g)	D664	0.49	-	0.75	nd
8	Water and sediment (%)	D2709	0.03	-	3.9	nd
9	Methyl linolenate (%)	EN 14103	7.54	7.45%	nd	nd
10	Unsaturated ester (>4 double bonds) %	Internal method—GC	0	nd	nd	nd
11	Oxidation stability (IP, at 140 °C, h)	ASTM-D7545 and prEN16091	3.08	2.80 (min)	nd	nd

Table 4. Microalgae biodiesel fuel properties [5,6,144,145,146].

S. No.	Parameters	Method	Microalgae Biodiesel Chlorella vulgaris	Chlorella minutissima	Spirulina platensis	Chlorella emersonii
1.	Calorific value (MJ/kg)	D240	39.45	39.15	45.63	36.9
2.	Density at 15 °C (kg/m3)	D7777	889	0.877	0.8637	nd
3	Viscosity at 40 °C (mm2/s) or cSt	D445	5.72	4.75	12.4	3.96
	Flash point (°C)	D93	155	-	189	212
4	Pour point (°C)	D97	−12	−3.76	−9	7
5	Copper strip corrosion	D130	1	-	1a	nd
6	Cetane number		57.03	-	70	52.5
7	Acid number (mg KOH/g)	D664	0.49	-	0.75	nd
8	Water and sediment (%)	D2709	0.03	-	3.9	nd
9	Methyl linolenate (%)	EN 14103	7.54	7.45%	nd	nd
10	Unsaturated ester (>4 double bonds) %	Internal method—GC	0	nd	nd	nd
11	Oxidation stability (IP, at 140 °C, h)	ASTM-D7545 and prEN16091	3.08	2.80 (min)	nd	nd

6. Conclusions

This review shed light on the key factors influencing biodiesel production from microalgae, encompassing both upstream and downstream processes. Lipid accumulation in microalgae is significantly affected by environmental parameters like light, pH, nitrogen, and phosphate-averse conditions. Optimizing these crucial factors is essential before scaling up cultivation as they vary from species to species. The literature study also concludes that the use of wastewater as growth media can play a major role in the cost-reduction of biomass production.

Downstream processes, especially lipid extraction and biodiesel synthesis, are pivotal in achieving efficient biodiesel production. Lipid extraction is one of the challenging tasks; however, integrating pretreatment methods like microwave or ultrasonic techniques facilitates lipid extraction by disrupting cell walls. For biodiesel synthesis, the selection of a catalyst is a crucial step. Recently, heterogenous nano-catalysts outperformed traditional catalysts (Base catalysts like NaOH and KOH) due to their superior active sites, higher activity, stability, and reusability. In addition, one of the biggest challenges in microalgae biodiesel is its poor oxidation stability, as microalgae biodiesel is rich in unsaturated fatty alkyl esters. This can be mitigated by incorporating antioxidants.

To commercialize the microalgae biodiesel, future research should focus on breeding microalgae strains with high lipid content and rapid growth, optimizing cultivation conditions under adverse conditions, low cost and ecofriendly catalyst designing for transesterification processes, zero waste economy with a biorefinery approaches.