Industrial Biotechnology Today: Overview of Main Products

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Contents

Introduction

1. Biopolymers and their applications

1.1 Biopolymers produced by bacteria

1.2 Biopolymers produced by fungi and yeasts

1.3. Biopolymers produced by algae

2. Classification of enzymes and their application in various branches of industrial biotechnology

2.1. Bacterial enzymes

2.2. Fungal enzymes

2.3. Microalgae enzymes

2.4. Proteases

2.5. Lipases

2.6. Amylases

2.7. Pullulanases

2.8. Pectinases

2.9. Xylanase

2.10. Laccases

2.11. Transglutaminases

2.12. Phytases

2.12. Xylose isomerase

3. Amino acids and their production by industrial biotechnology

3.1. Production of amino acids

3.2. Amino acids produced by bacteria

4. Organic acids and their applications

4.1. Bacteria — producers of organic acids

4.2. Fungi — producers of organic acids

4.3. Yeast — producers of organic acids

4.4. Microalgae — producers of organic acids

5. Lipids for industrial biotechnology

5.1. Bacterial and yeast lipids

5.2. Fungal lipids

5.3. Lipids of microalgae

5.4. Lipid extraction

6. Antibiotics as products of industrial biotechnology

6.1. Classification of antibiotics

6.1.1. β-lactam antibiotics

6.1.2. Macrolides

6.1.3. Tetracyclines

6.1.4. Aminoglycosides

6.1.5. Peptide antibiotics

6.2. Main steps in biotechnological production of antibiotics

7. Vitamins obtained through microbial synthesis

7.1. Bacteria as sources of vitamins

7.2. Role of fungi in vitamin production

7.3. Vitamin production by microalgae

8. Hormones produced by microbial synthesis

8.1. Synthesis of hormones and steroids using bacteria

8.2. Biotechnological synthesis of hormones by fungi

8.3. Algae as a source of hormones and steroids

Conclusion

Conflict of interest

References

Introduction

Industrial biotechnology is one of the key areas of modern science and economics that develop at the intersection of biology, chemistry and engineering technologies. It is based on the use of biological processes and organisms, such as bacteria, fungi, microalgae and their metabolites, to solve urgent economic and environmental problems.

The global industrial biotechnology market was valued at over USD 400 billion in 2020 and is further expected to grow at an average of 7-10% in the coming years [1]. The main segments of the global industrial biotechnology market include such areas as biopharmaceuticals (production of drugs using microorganisms and cells), biopolymers (use of natural sources to create environmentally friendly packaging materials), biofuels (production of bioethanol and biodiesel to replace fossil fuels), and industrial enzymes used in the food and cosmetic industries [2]. The biotechnology market in Russia lags far behind the global indicators, but it has demonstrated positive dynamics in recent years. According to some estimates, the Russian biotechnology market is about 35-40 billion rubles, and is expected to grow by 19-22% in the next five years [3]. Despite the growth, the Russian biotechnology industry faces a number of challenges, namely: lack of investments, high barriers to market entry, and insufficient cooperation between research institutions and industry [4]. The main challenges is also the need to create an effective regulatory framework and to develop strategies to attract investment.

Industrial biotechnology processes can be divided into two main groups: production of biomass and production of metabolic products. But this classification does not take into account the key technological features of industrial biotechnology. It is important to analyze the stages and specificity of biotechnological production taking into account the final goal of the process. Figure 1 demonstrates a biotechnological production scheme.

The first step in any industrial process is to select an efficient producer by screening strains from various environments. It is also possible to use modern technologies such as CRISPR/Cas9 and genetic engineering, which allow modifying the metabolism of microorganisms to increase their ability to synthesize specific compounds (e.g., antibiotics or biopolymers) [5].  The next step is fermentation, which involves the use of microorganisms to convert carbon sources into various products such as enzymes, acids, alcohols and polymers. An example is the use of Saccharomyces cerevisiae to produce ethanol from sugars. In this case, it is necessary to ensure optimal conditions for the cultivation of yeast cells (temperature, pH and substrate ratio), which are critical for obtaining the desired yield of the target product [6]. One of the important economic components of the industrial fermentation process is the use of inexpensive and renewable carbon sources, such as agricultural waste or industrial by-products. Khan, M. I et al. demonstrated that bacteria of the genera Rhodococcus and Pseudomonas can utilize plant wastes to produce bioplastics and biofuels, which reduces the cost of production and improves the environmental conditions [7]. In addition, modern bioreactors are always equipped with sensors to monitor various cultivation parameters, which also contributes to maximizing the yield of target products and ensures high efficiency of the processes. López N. I. et al. were able to increase the yield of polyhydroxyalkanoates during continuous cultivation by optimizing the culture growth parameters [8]. At the final stage of industrial production, efficient extraction and purification of the target products must be carried out, which often involves such methods as centrifugation, precipitation, and chromatography [9].

Industrial biotechnology has great potential to address global challenges such as food security, development of clean energy sources and creation of new materials with improved properties. However, to fully unlock such its potential, further research and development, as well as favorable conditions for the introduction of biotechnological solutions into practice are needed. The advances in molecular biology, genetic engineering and microbiology open up new horizons for the creation of innovative products that can be applied in medicine, agriculture, ecology, food and energy industries. This review discusses the main types of industrial biotechnology products that have applications in various human endeavors. Special attention is paid to biopolymers, enzymes, biologically active substances and other products of biotechnological proesses.

Industrial biotechnology products can be segmented in different ways, for example, according to their type, source, or application area. The following types of industrial biotechnology products will be presented in this section: biopolymers, enzymes, amino acids, organic acids, lipids, antibiotics, vitamins, and hormones.

Figure 1. Biotechnological production scheme.

Biopolymers synthesised by microorganisms represent an important class of materials that are widely applied in various industries, including medicine, food, agriculture and biotechnology. Polyhydroxyalkanoates, exopolysaccharides, chitosan, alginates and other polymers have unique properties including biocompatibility, biodegradability and the ability to be produced from renewable resources.

Bacteria are one of the most studied sources of biopolymers due to their ability for high levels of biosynthesis under strictly controlled conditions.

Polyhydroxyalkanoates have more than 150 varieties and are one of the most studied classes of biopolymers synthesised by the bacteria Cupriavidus necator, Schelegelella thermodepolymerans, Pseudomonas spp., Ralstonia eutropha and Bacillus spp. Polyhydroxyalkanoates accumulate in microbial cells as storage materials at the deficiency of nutrients, such as nitrogen or phosphorus, and in the presence of excess carbon [10]. These polymers can be used in the production of packaging materials, medical implants, biodegradable films, and in agriculture [11, 12]. Polyhydroxyalkanoates are produced inductrially in Brazil, Austria, and China where various organic substrates are used including agricultural waste.

Exopolysaccharides such as xanthan and hyaluronic acid are produced by bacteria of the genera Xanthomonas, Leuconostoc and Streptococcus. Xanthan, produced by aerobic fermentation of sugars, is widely used in the food industry as a thickener and stabilizer, while hyaluronic acid, due to its moisturizing and regenerating properties, is applied in cosmetics and medicine [13]. Dextran and levan synthesized by bacteria of the genera Leuconostoc, Lactobacillus, Streptococcus, Aerobacter and Weissella [14] are used in the pharmaceutical industry, including the production of plasma substitutes and delayed drug release systems [15]. Exopolysaccharide curdlan is obtained by deep fermentation of bacteria of the genera Rhizobium, Agrobacterium, Alcaligenes, Cellulomonas, and Bacillus [16]. Due to its gelling ability, curdlan is used in food industry and biomedical research.

Yeasts are widely used in the production of polysaccharides such as β-glucans and mannans.

Chitosan, a chitin derivative, is synthesized by fungi Aspergillus niger and Mucor rouxii. This biopolymer demonstrates antimicrobial, antioxidant and biocompatible properties, which makes it promising for use in medicine, e.g. as wound coatings and for drug delivery [17].

β-glucans produced by the yeasts Saccharomyces cerevisiae, Candida albicans and Kluyveromyces lactis, and by the fungi Pleurotus ostreatus have immunomodulatory properties and are used in production of food supplements and in medicine as pharmaceuticals imporving the work of immune system [18].

Mannan-oligosaccharides synthesized by yeast S. cerevisiae are used as prebiotics in animal feeds to improve intestinal health and immune response [19].

Fucin is a biopolymer synthesized by some species of fungi, including those of the genera Fusarium and Aspergillus. This biopolymer is formed during metabolism in fungi and is released into the environment as exopolysaccharide. Fucin production depends on cultivation conditions such as nutrient medium composition, pH, temperature, and aeration [20]. Optimization of these parameters can increase the yield of the biopolymer and make its inductrial production economically viable [21]. Fucin is biodegradable and therefore attractive for use in various areas including medicine (e.g., development of biodegradable films and drug delivery systems) and agriculture (e.g., production of biodegradable packaging materials) [22].

Pullulan is mainly obtained as water-soluble polysaccharide from the yeast-like fungus Aureobasidium pullulans. Other known fungi producing pullulan are Tremella mesenterica, Cryphonectria parasitica, Teloschistes flavicans, Rhodotorula bacarum, Cytaria harioti and C. darwinii [23]. Pullulan is widely used in various industries including pharmaceuticals, cosmetics, biomedicine, and food production. Due to its high adhesive properties, it is in demand in paper production.

Alginates synthesized by brown algae (e.g., Laminaria spp.) are widely used in the food industry as thickeners and stabilizers, and in pharmaceutical industry to create hydrogels and wound coatings [24].

Carrageenans derived from red algae (e.g., Chondrus crispus) are used in food industry to improve the texture of products and in pharmaceuticals to create dosage forms and microcapsules [25]. Industrial production of alginates and carrageenans is actively developing in countries with rich aquaculture, such as Indonesia and Norway.

Due to their unique properties and ability to be produced from renewable resources, biopolymers help to reduce dependence on fossil resources and thereby reduce the negative impact on the environment. In general, industrial production of biopolymers requires optimization of microbial cultivation conditions, including strain selection, nutrient medium composition, pH, temperature,  and aeration. Modern biotechnological approaches such as metabolic engineering and CRISPR/Cas9 can improve microbial productivity and reduce production costs [26]. In addition, further research in metabolic engineering and optimization of production processes will expand the applications of biopolymers and make them more accessible for industrial use.

Environmental safety requirements for most industrial areas are constantly growing. That is why in various industrial areas chemical catalysts are replaced by enzymes, and primarily of a microbiological origin. The latter is due to the economic component of the fermentation process. In addition, microbial enzymes have some advantages compared to enzymes of plant and animal origin, amomg them a wide variety of microbial producers, easy production of enzyme in fermenters, the possibility of continuous cycles (daily), easy controllability of the process, etc.

Despite the long history of enzyme technology applications, large-scale production of enzymes in the form of purified and well-characterized preparations became possible only with the development of recombinant DNA technology. They have become widely used in a variety of industrial products and processes, including chemical, detergent, textile, food, feed, tanning and pulp and paper industries.

The development of techniques such as protein engineering and site-directed mutagenesis provides an opportunity to design enzymes with novel properties [27].

Industrial technologies of enzyme production using microorganisms differ depending on the type of the used producer and production conditions.

The bacteria Bacillus subtilis, Bacillus licheniformis and Escherichia coli are often used to obtain enzymes such as amylases, proteases and lipases [28] mostly by submerged fermentation method, where the cell mass is cultured in a liquid nutrient medium under high aeration. This method provides rapid bacterial growth and high enzyme yield [29]. The enzymes are isolated most often by centrifugation and protein precipitation.

Half of the commercially produced enzymes are of fungal origin due to the easy cultivation of fungi in bioreactors and high yields [30]. The fungi Aspergillus. niger, Trichoderma reesei, and Penicillium spp. are the major producers of cellulase, pectinase, and amylopectinase [31]. The enzymes are most commonly obtained by surface fermentation of fungal cells on solid nutrient media such as wheat bran or rice hulls. This method is effective for production of exoenzymes [32]. The submerged fermentation method is also used to cultivate fungi in liquid media with high carbohydrate content [33].

Microalgae such as Chlorella vulgaris and Spirulina platensis are used to produce lipases and phosphatases [34]. Microalgae synthesize enzymes in open or closed photobioreactors where they grow in liquid cultures under optimal light and in the presence of carbon dioxide. Since some algae act as autotrophs, their enzymatic productivity depends on light intensity and CO₂ availability, which requires the use of specific bioreactors [35].

The emergence of new applications of enzymes is leading to an increased demand for industrial enzymes, and the industry is responding with a steady stream of innovative products. Among the industrial enzymes currently in use, hydrolases such as proteases and lipases are the most common and are widely used in various industries e.g. in production of detergents, dairy products, and chemicals. Carbohydrases, especially amylases and cellulases, represent a significant group of enzymes [36-37].

Enzymes can be classified according to the main directions of their application: technical enzymes (used in the production of synthetic detergents; in the textile, leather, pulp and paper industries; in the production of biofuels); enzymes for the food and beverage industry; feed enzymes [38]. Table 1 summarizes the enzymes most frequently used in various industries.

Proteases (proteinases, proteolytic enzymes) are a group of enzymes capable of hydrolyzing peptide bonds in proteins. They are divided into exopeptidases (cleaving end peptide bonds) and endopeptidases (acting within proteins). Proteases are classified according to substrate specificity, catalytic mechanism, pH optimum, and other properties.

Subtilisins are serine proteases actively used in detergents due to their stability and low substrate specificity [40]. These ptoteases are produced by Bacillus spp. bacteria; they are active in the pH range of 6-11, with an optimum of 9-11. In 1985, the first genetically modified subtilisin, less sensitive to oxidation by hydrogen peroxide, was created [41]. Subsequent studies improved such subtilisin properties as thermostability, activity in organic solvents and at extreme pH [42].

Proteases are applied in the food industry to hydrolyze proteins and produce nutritious protein hydrolysates [43]. Acidic proteases used in the dairy industry replace rennet enzyme in cheese production [44]. Alkaline proteases are used to process waste materials such as keratinous materials and in production of feed [45]. In the leather industry, they replace chemicals for soaking hides and removing hair, which reduces contamination and energy costs [46].

Research on proteases is mainly focused on the development of new enzymes with improved characteristics and new applications. These works represent a significant potential for the improvement of industrial processes [47-48].

Lipases catalyze the hydrolysis of triglycerides to glycerol and fatty acids; they are widely distributed in nature. Lipases possess a catalytic triad (serine, histidine, and aspartate) and interfacial activation, which makes them highly efficient in applications with hydrophobic surfaces [49]. Lipases also perform esterification and transesterification, and are therefore actively employed in the food industry, production of biofuels and detergents, and in organic chemistry [50-51].

In the food industry, lipases are used to transesterify palm oil to create cocoa butter substitutes, synthesize flavor compounds, remove phospholipids, and create low-calorie products. Immobilized lipases are economically advantageous due to the possibility of their multiple uses [52]. In production of detergents used to remove grease stains, the best choice are enzymes resistant to alkaline environments such as lipases produced by Thermomyces lanuginosus and Pseudomonas mendocina [53]. In the leather industry, lipases used to degrease skins help to reduce the environmental pollution [54].

Table 1. Enzyme preparations and scope of their application [39].

In biodiesel production, lipases act as an environmentally friendly alternative to chemical catalysts, providing high product purity and efficient glycerol separation. Their high cost is partially decreased due to immobilization, the use of recombinant DNA and protein engineering [54-55]. Lipases are also in demand in organic chemistry for the synthesis of chiral compounds inpharmaceuticals and agrochemicals.

Amylases (glycoside hydrolases) hydrolyze α-1,4-glycoside bonds in starch, breaking it down to sugars. Microbial amylases are divided into exo- (glucoamylases and β-amylases) and endoamylases (α-amylases). Glucoamylases hydrolyze α-1,4 and α-1,6 bonds, releasing glucose, while β-amylases release maltose [56]. Endoamylases break internal bonds in starch to form oligosaccharides. Alpha-amylases, thermostable enzymes derived from fungi and bacteria, are used in food, textile, paper and fermentation industries [57-58]. Amylases are in demand due to their stability and activity under extreme conditions. For example, Bacillus sp. strains (B. subtilis, B. licheniformis) produce enzymes active in the pH range of 1.0-11.5 and temperatures of 25-90 °C, which is suitable for industrial processes.

Amylases are used to enzymatically liquefy and saccharify starch, replacing the less efficient acid hydrolysis. Amylases are also used in ethanol production: α-amylases break down starch into sugars, which are then fermented into ethanol by S. cerevisiae. In detergents, α-amylases help to remove starch even at low temperatures [58].

In the paper industry, α-amylases such as Amizyme^® and Thermamyl^® reduce the viscosity of starch for paper gluing, which improves paper stiffness and quality. In baking, enzymes break down starch into dextrins, which increases volume, improves texture, flavor, and color and extends the shelf life of bread [59].

Polysaccharides such as starches are polymers of D-glucopyranose with α-1,4 bonds and α-1,6 branching bonds. Complete hydrolysis of starch requires enzymes capable of cleaving both types of bonds. Pullulanases, widely distributed in nature, hydrolyze α-1,6 bonds by breaking the branching bonds in starch, amylopectin, and glycogen. Type I pullulanases (branching removal enzymes) act only on α-1,6 bonds, while type II pullulanases (amylopullulanases) can hydrolyze both α-1,6 and α-1,4 bonds [60]. For example, in the food industry, pullulanases are used to form maltotriose syrup, which has mild sweetness, low freezing point and moisture retention properties. Such syrup is used in the production of desserts, baked goods, beer and pharmaceutical products, including solutions for intravenous administration.

Pectinases (pectinolytic enzymes) hydrolyze pectin substances - plant polysaccharides with a framework of α-D-galacturonate links linked to α-1,4. They are divided into three groups: protopectinases (convert insoluble protopectin into soluble pectin), esterases (remove methoxy esters), and depolymerases (break α-1,4-linkages in pectin) [61]. The most common application of pectinases is in the extraction and clarification of fruit juices, where they degrade pectins, facilitating pressing, clear juice production and removal of turbidity. Cellulases and xylanases are also used to improve the efficiency of this process [62].

Pectinases are actively used in the textile industry, where together with other enzymes (amylases, lipases) they remove adhesives before dyeing fabrics, thus reducing the use of aggressive chemicals [63]. In addition, these enzymes are used to treat cotton, replacing alkaline treatment with the more environmentally friendly biopecanation, which improves water absorption and preserves fibers. In bast fibers, pectinases in combination with xylanases effectively remove gum thus reducing the adverse effect on the environment.

Pectinases are also used for the enzymatic treatment of wastewater from the plant food industry, promoting its decomposition [64]. In feed additives, enzymes increase nutrient digestibility in animals and reduce waste. Further studies on their stability, specificity and properties are needed to optimize the use of pectinases.

Plant biomass contains about 23% lignin, 40% cellulose and 33% hemicellulose, including xylan. Xylan is a heteropolymer with D-xylose in the backbone, branched L-arabinose and glucuronic acid. Its hydrolysis requires a complex of enzymes such as endoxylanases, β-xylosidases, esterases, etc. Endoxylanases cleave the main chain of xylan and β-xylosidases convert xylooligosaccharides into xylose. Xylanases are actively produced by Aspergillus and Trichoderma fungi [65].

The main application of xylanases is in the pulp and paper industry to remove lignin from kraft pulp, which reduces the use of chlorine bleaches and improves pulp quality. Enzymes are also used to improve dough properties in baking, increase bread volume and prolong its freshness [66]. In animal feed production, xylanases hydrolyze arabinoxylans in cereals, reducing their viscosity in the intestine and promoting weight gain [67].

In agriculture and the food industry, xylanases help to process wastes, converting xylan to xylose, which can be used in wastewater or to produce oligosaccharides. These enzymes also contribute to the synthesis of phytosterols, phytoalexins and alkylglycosides applied as surfactants [68].

Laccases also known as benzenediol oxygen oxidoreductase enzymes are able to oxidize a variety of organic substrates including ortho- and para-diphenols, aminophenols, polyphenols, polyamines, lignins and aryldiamines, as well as some inorganic ions. The laccase molecule is a glycoprotein, usually dimeric or tetrameric, with four copper atoms on each monomer distributed in three redox reaction centers. Laccases are widely used in biobleaching, delignification of cellulose, removal of dyes and pollutants, creation of biosensors, biofuel cells, and in food and textile industries [69].

In the pulp and paper industry, they provide environmentally friendly delignification and clarification without the use of chlorine. In the textile industry, laccases are used to degrade synthetic dyes and bleach fabrics. In bioremediation, they degrade xenobiotics including polycyclic hydrocarbons [69].

In the food industry, laccase are used to remove phenols from beverages (juices, beer, wine), improving their transparency. They also affect the properties of dough, increasing its stability by crosslinking biopolymers [69].

Transglutaminases are a class of enzymes that participate in the catalytic formation of a covalent bond between the free amino group of lysine (acyl acceptor) and the gamma-carboxamide group of glutamine (acyl donor) in proteins and peptides. This leads to modification of proteins by cross-linking intra- or intermolecularly, which in turn improves the end use of the protein [70]. In addition to cross-linking, transglutaminases are also capable of catalyzing the deamination of glutamine residues, which are acceptors of amino groups. In the absence of substrates with amino groups, water molecules can act as acceptors for the deamination of glutamine residues. Transglutaminases are found in animals, plants, and microorganisms and differ in size and calcium dependence: mammalian enzymes require calcium, while microbial enzymes do not [70].

In the food industry, transglutaminases improve the properties of proteins (dairy, soy, meat, and fish), increasing their nutritional value, elasticity, and moisture content. These enzymes are also used to create heat resistant films and increase the shelf life of products by protecting lysine residues [71]. In addition, transglutaminases find application in the textile industry to improve the quality of wool [72].

Phytases (phosphatases) play an important role in phytate hydrolysis and make phosphate available to organisms [73]. Phytases have been found in various organisms and can be divided into several groups depending on the carbon content of the phytate myo-inositol ring [73]. Some of them may also be called histidic acid phytases, α-propeller phytases, cysteine phytases or purple acid phytases based on their catalytic mechanism [73]. Phytases are widely used in animal feeds (pigs, poultry, and fish), increasing phytate-phosphorus uptake by 20-45% and reducing inorganic phosphorus requirements [73]. This reduces phosphate pollution of the environment and lowers costs for feed producers [74-75].

Phytases can be used in baking, plant protein production, corn milling and food additives. They are also promising for the development of functional foods.

Xylose isomerase is an enzyme that catalyzes the reversible isomerization of D-glucose to D-fructose and of D-xylose to D-xylulose. This enzyme belongs to the isomerase family, it has been found in nearly one hundred species of bacteria.  The production of glucose syrups began in Russia in 1811, when the German chemist Konstantin Kirchhoff from the Imperial Academy of Sciences in St. Petersburg obtained glucose syrup by heating starch with sulfuric acid [76]. Already in the middle of the 20th century, the discovery of the enzyme xylose isomerase, which converts glucose into fructose, played a key role. Takasaki from the Japanese Fermentation Institute identified a thermostable xylose isomerase from Streptomyces, which became the basis for the commercial production of high fructose syrups [77]. In 1967, Clinton Corn Processing (USA) adopted the Japanese technology to produce glucose-fructose syrups (GFS) and then improved the process using immobilization of the enzyme to produce syrup with 42% fructose [77]. Chromatographic separation was used to further increase the fructose concentration. GFSs with high fructose content have found applications as sugar substitutes in foods [78-79]. Technology development continues, aiming to reduce production costs [80].

Currently, enzymes operating under mild conditions are actively used in industry and various processes due to their efficient catalytic properties. However, thanks to new advances in protein engineering and directed evolution, we can expect the development of new enzymes with improved characteristics that will be used in both established technical fields and in completely new areas where the use of enzymes was previously impossible.

Since the beginning of the 20th century, the demand for amino acids has continued to grow due to their role in protein synthesis and participation in metabolism. These compounds are widely used in food, pharmaceutical, cosmetic industry and agriculture to improve enzymatic activity, immunity and muscle metabolism [81-83]. The importance of amino acids is emphasized by their deficiency, which can cause serious diseases [84-85].

Recent advances in genetic engineering have made it possible to obtain high-quality products with the desired properties [86]. Discoveries in this field have led to the development of innovative techniques for the isolation of amino acids, including fermentation using microbial cultures, extraction from protein hydrolysates, and chemical synthesis. Such techniques enable the development of sustainable products for food and agriculture [86]. The growing interest to amino acids has stimulated the development of innovative technologies, especially with an emphasis on sustainability.

Kikunae Ikeda, who researched monosodium glutamate in 1907, paved the way for the commercial production of flavor enhancers and initiated research in this field. Today, amino acids are used in various areas including food and feed additives, cosmetics, polymers and chemicals. The main challenge remains the optimization of methods for their industrial synthesis to improve efficiency and sustainability [87].

There are three main approaches to obtaining amino acids: extraction from protein hydrolysates, chemical synthesis, and microbiological techniques.

Extraction from protein hydrolysates is based on the unique characteristics of amino acids, such as pH, and is used to extract, for example, L-cysteine from keratin-containing materials (hair, feathers). This method is efficient and economical because it utilizes industrial waste [88]. Extraction using alkali generates environmental problems due to the formation of wastewater with toxic compounds [89]. The development of methods using supercritical water has improved the results while minimizing the negative environmental impact [90-91]. However, resource limitations of protein materials remain a problem.

Chemical synthesis involves reactions such as the Strecker method, but it requires expensive catalysts, is characterized by the use of toxic compounds (e.g., cyanide) and lack of enantioselectivity [92-93]. The well-known Bucherer-Bergs method is used for the synthesis of racemic amino acids, but it requires biochemical methods to separate isomers and is energy-consuming [94].

The microbiological process involves fermentation using microorganisms such as E. coli or S. cerevisiae. This method allows the production of optically pure amino acids with high efficiency and low amount of by-products [86]. Enzyme immobilization has been shown to increase process stability and product yield [95]. Fermentation is considered the most economical and environmentally friendly method despite the high costs of aeration, sterilization and equipment [96]. This approach prevails in the production of L-amino acids for the food industry and pharmaceuticals because it ensures high purity of the products.

The most commonly used microorganisms for amino acid fermentation are Corynebacterium glutamicum and E. coli [86]. Genetically modified C. glutamicum efficiently produces amino acids such as L-lysine and glutamic acid with high yields reaching 50% [97-98]. This glucose-preferring strain is used to produce L-glutamate, L-lysine, L-phenylalanine, L-tryptophan and other amino acids [99-100]. However, its growth is inhibited at high substrate or product concentrations, such as glucose above 50 g/L or L-glutamic acid above 12 g/L [101].

E. coli is used for the synthesis of such amino acids as L-methionine, L-lysine, and L-threonine [86]. Site-specific mutagenesis and modification of metabolic pathways make it possible to create strains for the production of branched-chain amino acids such as L-valine, L-leucine, and L-isoleucine, which makes them promising for feed, cosmetics, and pharmaceuticals [102].

Genetic engineering has made it possible to use waste as a raw material for fermentation, making the process economical and environmentally friendly. Innovative technologies such as nanofiltration membranes for integrating purification and production, as well as reactor modeling, are contributing to increased efficiency and lower costs. Looking forward, the use of bacteria and microalgae to convert waste into amino acid-rich proteins opens up new opportunities for protein supplementation and sustainable production [86].

Organic acids are low molecular weight organic compounds containing one or more acidic functional groups such as carboxyl, sulfonic, hydroxyl or thiol groups. Because of these functional groups, organic acids serve as important building blocks for the production of chemical compounds that have a wide range of applications in global markets. In the past, such acids were often derived from petroleum, but given the unsustainability of this resource and associated significant greenhouse gas emissions, a shift to biotechnological production methods is becoming necessary [103].

Bacteria are important producers of various organic acids. Among the most well studied compounds are lactic, succinic and acetic acids.

Lactic acid is produced by various strains of Lactobacillus casei, as well as Rhisopus oryzae and Endomycopsis fibuligera [104]. It is widely used in the production of biodegradable plastics (e.g., polylactide) and in the food industry as a preservative and acidity regulator.

Acetic acid is synthesized by the bacteria Acetobacterium woodi and Clostridium aceticum by microbiological conversion of hydrogen and carbon dioxide [105].

Succinic acid (butanoic acid) is an important chemical compound that is widely used in various industries [106]. The main bacteria involved in the synthesis of succinic acid are: E. coli, Basfia succiniproducens, C. glutamicum. This acid has gained commercial importance as a fuel additive and raw material for the chemical industry [107].

Fungi also play a significant role in the biosynthesis of organic acids. Of particular interest are strains of the genera Aspergillus and Penicillium, which are used to produce citric and oxalic acids.

Citric acid (2-hydroxy-propane-1,2,3-tricarboxylic acid) is a widely demanded compound in the food and pharmaceutical industries due to its safety, high water solubility, and pronounced chelating and buffering properties. In addition, citric acid is widely used in the production of detergents, cosmetic products, and a number of other industrial applications [108]. Modern industrial production of citric acid is based on the use of A. niger and various Rhizopus species in the process of deep fermentation in media supplied with starch or sucrose [109]. To increase the competitiveness of this production, researchers are trying to reduce costs by switching to cheaper substrates such as agro-industrial waste [110]. However, such substrates require pre-treatment to ensure nutrient digestibility, which can increase the complexity and cost of the process, especially if high product purity is required.

Another example is gluconic acid synthesized by Aspergillus spp. which is used in the textile and paper industries and is also applied as a rust remover [111]. Synthesis of this acid is carried out by microbial fermentation using various carbon sources containing glucose hydrolysates.

Itaconic acid is an α,β-unsaturated biaxonic carboxylic acid produced by the fungi Aspergillus itaconicus and Aspergillus terreus [112]. This compound finds wide application in the chemical industry, mainly in the production of various polymers including carboxylate rubbers, styrene butadiene polymers and nitrile latexes [113]. In addition, itaconic acid serves as a feedstock for the synthesis of pyrrolidones, butyrolactone, methylbutanediol, and is also used in the production of detergents, herbicides, and solvents [114].

Fumaric acid is a trans-isomer of butenedioic acid, which is produced by various representatives of mucor fungi, including the genera Mucor, Rhizopus and Cunninghamella [115]. This biotechnological process is based on the ability of these microorganisms to convert carbohydrates to fumaric acid through tricarboxylic acid metabolic pathways. The resulting fumaric acid is widely used in various industries: food industry (food additives, flavorings), pharmaceuticals (synthesis intermediates), cosmetics (acidity regulators), and chemical industry (polymer production). Fumaric acid is used to obtain maleic acid, which is applied in the production of: resins, paints and varnishes [115].

There are various yeast species capable of producing organic acids. One of the most studied is Yarrowia lipolytica, as well as a number of other non-fermenting yeasts such as Candida sp., Pichia sp. and Kluyveromyces sp. These microorganisms have high enzymatic activity and are able to efficiently convert simple and complex substrates into target products - organic acids. The mechanism of organic acid biosynthesis in yeast is due to the conversion of carbon sources such as glucose, xylose or hydrocarbons into metabolites of the intermediate and terminal Krebs cycle as well as other anabolic pathways. A peculiarity of the yeast Y. lipolytica is their ability to process hydrophobic substrates (e.g., fatty acids and oils), which makes them extremely promising for processing plant and animal wastes [116]. For example, this yeast can produce citric and dicarboxylic acids by decomposition of alkanes.

In addition to citric acid, yeast shows high efficiency in the production of succinic acid. This metabolite is considered as “platform chemistry” with diverse applications ranging from biomaterials to pharmaceutical ingredients. The strains of Candida utilis and partially modified S. cerevisiae were used in the large-scale production of succinic acid, and application of genetic techniques for strain improvement increased the yield of this acid in industrial processes [117].

Another important area is lactic acid biosynthesis. The cells of the yeast Kluyveromyces lactis produce lactic acid under favorable conditions of anaerobic fermentation [118]. This process is used to produce biodegradable plastics, which is relevant in the context of the global environmental program aimed at reducing plastic waste.

Microalgae are promising organisms for the synthesis of organic acids on an industrial scale due to their ability for rapid fermentation and high carbon dioxide assimilation efficiency [119]. They can produce a variety of organic acids such as myristic, palmitic, and lanolic acids, as well as other metabolites including lipids and carbohydrates [120]. The main advantages of their use are their ability to absorb CO2 in a highly efficient manner, which helps to reduce greenhouse gas emissions [121], and their growth rate, which allows more production in a shorter time. [122]. Microalgae are attractive for application in various ecosystems as they are able to survive in both fresh and salt water [123]. Currently, there are difficulties in the development of bioproducts using microalgae on an industrial scale, which are related to the optimization of cultivation conditions and extraction of metabolites [124].

Despite the obvious advances, the production of organic acids faces a number of limitations. For example, the use of expensive pure substrates, low yields in some cases, and difficulty in scaling up processes require the development of more efficient synthesis methods. Ongoing research in this area is aimed at reducing production costs, increasing productivity and expanding the functionality of organic acids to address a range of challenges related to environmental safety.

Lipids are important components of cell membranes, they provide the structural integrity of cells and transport of substances [125.]. In industry, lipids are widely used for the production of biodiesel, food additives, creams, lotions, and pharmaceuticals [126]. In recent years, microbial lipids have become a promising alternative to traditional lipids due to their rapid production, environmental friendliness, and ability to be grown on various substrates, including agricultural waste.

The application of microbial lipids in the food industry, such as the production of omega-3 and other fatty acids, has received special attention [127]. Polyunsaturated fatty acids such as n-3 (ω-3) and n-6 (ω-6) are healthy and are intensively used in fortified foods [128]. Microorganisms are also used to produce high-value cocoa butter, the cost of which is increasing due to growing demand [129].

Modern technologies, including CRISPR/Cas9 gene editing, have significantly increased the efficiency of microbial lipid production and improved fermentation processes [130].

The production of bacterial biomass to obtain lipids is becoming promising due to their potential use in biofuels and production of sustainable energy. Many bacteria such as Rhodococcus opacus, Acinetobacter calcoaceticus and Acinetobacter spp. accumulate up to 87% of their mass as triacylglycerides, which are converted into biodiesel [131-132]. Compared to oilseed crops, bacteria grow faster, produce higher lipid yields, and efficiently utilize various carbon substrates, including industrial wastes [133].

The accumulation of triacylglycerides is associated with cellular stress and consists of two stages: active growth in a carbon-rich medium and the “oil” phase, which is triggered by nitrogen deficiency [134]. The choice of substrate (glucose, sucrose or cheap waste) affects the efficiency of the process. For example, R. opacus accumulated up to 64.47% of lipids using sugarcane cake hydrolysate as cheap substrate [135].

Modern enzymatic methods, such as additional supplies of essential nutrients, have improved lipid yield by prolonging the growth phase and controlling the resource limitation [136]. For example, addition of glycerol to the culture medium enhanced lipid accumulation in R. opacus [137]. Thus, methods for optimizing substrates and culture conditions are the key to increasing the profitability of microbial lipid production.

The major yeast genera used for lipid production include Yarrowia, Candida, Rhodotorula, Rhodosporidium, Cryptococcus, Trichosporon and Lipomyces, but only about 30 species are able to accumulate more than 20% of their biomass as lipids [138-139]. The most studied are Yarrowia lipolytica, Rhodotorula glutinis, Rhodosporidium toruloides, Cryptococcus curvatus and Lipomyces starkeyi; Trichosporon fermentans also produce high amounts of lipids [140]. Lipid production is stimulated by excess carbon from glucose, glycerol or fatty acids under nitrogen limitation, leading to the synthesis of triacylglycerides (TAGs), the main lipid stores, accounting for up to 90% of the total lipid content [141-142].

High carbon to nitrogen (C/N) ratio and the activity of enzymes such as glycerol-3-phosphatacyltransferase enhance lipogenesis [143-144]. Oxygen saturation stimulates the synthesis of unsaturated fatty acids, and oxygen is also important for the Krebs cycle [145]. Environmental factors such as temperature, pH, and phosphorus for membrane phospholipids significantly affect the process [146-147].

Lipids accumulated by fungi include triglycerides, fatty acids, and polar lipids that serve as energy reserves and components of cell membranes. The fungi A. niger can produce biomass with up to 52% protein and 1% to 10% lipid content [148-149], while Mucor circinelloides and A. terreus are highly lipid-producing, with accumulation levels up to 38% of dry biomass. For example, A. terreus is rich in oleic and stearic acids, making it promising for biodiesel production [150-151].

Lipid accumulation occurs in two stages: active biomass growth and transition to lipogenesis, when nitrogen sources are depleted but carbon remains abundant [152-153]. The process is based on carbohydrate metabolism and synthesis of acetyl-CoA converted into lipids [154].

Fungal fermentation can be carried out by two methods: deep and solid state cultivation. Deep fermentation allows better control of environmental parameters and reduces the risk of contamination, which makes it preferable for industrialized production [155-156]. It is widely used for the production of fungal lipids and biomass, especially due to its efficiency and technological sophistication.

The composition of microalgae biomass varies greatly depending on species and growth conditions. The protein content ranges from 18 to 56%, lipids from 7 to 48%, and carbohydrates from 15 to 46% [157]. For example, the protein content of Spirulina maxima reaches 71% protein,  microalgae contain essential amino acids, polyunsaturated fatty acids, carotenoids and chlorophyll [158-159]. C. vulgaris and Scenedesmus spp. are characterized by high protein (56.1% and 49%) and lipid content (12.5% and 12.1%) [160]. They exhibit diverse fatty acid profiles: C. vulgaris is rich in polyunsaturated fatty acids, while S. platensis is rich in saturated fatty acids, including palmitic acid.

Fungi and yeast are superior to microalgae and bacteria in lipid production due to their ability to metabolize a wide range of substrates, which contributes to the yield of fatty acids such as palmitic (C16:0), oleic (C18:1), linoleic (C18:2) and α-linoleic (C18:3) [161-162]. These compounds are in demand in food, cosmetic and pharmaceutical industries due to their properties and nutritional value. Linoleic acid and γ-linoleic acid are used in nutrition and in the treatment of inflammation, while saturated fatty acids such as myristic acid (C14:0) are applied in personal care products [163]. Lipid production by fungi and yeast shows commercial potential due to high yields and a wide range of lipid applications, including biofuels.

Lipids produced by microorganisms include fatty acids with chain lengths ranging from 6 to 36 carbon atoms and are classified as saturated, monounsaturated or polyunsaturated ones [164-165]. These fatty acids are widely used in biofuels, and monounsaturated or polyunsaturated fatty acids produced by microalgae and traustochytrids find applications in nutraceuticals [164].

Lipid extraction remains a key challenge for industrial production due to their diverse polarity and location (intracellular and extracellular). Extracellular lipids are extracted more easily using filtration or centrifugation, whereas intracellular lipids require disruption of the cell wall [166]. This is accomplished by wet or dry biomass methods. The wet solvent extraction method is preferred due to low energy consumption, but it depends on lipid availability and mass transfer process [131, 166].

Chemical solvent extraction methods are efficient but pose environmental and financial risks [167]. Mechanical methods, such as bead milling and homogenization, are safer but require significant energy input [168]. Physical methods, including pulsed electric field technology and ultrasonic processing, are environmentally friendly and scalable, demonstrating high lipid yields and low energy consumption [169-170].

Overall, it is noteworthy that microbial lipids are a promising source for the production of sustainable bioproducts such as biosurfactants, carotenoids, dietary supplements, biofuels, pharmaceutical and cosmetic products [171-172], highlighting their versatile industrial potential [173-174]. Their versatility covers energy, nutrition, cosmetics and biodegradable materials. The production of lipids from agro-industrial wastes increases the efficiency of synthesis, reduces land use and competition with food crops, making them a sustainable feedstock for biofuels and reinforcing their importance in the bioeconomy [171, 175].

Antibiotics are a heterogeneous group of biologically active molecules that inhibit the viability of microorganisms. Initially, they included only compounds of natural origin produced by microorganisms, but later the definition included synthetic drugs [176-177]. The first anti-infectious drug, salvarsan, was synthesized by Paul Ehrlich in 1907 and was used to treat syphilis [178-179]. The discovery of penicillin by Alexander Fleming in 1928 ushered in the era of antibiotics, but its commercial production started only in the 1940s thanks to Penicillium chrysogenum [180-181].

The golden age of antibiotics is considered to be the 40s-60s of the 20th century, when many natural and synthetic compounds were discovered [182-183]. The classical producers are fungi and bacteria, in particular actinobacteria, which synthesize about 90% of commercial preparations [184]. In recent years, microalgae and cyanobacteria have been investigated for their potential to develop new antimicrobial drugs [185-186]. Current developments are focused on chemical modification of known microbial metabolites, allowing the development of semi-synthetic antibiotics with improved characteristics including broad spectrum of action and resistance [176]. Current research shows the high potential of entomopathogenic fungi [187] and microalgae [188] as sources of new antibiotics.

There are several classifications of antibiotics: by type of pharmacological effects (bactericidal and bacteriostatic), by activity (broad and narrow spectrum of action), but the most common types of classifications are based on the molecular structure and mode of action of antibiotics [189-190]. According to the mode of action, antibiotics are divided into inhibiting cell wall synthesis, disrupting the structure or function of the cell membrane and protein synthesis, affecting the structure and function of nucleic acids, or blocking key metabolic pathways [191]. Based on the molecular (chemical) structure, antibiotics are classified into β-lactams, macrolides, tetracyclines, quinolones, aminoglycosides, sulfonylamides, peptide antibiotics, and oxazolidines [192]. It should be noted that β-lactams, aminoglycosides, glycopeptides, quinolones, and lipopeptides acr as bactericides, while sulfonamides, tetracyclines, and oxazolidinones exert a bacteriostatic effect [193-194]. Among macrolides there are preparations of both bactericidal and bacteriostatic character. Since one of the ways to obtain new antibiotics is to modify the chemical structure of already known antibiotics, some classes of antibiotics differing in chemical structure and including natural and semi-synthetic compounds will be discussed in more detail below.

Antibiotics of the β-lactam class contain a β-lactam ring in their chemical structure, which inhibits peptidoglycan synthesis by binding to penicillin-binding proteins of bacteria. This disrupts the cell wall, causing cell lysis and cell death [195-196]. The β-lactams include penicillins, cephalosporins, monobactams, and carbapenems, which differ in side chains and structure [197].

Penicillins contain a β-lactam ring connected to a thiazolidine ring. They are enzymatically produced on the basis of 6-aminopenicillanic acid [198], the synthesis of which depends on the nutrient medium and genetic modification of strains [199]. The production of semi-synthetic penicillins involves chemical addition of side chains to 6-aminopenicillanic acid to produce compounds such as ampicillin and amoxicillin [200-201].

Cephalosporins produced by A. chrysogenum are characterized by stability and spectrum of activity. The production of semi-synthetic cephalosporins is based on the modification of 7-aminocephalosporanic acid [202]. Genetic engineering is used to increase titers and develop new compounds [203-204].

Monobactams, such as aztreonam, have a monocyclic structure and are entirely synthetic in origin [205].

Carbapenems, such as thienamycin, were first isolated in 1976 from Streptomyces cattleya [206-207]. However, due to their low yield and instability, their commercial production is based on chemical synthesis.

Macrolides are antibiotics containing 14-, 15-, 16-, or 18-membered lactone rings with attached aminosugars [208-209]. The first macrolide, erythromycin A, was isolated from Saccharopolyspora erythreus (formerly Streptomyces erythreus) in 1952. It became the prototype for the creation of semi-synthetic analogs such as azithromycin and clarithromycin [210-211].

14-membered oleandomycin is synthesized by Streptomyces antibioticus, and 18-membered fidaxomicin by Dactylosporangium aurantiacum [212-213]. The composition of the nutrient medium strongly affects the biosynthesis of macrolides: for example, propyl alcohol is added to the medium for erythromycin production and ammonium nitrogen, instead of nitrate nitrogen, is added for nystatin production (by S. noursei) [214].

Tetracyclines are broad-spectrum antibiotics based on the tetracyclic nucleus (A, B, C, D) [215]. The first tetracycline, chlortetracycline (aureomycin), was isolated in 1945 from Streptomyces by Benjamin Duggar, and in 1949 oxytetracycline (terramycin) was discovered by Alexander Finlay [216]. Their study made it possible to create synthetic and semi-synthetic analogs, as well as to modify the cultivation medium to obtain compounds with altered structure, for example, tetracycline instead of chlortetracycline in the absence of chlorine [217]. Semi-synthetic doxycycline is obtained from oxytetracycline, for example, by dehydration at C6 using a catalyst (rhodium on carbon) [218].

Tetracyclines are divided into generations: natural (1st), semi-synthetic (2nd) and synthetic (3rd) [219-220]. Their production was for a long time based on the fermentation by strains of the genus Streptomyces or semi-synthetic processes, later the third generation clinical preparations were approved [221].

Aminoglycosides contain two or more aminosaccharides linked by glycosidic bonds to an aminocyclic ring and are among the oldest antibiotics along with β-lactamases, macrolides, and tetracyclines [222-223]. Streptomycin was the first compound of this class to be discovered in 1943, derived from Streptomyces griseus; the second was kanamycin in the culture fluid of Streptomyces kanamyceticus [224]. The high toxicity of streptomycin necessitated the search for new less toxic representatives of aminoglycosides, which led to the discovery of such antibiotics as gentamicin (isolated during fermentation from Micromonospora purpurea), neomycin (isolated from Streptomyces fradiae), tobramycin (isolated from Streptomyces tenebrarius), and amikacin (a semi-synthetic derivative of kanamycin), among others [225].

Antimicrobial peptides are natural compounds with a broad spectrum of action against bacteria, viruses, fungi, and even cancer cells. They are synthesized to compete with microorganisms [226-228]. The first peptide antibiotic, gramicidin, was discovered in 1939 by René Dubo from bacteria of the genus Bacillus [229]. The classification of antimicrobial peptides varies and includes classes such as polypeptide, lipopeptide and glycopeptide antibiotics.

Polypeptides, such as polymyxins, are cationic cyclic decapeptides synthesized by Bacillus polymyxa. Polymyxins B and E (colistin) are used in clinical practice [230-231].

Lipopeptides such as daptomycin are produced by Streptomyces roseosporus and are used to treat various infections due to their antimicrobial and surface action [232-233].

Glycopeptides like vancomycin and teicoplanin contain a peptide framework with aromatic amino acids and sugars. Amycolatopsis orientalis and Actinoplanes teichomyceticus are the producers. Modern semi-synthetic analogs, such as telavancin and oritavancin, have improved properties, including efficient binding to the target and an extended half-life [234-235].

Thus, the development of industrial antibiotic production has not only focused on improving the performance of strain-producing strains, but also on the development of new nutrient media formulations and various methods of catalyzing the antibiotic molecule to remove unwanted side chains and/or attach new ones [236-237].

Industrial production of antibiotics involves many steps aimed at increasing product yield and reducing production costs, using modern technologies such as computerized control of fermentation parameters [238-241]. For example, the technology of penicillin production has evolved significantly: instead of a batch process using lactose, semi-continuous cultivation on substrates such as mixtures of glucose and sucrose with computer-controlled parameters has been used [200].

Industrial production includes the following main steps: obtaining the producer strains, development of cultivation conditions, isolation and purification of the antibiotic, scale-up and production fermentation [242]. Genetic and metabolic engineering enables the development of highly efficient producer strains [243-245]. Cultivation conditions include the selection of media with sources of carbon, nitrogen, and phosphorus, as well as factors such as trace elements and amino acids, which can enhance biosynthesis [198, 242]. For example, for A. chrysogenum, the addition of methionine stimulates the synthesis of cephalosporins [200].

Antibiotics, being secondary metabolites, are synthesized in the stress phase of the growth of microbial cells following the exponential phase. This phenomenon is used in two-stage cultivation: first, the biomass is increased, and then the active synthesis of antibiotic is started in a special medium [246-247]. The obtained preparations can be chemically modified to produce semi-synthetic antibiotics.

After fermentation, the culture liquid is treated; the product is isolated and purified. Methods include organic solvent extraction, adsorption, ion exchange and chemical precipitation, the choice of approach depending on the characteristics of the antibiotic [239, 242]. For example, penicillins G and V are extracted with organic solvents, whereas aminoglycosides require other methods due to their polarity [240]. In the final step, the drug is treated to remove water, often using lyophilic drying to preserve biological activity [248-249].

Thus, modern fermentation and downstream processing technologies, accompanied by genetic engineering tools and process optimization, allow the successful production of antibiotics on an industrial scale.

In recent decades, there has been considerable interest in environmentally friendly and sustainable methods of vitamin production. Vitamins essential for the maintenance of human health have traditionally been obtained from plant sources or synthetically. However, the use of microorganisms such as bacteria, fungi and microalgae opens new horizons for biotechnological production of vitamins on an industrial scale [250-251].

Bacteria are key players in vitamin biosynthesis, e.g., species of the genera Corynebacterium and Bacillus are used for the production of vitamins B - B12, B2 and B6. For example, BASF (Germany) is engaged in the production of riboflavin using a genetically modified strain of B. subtilis [252].  C. glutamicum is widely used in industry for vitamin B12 biosynthesis due to its ability to assimilate various carbon and nitrogen sources [253]. Propionibacterium shermanii, Propionibacterium freudenreichii and Pseudomonas denitrificans are highly productive when fermentation conditions (pH, temperature) and nutrient medium composition are optimized; therefore, they are used for industrial synthesis of cyanocobalamin (vitamin B12) [254]. One of the most significant examples is the production of ascorbic acid (vitamin C) based on the combined Reichstein-Grüssner technology, which includes sequential stages of chemical synthesis and biotechnological conversion. The process involves the bacterium Gluconobacter oxydans, which transforms glucose into sorbitol with its subsequent oxidation to sorbose [255]. The cells of Brevibacterium spp. efficiently uptake carbohydrates and produce pantothenic acid (vitamin B5) during their life cycle, which plays a key role in cell metabolism and coenzyme A synthesis [256]. Photosynthetic bacteria such as Rhodobacter sphaeroides and Corynebacterium play a significant role in the biosynthesis of vitamin A through its precursor β-carotene [257].

Fungi are also extensively used in the biotechnology industry due to their ability to produce vitamins, especially ergosterol, the precursor of vitamin D2. For example, the yeast S. cerevisiae is able to accumulate large amounts of ergosterol, which is converted to vitamin D2 when exposed to ultraviolet radiation [258]. A. oryzae and A. niger are employed in vitamin B2 production due to their high fermentation activity and easy adaptation to industrial conditions [259]. Eremothecium ashbyi is also one of the microorganisms used for the production of vitamin B2 (riboflavin) [260]. The fungi C. maltosa are used in the industrial production of ergosterol and ubiquinones by extraction from microbial lipids, which are by-products in the production of protein-vitamin concentrates [261].

Various mold fungi are capable of synthesizing vitamin A (in the form of β-carotene), and their content of the provitamin is many times higher than its amount in plant sources. For example, when cultivating Blakeslea trispora, its biomass accumulates 3-8 thousand µg of β-carotene per 1 g of mycelium, while in carrots it only makes up 60 µg. B. trispora is one of the most efficient carotenoid producers, demonstrating stable productivity under controlled fermentation conditions lasting 6-7 days [262]. For industrial production of β-carotene, a special fermentation process is used, where female and male mycelial strains are grown separately and then mixed in a bioreactor at a ratio of 1:15, which allows achieving a concentration of β-carotene up to 2 000 mg/L [263].

Microalgae represent one of the most promising platforms for industrial vitamin production [264]. Microalgae capable of producing carotenoids, including β-carotene, have received special attention. Microalgae such as Chlorella and Spirulina are capable of synthesizing vitamins A, E and D [265]. These algae can be used both as food and as a source of vitamins for dietary supplements. In addition, they contain many other nutrients such as proteins, minerals and antioxidants [266]. Of particular importance is that the content of some vitamins in microalgae far exceeds their concentration in terrestrial plants [267].

Vitamins and carotenoids obtained from microalgae are widely used in the food industry (food fortification), pharmaceuticals (production of vitamin supplements), and cosmetology (creation of cosmetics). However, despite significant prospects, the development of industrial microalgae biotechnology is limited by high production costs [263]. The solution to this problem requires optimization of all production stages - from strain selection to extraction and purification methods.

Thus, microbial synthesis of vitamins demonstrates high efficiency and environmental safety, which makes it a priority in the modern vitamin industry.

Hormones and steroids play an important role in medicine, agriculture and biotechnology [268]. Current methods for the production of these biomolecules include not only chemical synthesis but also biosynthesis using bacteria, fungi and algae [269]. These methods provide the possibility of producing complex molecules with high efficiency, lower costs and less environmental impact compared to traditional approaches [270]. In recent years, microorganisms have been considered as promising biocatalytic platforms that can produce both natural and modified analogs of steroidal compounds and hormones.

Synthesis of hormones using bacteria can be cost-effective and allows the production of hormones without the need to extract them from organic sources. The bacteria E. coli and B. subtilis are model organisms for heterologous expression of various peptide hormones [271]. A significant achievement of biotechnology was the development of a method of insulin production using recombinant DNA technologies [272]. This method involves the introduction of genes encoding the production of preproinsulin into the genome of bacterial cells, which provides them with the ability to produce this hormone on a large scale [273]. Bacteria can also synthesize other hormones, such as somatotropin, which is used to stimulate growth of farm animals [274].

One of the important areas of research is the study of the use of actinomycetes and other bacteria for biotransformation of complex steroid compounds.  Studies have shown that bacterial strains Rhodococcus ruber, Mycobacterium sp. and others possess key enzymes that can selectively change the structure of steroids at given positions. For example, the process of steroid hydroxylation at positions C-9 and C-11, which is an important step in the production of drugs such as cortisone and prednisolone, has been described [275]. Donova with co-authors, in their study of steroid catabolism by bacteria of the genus Mycobacterium, which can degrade the steroid molecule with the removal of the side chain, demonstrated a process important for the production of such intermediates as androstenone and androstadienedione [276].

Fungi of the genera Aspergillus, Penicillium and Rhizopus, are key biotechnological agents in the transformation of steroids [277]. Their unique ability to specifically hydroxylate steroid compounds makes it possible to obtain important pharmaceuticals including cortisone and prednisolone [278]. For example, Rhizopus nigricans has traditionally been used for the hydroxylation of progesterone to produce pharmaceuticals with anti-inflammatory and immunosuppressive properties. The mechanism of progesterone hydroxylation by the fungus Rh. nigricans, involves the conversion of progesterone to the less toxic 11α-hydroxyprogesterone, which is then excreted from the mycelium. The hydroxylation process is inducible and involves specific progesterone receptors [279].

The production of sterols by fungi occupies a special place in modern biotechnology [280], among which ergosterol is an important precursor for the synthesis of vitamin D [281]. The unique enzymatic systems of fungi provide highly selective modification of steroid molecules [282], which greatly simplifies the production of the required compounds. With an improved understanding of fungal metabolic pathways [283] and advances in genetic engineering [284], it has become possible to create specialized strains with improved production characteristics. These “super strains” demonstrate the ability to synthesize complex hormonal compounds on a large scale [285], which opens up new perspectives for their industrial applications.

Despite the prospects of hormone synthesis by fungi, scaling up the process is difficult. The key limitations are the relatively low growth rate of fungal cultures and insufficient concentration of target metabolites, which requires careful optimization of fermentation conditions [286].

Algae are a unique natural source of steroidal compounds and phytohormones with potential biological activity [287]. Of particular interest are representatives of Sargassum and Chlorella, characterized by the presence of specific steroidal components, which makes them promising raw materials for the production of biologically active additives [288]. The phytohormonal complex of algae includes important plant growth regulators: auxins, cytokinins and gibberellins. These compounds are not only involved in the regulation of growth and development of algae themselves, but can also have a stimulating effect on higher plants when used as biostimulants [289]. Thus, biotechnological processes based on the use of algae can be employed in production of new sources of phytohormones for agriculture, as well as functional additives to human diet and animal feed.

Conclusion

The products of industrial biotechnology play an important role in various fields of human activity. Despite the active use of plants and animals in medical, cosmetic, agricultural and food industries, it is the microbiological area that is the key player in industrial biotechnology. It is connected with obtaining various biotechnological products using microbiological approaches: antibiotics, enzymes, biopreparations based on microbial biomass, amino acids, feed proteins, bioethanol, biogases, vitamins, organic acids, polysaccharides, as well as bioproducts obtained on the basis of recombinant DNA technology (hormones, biologically active substances, etc.). Industrial biotechnology is actively developing; it contributes to the identification of new microorganisms-producers, improvement of technological processes and creation of innovative products. At the same time, the tasks of increasing the resistance of microorganisms, optimizing fermentation processes and reducing production costs remain relevant. Industrial microbial biotechnology has a significant potential to stimulate economic growth, improve public health and environmental protection. Continued scientific research in this area is of fundamental importance.

Conflict of interest

            The authors declare no conflict of interest.

References

1. Mordor Intelligence. Biotech market - Growth, trends, COVID-19 impact, and forecasts (2021 - 2026). URL: https://www.mordorintelligence.com
2. Nielsen J., Tillegreen C. B., Petranovic D. Innovation trends in industrial biotechnology. Trends Biotechnol., 40 (10), 1160–1172 (2022). DOI: 10.1016/j.tibtech.2022.03.007
3. Biotechnology in Russia: market, trends and forecasts. 2021. URL: RBC.
4. Aleksandrova E. V. V., Polskova N. V. Development of biotechnology as a factor of economic security of Russia. Vestnik Technosphernoy Bezopasnosty i Selskogo Razvitiya, No. 1 (30), 2022. URL:: https://cyberleninka.ru/article/n/razvitie-biotehnologiy-kak-faktor-ekonomicheskoy-bezopasnosti-rossii
5. Lian J., HamediRad M., Zhao H. Advancing Metabolic Engineering of Saccharomyces cerevisiae Using the CRISPR/Cas System. J., 13 (9), e1700601 (2018). DOI: 10.1002/biot.201700601
6. Zabed H., Sahu J. N., Suely A., Boyce A. N., Faruq G. Bioethanol production from renewable sources: Current perspectives and technological progress. Sustain. Energy Rev., 71, 475–501 (2017). DOI: 10.1016/j.rser.2016.12.076
7. Khan, M. I., Shin, J. H., Kim, J. D. The promising future of microalgae: current status, challenges, and optimization of a sustainable and renewable industry for biofuels, feed, and other products. Microbial Cell Factories, 17(1), 36 (2018). DOI: 10.1186/s12934-018-0879-x
8. López N. I., Pettinari M. J., Nikel P. I., Méndez B. S. Polyhydroxyalkanoates: Much More than Biodegradable Plastics. Adv. Appl. Microbiol., 93, 73–106 (2015). DOI: 10.1016/bs.aambs.2015.06.001
9. Sharipova A. R. Methods of isolation and purification of biotechnological products.Science Time, №9 (33), 2016. URL: https://cyberleninka.ru/article/n/metody-vydeleniya-i-ochistki-biotehnologicheskoy-produktsii
10. Chen G.-Q., Patel M. K. Plastics derived from biological sources: present and future: a technical and environmental review. Rev., 112 (4), 2082–2099 (2012). DOI: 10.1021/cr200162d
11. Możejko-Ciesielska J., Kiewisz R. Bacterial polyhydroxyalkanoates: Still fabulous? Res., 192, 271–282 (2016). DOI: 10.1016/j.micres.2016.07.010
12. Hammami K., Souissi Y., Cherif A., Neifar M. Challenges for polyhydroxyalkanoates production: extremophilic bacteria, waste streams, and optimization strategies. MedCrave Online J. Appl. Bionics Biomech., 7 (1), 101–107 (2023). DOI: 10.15406/mojabb.2023.07.00181
13. Freitas F., Alves V. D., Reis M. A. Advances in bacterial exopolysaccharides: from production to biotechnological applications. Trends Biotechnol., 29 (8), 388–398 (2011). DOI: 10.1016/j.tibtech.2011.03.008
14. Kang H. K., Nguyen T. T. H., Jeong H. N., et al. Molecular cloning and characterization of a novel glucansucrase from Leuconostoc mesenteroides subsp. mesenteroides LM34. Bioprocess Eng., 19, 605–612 (2014). DOI: 10.1007/s12257-014-0116-3
15. Rehm B. H. Bacterial polymers: biosynthesis, modifications and applications. Rev. Microbiol., 8 (8), 578–592 (2010). DOI: 10.1038/nrmicro2354
16. Mangolim C. S., Silva T. T. d., Fenelon V. C., Koga L. N., Ferreira S. B. d. S., Bruschi M. L., et al. Description of recovery method used for curdlan produced by Agrobacterium sp. IFO 13140 and its relation to the morphology and physicochemical and technological properties of the polysaccharide. PLoS ONE, 12 (2), e0171469 (2017). DOI: 10.1371/journal.pone.0171469.
17. Rinaudo M. Chitin and chitosan: Properties and applications. Prog. Polym. Sci., 31 (7), 603–632 (2006). DOI: 10.1016/j.progpolymsci.2006.06.001
18. Novak M., Vetvicka V. Beta-glucans, history, and the present: immunomodulatory aspects and mechanisms of action. Immunotoxicol., 5 (1), 47–57 (2008). DOI: 10.1080/15476910802019045
19. Spring P., Wenk C., Dawson K. A., Newman K. E. The effects of dietary mannaoligosaccharides on cecal parameters and the concentrations of enteric bacteria in the ceca of salmonella-challenged broiler chicks. Sci., 79 (2), 205–211 (2000). DOI: 10.1093/ps/79.2.205
20. Singh R. S., Saini G. K., Kennedy J. F. Pullulan: Microbial sources, production and applications. Polym., 73 (4), 515–531 (2008). DOI: 10.1016/j.carbpol.2008.01.003
21. Farina J. I., Sineriz F., Molina O. E., Perotti N. I. High scleroglucan production by Sclerotium rolfsii: Influence of medium composition. Lett., 23 (9), 717–720 (2001). DOI: 10.1023/A:1005351123156
22. Giavasis I. Bioactive fungal polysaccharides as potential functional ingredients in food and nutraceuticals. Opin. Biotechnol., 26, 162–173 (2014). DOI: 10.1016/j.copbio.2014.01.010
23. Selvasekaran P., Mahalakshmi, Roshini F., Angalene L. A., Chandini, Sunil T., Chidambaram R. Fungal Exopolysaccharides: production and biotechnological industrial applications in food and allied sectors. In: Yadav A. N. (ed.) Recent Trends in Mycological Research. Springer International Publishing, pp. 311–357 (2021).
24. Draget K. I., Smidsrød O., Skjåk-Bræk G. Alginates from algae. Biopolymers Online (2005). DOI: 10.1002/3527600035.BPOL6008
25. Necas J., Bartosikova L. Carrageenan: a review. Med., 58 (4), 187–205 (2013).
26. Keasling J. D. Manufacturing molecules through metabolic engineering. Science, 330 (6009), 1355–1358 (2010). DOI: 10.1126/science.1193990
27. Jackson C. J., Gillam E. M. J., Ollis D. L. Directed evolution of enzymes. In: Lew M., Hung-Wen L. (eds.) Comprehensive Natural Products II, pp. 723–749. Oxford: Elsevier (2010). DOI: 10.1016/B978-0-08-102690-8.00675-8
28. Gupta R., Gupta N., Rathi P. Bacterial lipases: an overview of production, purification and biochemical properties. Microbiol. Biotechnol., 64 (6), 763–781 (2004). DOI: 10.1007/s00253-004-1568-8
29. Schallmey M., Singh A., Ward O. P. Developments in the use of Bacillus species for industrial production. J. Microbiol., 50 (1), 1–17 (2004). DOI: 10.1139/w03-076
30. Bala A., Singh B. Development of an environmental-benign process for efficient pretreatment and saccharification of Saccharum biomasses for bioethanol production. Energy, 130, 12–24 (2019). DOI: 10.1016/j.renene.2018.06.033
31. Singhania R. R., Sukumaran R. K., Patel A. K., Larroche C., Pandey A. Advancement and comparative profiles in the production technologies using solid-state and submerged fermentation for microbial cellulases. Enzyme Microb. Technol., 46 (7), 541–549 (2010). DOI: 10.1016/j.enzmictec.2010.03.010
32. Pandey A., Selvakumar P., Soccol C. R., Nigam P. S. Solid state fermentation for the production of industrial enzymes. Sci., 77, 149–162 (1999).
33. Jørgensen H., et al. Production of cellulases and hemicellulases by three Penicillium species: Effect of substrate and evaluation of cellulase adsorption by capillary electrophoresis. Enzyme Microb. Technol., 36 (1), 42–48 (2005). DOI: 10.1016/j.enzmictec.2004.03.023
34. Chojnacka K., Noworyta A. Evaluation of Spirulina sp. growth in photoautotrophic, heterotrophic and mixotrophic cultures. Enzyme Microb. Technol., 34 (5), 461–465 (2004). DOI: 10.1016/j.enzmictec.2003.12.002
35. Markou G., Nerantzis E. Microalgae for high-value compounds and biofuels production: a review with focus on cultivation under stress conditions. Adv., 31 (8), 1532–1542 (2013). DOI: 10.1016/j.biotechadv.2013.07.011
36. Kirk O., Borchert T. V., Fuglsang C. C. Industrial enzyme applications. Opin. Biotechnol., 13 (4), 345–351 (2002). DOI: 10.1016/S0958-1669(02)00328-2
37. Schäfer T., Borchert T. W., Nielsen V. S., et al. Industrial enzymes. Adv. Biochem. Eng. Biotechnol., 105, 59–131 (2007). DOI: 10.1007/10_2006_039
38. Tolkacheva A.A., Cherenkov D.A., Korneeva O.S., Ponomarev P.G. Industrial enzymes - review of the market of enzyme preparations and prospects for its development. Bulletin of Voronezh State University of Engineering Technologies,79 (4 (74)), 197-203. (2017).
39. Lukin A.A., Danilov M.B., Pirozhinsky S.G. Peculiarities of microbial enzymes application in branches of production sphere. International Research Journal. 8 (98), 94-98 (2020) DOI: 10.23670/IRJ.2020.98.8.013
40. Maurer K.-H. Detergent proteases. Curr. Opin. Biotechnol., 15, 330–334 (2004). DOI: 10.1016/j.copbio.2004.06.005
41. Estell D. A., Graycar T. P., Wells J. A. Engineering an enzyme by site-directed mutagenesis to be resistant to chemical oxidation. Biol. Chem., 260 (11), 6518–6521 (1985).
42. Bryan P. N. Protein engineering of subtilisin. Biochim. Biophys. Acta, 1543 (2), 203–222 (2000). DOI: 10.1016/S0167-4838(00)00235-1
43. Sumantha A., Larroche C., Pandey A. Microbiology and industrial biotechnology of food-grade proteases: A perspective. Food Technol. Biotechnol., 44, 211–220 (2006).
44. Koka R., Weimer B. C. Investigation of the ability of a purified protease from Pseudomonas fluorescens r098 to hydrolyze bitter peptides from cheese. Dairy J., 10, 75–79 (2000). DOI: 10.1016/S0958-6946(00)00023-6
45. Cheng S. W., Hu H. M., Shen S. W., Takagi H., Asano M., Tsai, Y. C. Production and characterization of keratinase of a feather-degrading Bacillus licheniformis PWD-1. Biotechnol. Biochem., 59 (12), 2239–2243 (1995). DOI: 10.1271/bbb.59.2239
46. Arunachalam C., Saritha K. Protease enzyme: An eco-friendly alternative for leather industry. Indian J. Sci. Technol., 12, 29–32 (2009). DOI: 10.17485/ijst/2009/v2i12/29553
47. Rai S. K., Roy J. K., Mukherjee A. K. Characterisation of a detergent-stable alkaline protease from a novel thermophilic strain Paenibacillus tezpurensis sp. nov. AS-S24-II. Microbiol. Biotechnol., 85 (5), 1437–1450 (2010). DOI: 10.1007/s00253-009-2145-y
48. Doukyu N., Ogino H. Organic solvent-tolerant enzymes. Biochem. Eng. J., 48, 270–282 (2010). DOI: 10.1016/j.bej.2009.09.009
49. Brockman H. L. Lipases. Encycl. Chem., 729–732 (2013). DOI: 10.1016/B978-0-12-378630-2.00118-3
50. Reis P., Holmberg K., Watzke H., Leser M. E., Miller R. Lipases at interfaces: a review. Colloid Interface Sci., 147-148, 237–250 (2009). DOI: 10.1016/j.cis.2008.06.001
51. Gotor-Fernández V., Vicente G. Use of Lipases in Organic Synthesis. In: Polaina, J., MacCabe, A.P. (eds) Industrial Enzymes. Springer, Dordrecht. (2007). DOI: 10.1007/1-4020-5377-0_18
52. Hasan F., Shah A. A., Hameed A. Industrial applications of microbial lipases. Enzyme Microb. Technol., 39 (2), 235–251 (2006). DOI: 10.1016/j.enzmictec.2005.10.016
53. Reetz M. T. Lipases as practical biocatalysts. Curr. Opin. Chem. Biol., 6 (2), 145–150 (2002). DOI: 10.1016/S1367-5931(02)00297-1
54. Bajaj A., Lohan P., Jha P. N., Mehrotra R. Biodiesel production through lipase catalyzed transesterification: An overview. Mol. Catal. B Enzym., 62 (1), 9–14 (2010). DOI: 10.1016/j.molcatb.2009.09.018
55. Tan T., Lu J., Nie K., Deng L., Wang F. Biodiesel production with immobilized lipase: A review. Adv., 28 (5), 628–634 (2010). DOI: 10.1016/j.biotechadv.2010.05.012
56. Bijttebier A., Goesaert H., Delcour J. A. Amylase action pattern on starch polymers. Biologia, 63, 989–999 (2008). DOI: 10.2478/s11756-008-0169-x
57. van der Maarel M. J., van der Veen B., Uitdehaag J. C., Leemhuis H., Dijkhuizen L. Properties and applications of starch-converting enzymes of the alpha-amylase family. Biotechnol., 94 (2), 137–155 (2002). DOI: 10.1016/S0168-1656(01)00407-2
58. de Souza P. M., de Oliveira Magalhães P. Application of microbial α-amylase in industry - A review. J. Microbiol., 41 (4), 850–861 (2010). DOI: 10.1590/S1517-83822010000400004
59. Nielsen J. E., Borchert T. V. Protein engineering of bacterial alpha-amylases. Biophys. Acta, 1543 (2), 253–274 (2000). DOI: 10.1016/S0167-4838(00)00240-5
60. Singh R. S., Saini G. K., Kennedy J. F. Maltotriose syrup preparation from pullulan using pullulanase. Polym., 80, 401–407 (2010). DOI: 10.1016/j.carbpol.2009.11.040
61. Jayani R. S., Saxena S., Gupta R. Microbial pectinolytic enzymes: A review. Process Biochem., 40 (9), 2931–2944 (2005). DOI: 10.1016/j.procbio.2005.03.026
62. Jacob N. Pectinolytic Enzymes. In: Singh nee’ Nigam P., Pandey A. (eds) Biotechnology for Agro-Industrial Residues Utilisation. Springer, Dordrecht (2009). DOI: 10.1007/978-1-4020-9942-7_21
63. Kashyap D. R., Vohra P. K., Chopra S., Tewari R. Applications of pectinases in the commercial sector: a review. Technol., 77 (3), 215–227 (2001). DOI: 10.1016/S0960-8524(00)00118-8
64. Hoondal G. S., Tiwari R. P., Tewari R., Dahiya N., Beg Q. K. Microbial alkaline pectinases and their industrial applications: a review. Microbiol. Biotechnol., 59 (4-5), 409–418 (2002). DOI: 10.1007/s00253-002-1061-1
65. Beg Q. K., Kapoor M., Mahajan L., Hoondal G. S. Microbial xylanases and their industrial applications: a review. Microbiol. Biotechnol., 56 (3-4), 326–338 (2001). DOI: 10.1007/s002530100704
66. Subramaniyan S., Prema P. Biotechnology of microbial xylanases: enzymology, molecular biology, and application. Rev. Biotechnol., 22 (1), 33–64 (2002). DOI: 10.1080/07388550290789450
67. Butt M. S., Tahir-Nadeem M., Ahmad Z., Sultan M. T. Xylanases and their applications in baking industry. Food Technol. Biotechnol., 46 (1), 22–31 (2008).
68. Matsumura S., Sakiyama K., Toshima K. Preparation of octyl β-D-xylobioside and xyloside by xylanase-catalyzed direct transglycosylation reaction of xylan and octanol. Lett., 21 (1), 17–22 (1999). DOI: 10.1023/A:1005464025881
69. Rodríguez Couto S., Toca Herrera J. L. Industrial and biotechnological applications of laccases: a review. Adv., 24 (5), 500–513 (2006). DOI: 10.1016/j.biotechadv.2006.04.003
70. Yokoyama K., Nio N., Kikuchi Y. Properties and applications of microbial transglutaminase. Microbiol. Biotechnol., 64 (4), 447–454 (2004). DOI: 10.1007/s00253-003-1539-5
71. Zhu Y., Rinzema A., Tramper J., Bol J. Microbial transglutaminase—a review of its production and application in food processing. Microbiol. Biotechnol., 44, 277–282 (1995). DOI: 10.1007/BF00169916
72. Gembeh S.V., Jr H.M.F., Taylor M. M., Brown E.M., Marmer W.N. Application of transglutaminase to derivatize proteins: 1. Studies on soluble proteins and preliminary results on wool. J. Sci. Agric. 85, 418–424, (2005) DOI: 10.1002/jsfa.1999
73. Lei X. G., Porres J. M., Mullaney E. J., Brinch-Pedersen H. Phytase: Source, Structure and Application. In: Polaina J., MacCabe A. P. (eds) Industrial Enzymes. Springer, Dordrecht (2007). DOI: 10.1007/1-4020-5377-0_29
74. Ravindran V., Bryden W. L., Kornegay E. T. Phytates: Occurrence, bioavailability and implications in poultry nutrition. Avian Biol. Rev., 6, 125–143 (1995).
75. Maenz D. D. Enzymatic characteristics of phytases as they relate to their use in animal feed. In: Bedford M. R., Partridge G. G. (eds.) Enzymes in Farm Animal Nutrition, pp. 61–84. Wallington: CABI (2001). DOI: 10.1079/9780851993935.0061
76. Hull P. Glucose Syrups: Technology and Applications. Wiley-Blackwell (2010). DOI: 10.1002/9781444314748
77. White J. S. Sucrose, HFCS, and Fructose: History, Manufacture, Composition, Applications, and Production. In: Rippe J. (eds) Fructose, High Fructose Corn Syrup, Sucrose and Health. Nutrition and Health. Humana Press, New York, NY (2014). DOI: 10.1007/978-1-4899-8077-9_2
78. Singh P., Ban Y. G., Kashyap L., Siraree A., Singh J. Sugar and Sugar Substitutes: Recent Developments and Future Prospects. In: Sugar and Sugar Derivatives: Changing Consumer Preferences. Springer, Singapore (2020). DOI: 10.1007/978-981-15-6663-9_4
79. Zargaraan A., Kamaliroosta L., Yaghoubi A. S., Mirmoghtadaie L. Effect of substitution of sugar by high fructose corn syrup on the physicochemical properties of bakery and dairy products: A review. Food Sci. Res., 3 (4), 3–11 (2016). DOI: 10.18869/acadpub.nfsr.3.4.3
80. Песчанская, В. А., Андриевская, Д. В., Ульянова, Е. В. Перспективы использования глюкозно-фруктозных сиропов при производстве спиртных напитков. Пиво и напитки 3, 13–16, (2020). DOI: 10.24411/2072–9650–2020–10033
81. Sano C. History of glutamate production. Am. J. Clin. Nutr., 90 (3), 728S–732S (2009). DOI: 10.3945/ajcn.2009.27462F
82. Cesari M., Rossi G. P., Sticchi D., Pessina A. C. Is homocysteine important as risk factor for coronary heart disease? Metab. Cardiovasc. Dis., 15 (2), 140–147 (2005). DOI: 10.1016/j.numecd.2004.04.002
83. Wu G. Amino acids: metabolism, functions, and nutrition. Amino Acids, 37 (1), 1–17 (2009). DOI: 10.1007/s00726-009-0269-0
84. Paul B. D., Sbodio J. I., Xu R., et al. Cystathionine γ-lyase deficiency mediates neurodegeneration in Huntington's disease. Nature, 509 (7498), 96–100 (2014). DOI: 10.1038/nature13136
85. Wu G., Bazer F. W., Cudd T. A., Meininger C. J., Spencer T. E. Maternal nutrition and fetal development. Nutr., 134 (9), 2169–2172 (2004). DOI: 10.1093/jn/134.9.2169
86. Ikeda M. Amino acid production processes. Adv. Biochem. Eng. Biotechnol., 79, 1–35 (2003). DOI: 10.1007/3-540-45989-8_1
87. Kurihara K. Glutamate: from discovery as a food flavor to role as a basic taste (umami). J. Clin. Nutr., 90 (3), 719S–722S (2009). DOI: 10.3945/ajcn.2009.27462D
88. Renneberg R. High grade cysteine no longer has to be extraxted from hair. In Demain, AL, eds. Biotechnology for beginners. Academic Press: Amsterdam, pp.106 (2008)
89. Sereewatthanawut I., Prapintip S., Watchiraruji K., et al. Extraction of protein and amino acids from deoiled rice bran by subcritical water hydrolysis. Technol., 99 (3), 555–561 (2008). DOI: 10.1016/j.biortech.2006.12.030
90. Klejdus B., Lojková L., Kula E., et al. Supercritical fluid extraction of amino acids from birch (Betula pendula Roth) leaves and their liquid chromatographic determination with fluorimetric detection. Sep. Sci., 31 (8), 1363–1373 (2008). DOI: 10.1002/jssc.200700560
91. Pourali O., Asghari F. S., Yoshida H. Sub-critical water treatment of rice bran to produce valuable materials. Food Chem., 115 (1), 1–7 (2009). DOI: 10.1016/j.foodchem.2008.11.099
92. Zuend S. J., Coughlin M. P., Lalonde M. P., Jacobsen E. N. Scaleable catalytic asymmetric Strecker syntheses of unnatural alpha-amino acids. Nature, 461 (7266), 968–970 (2009). DOI: 10.1038/nature08484
93. Gröger H. Catalytic enantioselective Strecker reactions and analogous syntheses. Rev., 103 (8), 2795–2828 (2003). DOI: 10.1021/cr020038p
94. Singh S., Gogoi B. K., Bezbaruah R. L. Racemic resolution of some DL-amino acids using Aspergillus fumigatus L-amino acid oxidase. Microbiol., 63 (1), 94–99 (2011). DOI: 10.1007/s00284-011-9955-8
95. Hsiao H. Y., Walter J. F., Anderson D. M., Hamilton B. K. Enzymatic production of amino acids. Genet. Eng. Rev., 6, 179–219 (1988). DOI: 10.1080/02648725.1988.10647848
96. Sugimoto M. Amino Acids, Production Processes. In: Encyclopedia of Industrial Biotechnology (2010). DOI: 10.1002/9780470054581.eib025
97. Becker J., Zelder O., Häfner S., et al. From zero to hero—design-based systems metabolic engineering of Corynebacterium glutamicum for L-lysine production. Eng., 13 (2), 159–168 (2011). DOI: 10.1016/j.ymben.2011.01.003
98. Aoki R., Wada M., Takesue N., Tanaka K., Yokota A. Enhanced glutamic acid production by a H+-ATPase-defective mutant of Corynebacterium glutamicum. Biotechnol. Biochem., 69 (8), 1466–1472 (2005). DOI: 10.1271/bbb.69.1466
99. Hermann T. Industrial production of amino acids by coryneform bacteria. Biotechnol., 104 (1-3), 155–172 (2003). DOI: 10.1016/S0168-1656(03)00149-4
100. Zahoor A., Lindner S. N., Wendisch V. F. Metabolic engineering of Corynebacterium glutamicum aimed at alternative carbon sources and new products. Struct. Biotechnol. J., 3, e201210004 (2012). DOI: 10.5936/csbj.201210004
101. Khan N. S., Mishra I. M., Singh R. P., Prasad B. Modeling the growth of Corynebacterium glutamicum under product inhibition in L-glutamic acid fermentation. Eng. J., 25, 173–178 (2005). DOI: 10.1016/j.bej.2005.01.025
102. Park J. H., Lee S. Y. Fermentative production of branched chain amino acids: a focus on metabolic engineering. Microbiol. Biotechnol., 85 (3), 491–506 (2010). DOI: 10.1007/s00253-009-2307-y
103. Sauer M., Porro D., Mattanovich D., Branduardi P. Microbial production of organic acids: expanding the markets. Trends Biotechnol., 26 (2), 100–108 (2008). DOI: 10.1016/j.tibtech.2007.11.006
104. Upadhyaya B. P., DeVeaux L. C., Christopher, L. P. Metabolic engineering as a tool for enhanced lactic acid production. Trends Biotechnol., 32(12), 637–644. (2014) DOI: 10.1016/j.tibtech.2014.10.005
105. Wang B., Rutherfurd-Markwick K., Zhang X. X., Mutukumira A. N. Kombucha: Production and Microbiological Research. Foods, 11(21), 3456 (2022). DOI: 10.3390/foods11213456
106. Cheng K. K., Zhao X. B., Zeng J., et al. Downstream processing of biotechnological produced succinic acid. Microbiol. Biotechnol., 95 (4), 841–850 (2012). DOI: 10.1007/s00253-012-4214-x
107. Wang C., Li Q., Tang H., et al. Membrane fouling mechanism in ultrafiltration of succinic acid fermentation broth. Technol., 116, 366–371 (2012). DOI: 10.1016/j.biortech.2012.03.099
108. Soccol C. R., Vandenberghe L. P., Rodrigues C., Pandey A. New perspectives for citric acid production and application. Food Technol. Biotechnol., 44, 141–149 (2006).
109. Angumeenal A. R., Venkappayya D. An overview of citric acid production. LWT - Food Sci. Technol., 50 (2), 367–370 (2013). DOI: 10.1016/j.lwt.2012.05.016
110. Dhillon G. S., Brar S. K., Kaur S., Verma M. Screening of agro-industrial wastes for citric acid bioproduction by Aspergillus niger NRRL 2001 through solid state fermentation. Sci. Food Agric., 93 (7), 1560–1567 (2013). DOI: 10.1002/jsfa.5920
111. Singh O. V., Sharma A., Singh R. P. Optimisation of fermentation conditions for gluconic acid production by a mutant of Aspergillus niger. Indian J. Exp. Biol., 39 (11), 1136–1143 (2001).
112. Willke T., Vorlop K. D. Biotechnological production of itaconic acid. Microbiol. Biotechnol., 56 (3-4), 289–295 (2001). DOI: 10.1007/s002530100685 114.
113. Peace C. G., O’Neill L. A. The role of itaconate in host defense and inflammation. Clin. Invest., 132 (2), e148548 (2022). DOI: 10.1172/JCI148548
114. Zhao M., Lu X., Zong H., Li J., Zhuge B. Itaconic acid production in microorganisms. Biotech let, 40(3), 455–464 (2018). DOI: 10.1007/s10529-017-2500-5
115. Ilica R. A., Kloetzer L., Galaction A. I., Caşcaval D. Fumaric acid: production and separation. Biotechnol Lett, 41(1), 47–57, (2019). DOI: 10.1007/s10529-018-2628-y
116. Max B., Salgado J. M., Rodríguez N., Cortés S., Converti A., Domínguez J. M. Biotechnological production of citric acid. J. Microb., 41(4), 862–875 (2010). DOI:10.1590/s1517-83822010000400005
117. Slininger P.J., Liu S.C., Dien B.S. Yeast strains for organic acid production. US Patent 7.838.262, 2010.
118. Porro D., Branduardi P. Production of Organic Acids by Yeasts and Filamentous Fungi. In: Sibirny A. (eds) Biotechnology of Yeasts and Filamentous Fungi. Springer, Cham (2017). DOI: 10.1007/978-3-319-58829-2_7
119. Chisti Y. Biodiesel from microalgae. Biotechnol. Adv., 25 (3), 294–306 (2007). DOI: 10.1016/j.biotechadv.2007.02.001
120. Hu Q., Sommerfeld M., Jarvis E., et al. Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J., 54 (4), 621–639 (2008). DOI: 10.1111/j.1365-313X.2008.03492.x
121. Wang B., Li Y., Wu N., Lan C. Q. CO₂ bio-mitigation using microalgae. Microbiol. Biotechnol., 79 (5), 707–718 (2008). DOI: 10.1007/s00253-008-1518-y
122. Griffiths M. J., Harrison S. T. L. Lipid productivity as a key characteristic for choosing algal species for biodiesel production. Appl. Phycol., 21 (5), 493–507 (2009). DOI: 10.1007/s10811-008-9392-7
123. Borowitzka M. A. High-value products from microalgae—their development and commercialisation. Appl. Phycol., 25, 743–756 (2013). DOI: 10.1007/s10811-013-9983-9
124. Mata T. M., Martins A. A., Caetano N. S. Microalgae for biodiesel production and other applications: A review. Sustain. Energy Rev., 14 (1), 217–232 (2010). DOI: 10.1016/j.rser.2009.07.020
125. Lenaz G. The role of lipids in the structure and function of membranes. Biochem., 6, 233–343 (1979). DOI: 10.1007/978-1-4615-7945-8_5
126. Li-Beisson Y., Nakamura Y., Harwood J. Lipids: From Chemical Structures, Biosynthesis, and Analyses to Industrial Applications. Biochem., 86, 1–18 (2016). DOI: 10.1007/978-3-319-25979-6_1
127. Ochsenreither K., Glück C., Stressler T., Fischer L., Syldatk C. Production Strategies and Applications of Microbial Single Cell Oils. Microbiol., 7, 1539 (2016). DOI: 10.3389/fmicb.2016.01539
128. Shah A. M., Yang W., Mohamed H., Zhang Y., Song Y. Microbes: A Hidden Treasure of Polyunsaturated Fatty Acids. Nutr., 9, 827837 (2022). DOI: 10.3389/fnut.2022.827837
129. Ghazani S. M., Marangoni A. G. Microbial Lipids for Foods. Trends Food Sci. Technol., 119, 593–607 (2022). DOI: 10.1016/j.tifs.2021.10.014
130. Lakhawat S. S., Malik N., Kumar V., Kumar S., Sharma P. K. Implications of CRISPR-Cas9 in Developing Next Generation Biofuel: A Mini-review. Protein Pept. Sci., 23 (9), 574–584 (2022). DOI: 10.2174/1389203723666220907110310
131. Dong T., Knoshaug E. P., Pienkos P. T., Laurens L. M. L. Lipid Recovery from Wet Oleaginous Microbial Biomass for Biofuel Production: A Critical Review. Energy, 177, 879–895 (2016). DOI: 10.1016/j.apenergy.2016.06.002
132. Patel A., Karageorgou D., Rova E., et al. An Overview of Potential Oleaginous Microorganisms and Their Role in Biodiesel and Omega-3 Fatty Acid-Based Industries. Microorganisms, 8 (3), 434 (2020). DOI: 10.3390/microorganisms8030434
133. Koreti D., Kosre A., Jadhav S. K., Chandrawanshi N. K. A comprehensive review on oleaginous bacteria: an alternative source for biodiesel production. Bioprocess., 9 (1), 47 (2022). DOI: 10.1186/s40643-022-00527-1
134. Dourou M., Aggeli D., Papanikolaou S., Aggelis G. Critical steps in carbon metabolism affecting lipid accumulation and their regulation in oleaginous microorganisms. Microbiol. Biotechnol., 102 (6), 2509–2523 (2018). DOI: 10.1007/s00253-018-8813-z
135. Mahmood Z., Singh A. K. Rhodococcus opacus high-cell-density batch cultivation with a bagasse hydrolysate for possible triacylglycerol synthesis. Bioprocess., 7, 209–217 (2023). DOI: 10.4103/bbrj.bbrj_55_23
136. Behera B., Unpaprom Y., Ramaraj R., et al. Integrated biomolecular and bioprocess engineering strategies for enhancing the lipid yield from microalgae. Sustain. Energy Rev., 148, 111270 (2021). DOI: 10.1016/j.rser.2021.111270
137. Thanapimmetha A., Suwaleerat T., Saisriyoot M., Chisti Y., Srinophakun P. Production of carotenoids and lipids by Rhodococcus opacus PD630 in batch and fed-batch culture. Bioprocess Biosyst. Eng., 40 (1), 133–143 (2017). DOI: 10.1007/s00449-016-1681-y
138. Ageitos J. M., Vallejo J. A., Veiga-Crespo P., Villa T. G. Oily yeasts as oleaginous cell factories. Microbiol. Biotechnol., 90 (4), 1219–1227 (2011). DOI: 10.1007/s00253-011-3200-z
139. Sargeant L. A., Chuck C. J., Donnelly J., Bannister C. D., Scott R. J. Optimizing the lipid profile, to produce either a palm oil or biodiesel substitute, by manipulation of the culture conditions for Rhodotorula glutinis. Biofuels, 5 (1), 33–43 (2014). DOI: 10.4155/bfs.13.64
140. Zhu L. Y., Zong M. H., Wu H. Efficient lipid production with Trichosporon fermentans and its use for biodiesel preparation. Technol., 99 (16), 7881–7885 (2008). DOI: 10.1016/j.biortech.2008.02.033
141. Chopra J., Sen R. Process optimization involving critical evaluation of oxygen transfer, oxygen uptake and nitrogen limitation for enhanced biomass and lipid production by oleaginous yeast for biofuel application. Bioprocess Biosyst. Eng., 41 (8), 1103–1113 (2018). DOI: 10.1007/s00449-018-1939-7
142. Abeln F., Chuck C. J. The history, state of the art and future prospects for oleaginous yeast research. Cell Fact., 20 (1), 221 (2021). DOI: 10.1186/s12934-021-01712-1
143. Lopes H. J. S., Bonturi N., Kerkhoven E. J., Miranda E. A., Lahtvee P. J. C/N ratio and carbon source-dependent lipid production profiling in Rhodotorula toruloides. Microbiol. Biotechnol., 104 (6), 2639–2649 (2020). DOI: 10.1007/s00253-020-10386-5
144. Zhang H., Wang Z., Sun C., et al. A phospholipid:diacylglycerol acyltransferase is involved in the regulation of phospholipids homeostasis in oleaginous Aurantiochytrium sp. Biofuels Bioprod., 16 (1), 142 (2023). DOI: 10.1186/s13068-023-02396-y
145. Beopoulos A., Nicaud J. M., Gaillardin C. An overview of lipid metabolism in yeasts and its impact on biotechnological processes. Microbiol. Biotechnol., 90 (4), 1193–1206 (2011). DOI: 10.1007/s00253-011-3212-8
146. Hu M., Qiu L., Wang Y. The PHO Pathway Involved in Phosphate Metabolism in Yeast for Efficient Phosphorus Removal. E3S Web Conf., 53, 04023 (2018). DOI: 10.1051/e3sconf/20185304023
147. De Oliva-Neto P., Dorta C., Flavia A., Carvalho A., Gomes De Lima V. M. The Brazilian Technology of Fuel Ethanol Fermentation-Yeast Inhibition Factors and New Perspectives to Improve the Technology. Process. Energy Commun. Curr. Res. Technol. Dev., 1, 371–379 (2013).
148. Kam S., Kenari A. A., Younesi H. Production of Single Cell Protein in Stickwater by Lactobacillus acidophilus and Aspergillus niger. Aquat. Food Prod. Technol., 21, 403–417 (2012). DOI: 10.1080/10498850.2011.605539
149. Isaza-Pérez F., Ramírez-Carmona M., Rendón-Castrillón L., Ocampo-López C. Potential of residual fungal biomass: a review. Sci. Pollut. Res., 27 (12), 13019–13031 (2020). DOI: 10.1007/s11356-020-08193-6
150. Shah A. M., Mohamed H., Fazili A. B. A., Yang W., Song Y. Investigating the Effect of Alcohol Dehydrogenase Gene Knockout on Lipid Accumulation in Mucor circinelloides WJ11. J. Fungi, 8 (9), 917 (2022). DOI: 10.3390/jof8090917
151. Youssef, G.A.; Elrefaey, A.M.; El-Aassar, S. Potential Assessment of Oleaginous Fungi for Sustainable Biodiesel Production: Screening, Identification and Lipid Production Optimization. Preprints, (2020). DOI: 10.21203/rs.3.rs-127210/v1
152. Zhang X. Y., Li B., Huang B. C., et al. Production, Biosynthesis, and Commercial Applications of Fatty Acids From Oleaginous Fungi. Nutr., 9, 873657 (2022). DOI: 10.3389/fnut.2022.873657
153. Dzurendova S., Zimmermann B., Tafintseva V., et al. Metal and Phosphate Ions Show Remarkable Influence on the Biomass Production and Lipid Accumulation in Oleaginous Mucor circinelloides. Fungi, 6 (4), 260 (2020). DOI: 10.3390/jof6040260
154. van Aarle I. M., Olsson P. A. Fungal lipid accumulation and development of mycelial structures by two arbuscular mycorrhizal fungi. Environ. Microbiol., 69 (11), 6762–6767 (2003). DOI: 10.1128/AEM.69.11.6762-6767.2003
155. Athenaki M., Gardeli C., Diamantopoulou P., et al. Lipids from yeasts and fungi: physiology, production and analytical considerations. Appl. Microbiol., 124 (2), 336–367 (2018). DOI: 10.1111/jam.13633
156. Ali H., Zulkali M. D. Design Aspects of Bioreactors for Solid-State Fermentation: A Review. Biochem. Eng., 25, 255–266 (2011).
157. Finkel Z. V., Follows M. J., Liefer J. D., et al. Phylogenetic Diversity in the Macromolecular Composition of Microalgae. PLoS One, 11 (5), e0155977 (2016). DOI: 10.1371/journal.pone.0155977
158. Tibbetts S. M., Milley J. E., Lall S. P. Chemical Composition and Nutritional Properties of Freshwater and Marine Microalgal Biomass Cultured in Photobioreactors. Appl. Phycol., 27, 1109–1119 (2015). DOI: 10.1007/s10811-014-0428-x
159. de Oliveira M. A. C. L., Monteiro M. P. C., Robbs P. G., Leite S. G. F. Growth and Chemical Composition of Spirulina Maxima and Spirulina Platensis Biomass at Different Temperatures. Int., 7, 261–275 (1999). DOI: 10.1023/A:1009233230706
160. El-fayoumy E. A., Ali H. E. A., Elsaid K., Elkhatat A., Al-Meer S., Zul Helmi Rozaini M., Azmuddin Abdullah M. Co-production of high-density biomass and high-value compounds via two-stage cultivation of Chlorella vulgaris using light intensity and a combination of salt stressors. Biomass Convers. Biorefin., 14, 22673–22686 (2023). DOI: 10.1007/s13399-023-04442-z
161. Xue S. J., Chi Z., Zhang Y., Li Y. F., Liu G. L., Jiang H., Hu Z., Chi Z. M. Fatty acids from oleaginous yeasts and yeast-like fungi and their potential applications. Rev. Biotechnol., 38 (7), 1049–1060 (2018). DOI: 10.1080/07388551.2018.1428167
162. Fan Y. Y., Chapkin R. S. Importance of dietary gamma-linolenic acid in human health and nutrition. Nutr., 128 (9), 1411–1414 (1998). DOI: 10.1093/jn/128.9.1411
163. Venkatakrishnan P., Palanisamy P. Synthesis and Characterization of Myristic Acid---Infused Al₂O₃/CuO/MWCNT Nanocomposites for Energy Storage and Cooling Applications. Scr., 99 (2024).
164. Patel A., Karageorgou D., Rova E., Katapodis P., Rova U., Christakopoulos P., Matsakas L. An Overview of Potential Oleaginous Microorganisms and Their Role in Biodiesel and Omega-3 Fatty Acid-Based Industries. Microorganisms, 8 (3), 434 (2020). DOI: 10.3390/microorganisms8030434
165. Fahy E., Subramaniam S., Brown H. A., Glass C. K., Merrill A. H., Jr Murphy R. C., Raetz C. R., Russell D. W., Seyama Y., Shaw W., Shimizu T., Spener F., van Meer G., VanNieuwenhze M. S., White S. H., Witztum J. L., Dennis E. A. A comprehensive classification system for lipids. Lipid Res., 46 (5), 839–861 (2005). DOI: 10.1194/jlr.E400004-JLR200
166. Yellapu S. K., Bharti, Kaur R., et al. Recent developments of downstream processing for microbial lipids and conversion to biodiesel. Technol., 256, 515–528 (2018). DOI: 10.1016/j.biortech.2018.01.129
167. Joseph Antony Sundarsingh T., Ameen F., Ranjitha J., Raghavan S., Shankar V. Engineering Microbes for Sustainable Biofuel Production and Extraction of Lipids. Research and Future Perspectives. Fuel, 355, 129532 (2024). DOI: 10.1016/j.fuel.2023.129532
168. Silva J. M. E., Martins L. H. D. S., Moreira D. K. T., Silva L. D. P., Barbosa P. P. M., Komesu A., Ferreira N. R., Oliveira J. A. R. Microbial Lipid Based Biorefinery Concepts: A Review of Status and Prospects. Foods, 12 (10), 2074 (2023). DOI: 10.3390/foods12102074
169. Deng Y., Wang W., Zhao S., Yang X., Xu W., Guo M., Xu E., Ding T., Ye X., Liu D. Ultrasound-Assisted Extraction of Lipids as Food Components: Mechanism, Solvent, Feedstock, Quality Evaluation and Coupled Technologies-A Review. Trends Food Sci. Technol., 122, 83-96, (2022) DOI: 10.1016/j.tifs.2022.01.034
170. Karimi Sani I., Mehrnoosh F., Rasul N. H., Hassani B., Mohammadi H., Gholizadeh H., Sattari N., Kaveh M., Khodaei S. M., Alizadeh Sani M., Eghbaljoo H., Assadpour E., Zhang F., Jafari S. M. Pulsed electric field-assisted extraction of natural colorants; principles and applications. Food Biosci., 61, 104746 (2024). DOI: 10.1016/j.fbio.2024.104746.
171. Carruthers D. N., Lee T. S. Translating advances in microbial bioproduction to sustainable biotechnology. Bioeng. Biotechnol., 10, 968437 (2022). DOI: 10.3389/fbioe.2022.968437
172. Long X., He N., He Y., Jiang J., Wu T. Biosurfactant surfactin with pH-regulated emulsification activity for efficient oil separation when used as emulsifier. Technol., 241, 200–206 (2017). DOI: 10.1016/j.biortech.2017.05.120
173. Béligon V., Christophe G., Fontanille P., Larroche C. Microbial Lipids as Potential Source to Food Supplements. Opin. Food Sci., 7, 35–42 (2016). DOI: 10.1016/j.cofs.2015.10.002
174. Ummalyma S. B., Sirohi R., Udayan A., et al. Sustainable microalgal biomass production in food industry wastewater for low-cost biorefinery products: a review. Rev., 1–23 (2022). DOI: 10.1007/s11101-022-09814-3
175. Tomás-Pejó E., Morales-Palomo S., González-Fernández C. Microbial lipids from organic wastes: Outlook and challenges. Technol., 323, 124612 (2021). DOI: 10.1016/j.biortech.2020.124612
176. Russell A. D. Types of antibiotics and synthetic antimicrobial agents. In: Denyer S. P., Hodges N. A., German S. P. (eds.) Hugo and Russell’s Pharmaceutical Microbiology. 7th Ed. Blackwell Science, UK, pp. 152–186 (2004). DOI: 10.1002/9780470988329.ch10
177. Etebu E., Arikekpar I. Antibiotics: Classification and mechanisms of action with emphasis on molecular perspectives. J. Appl. Microbiol. Biotechnol. Res., 4, 90–101 (2016).
178. Gelpi A., Gilbertson A., Tucker J. D. Magic bullet: Paul Ehrlich, Salvarsan and the birth of venereology. Transm. Infect., 91 (1), 68–69 (2015). DOI: 10.1136/sextrans-2014-051779
179. Mohr K. I. History of Antibiotics Research. Curr. Top. Microbiol. Immunol., 398, 237–272 (2016). DOI: 10.1007/82_2016_499
180. Hare R. New light on the history of penicillin. Hist., 26 (1), 1–24 (1982). DOI: 10.1017/S0025727300040758
181. Raper K. B., Alexander D. F., Coghill R. D. Penicillin: II. Natural Variation and Penicillin Production in Penicillium notatum and Allied Species. Bacteriol., 48 (6), 639–659 (1944). DOI: 10.1128/jb.48.6.639-659.1944
182. Zaffiri L., Gardner J., Toledo-Pereyra L. H. History of antibiotics. From salvarsan to cephalosporins. J. Investig. Surg., 25 (2), 67–77 (2012). DOI: 10.3109/08941939.2012.664099
183. Hutchings M. I., Truman A. W., Wilkinson B. Antibiotics: past, present and future. Opin. Microbiol., 51, 72–80 (2019). DOI: 10.1016/j.mib.2019.10.008
184. Jose P. A., Jha B. New Dimensions of Research on Actinomycetes: Quest for Next Generation Antibiotics. Microbiol., 7, 1295 (2016). DOI: 10.3389/fmicb.2016.01295
185. Ayswaria R., Vijayan J., Nathan V. K. Antimicrobial peptides derived from microalgae for combating antibiotic resistance: Current status and prospects. Cell Biochem. Funct., 41 (2), 142–151 (2023). DOI: 10.1002/cbf.3779
186. Cepas V., Gutiérrez-Del-Río I., López Y., Redondo-Blanco S., Gabasa Y., Iglesias M.J., Soengas R., Fernández-Lorenzo A., López-Ibáñez S., Villar C.J., Martins C.B., Ferreira J.D., Assunção M.F.G., Santos L.M.A., Morais J., Castelo-Branco R., Reis M.A., Vasconcelos V., López-Ortiz F., Lombó F., Soto S.M. Microalgae and Cyanobacteria Strains as Producers of Lipids with Antibacterial and Antibiofilm Activity. Mar. Drugs, 19 (12), 675 (2021). DOI: 10.3390/md19120675
187. Wang W., Wang T.-T. Editorial: Fungal secondary metabolites as valuable chemical entities for medicines and agrochemicals. Microbiol., 14, 1150023 (2023). DOI: 10.3389/fmicb.2023.1150023.
188. Bhowmick S., Mazumdar A., Moulick A., Adam V. Algal metabolites: An inevitable substitute for antibiotics. Adv., 43, 107571 (2020). DOI: 10.1016/j.biotechadv.2020.107571.
189. Calderon C. B., Sabundayo B. P. Antimicrobial classifications: Drugs for bugs. In: Schwalbe R., Steele-Moore L., Goodwin A. C. (eds.) Antimicrobial Susceptibility Testing Protocols. CRC Press, Taylor and Francis Group (2007). DOI: 10.1201/9781420014495.ch2
190. Pancu D. F., Scurtu A., Macasoi I. G., Marti D., Mioc M., Soica C., Coricovac D., Horhat D., Poenaru M., Dehelean C. Antibiotics: Conventional Therapy and Natural Compounds with Antibacterial Activity—A Pharmaco-Toxicological Screening. Antibiotics, 10 (4), 401 (2021). DOI: 10.3390/antibiotics10040401
191. Wright G. D. Q&A: Antibiotic resistance: where does it come from and what can we do about it? BMC Biol., 8, 123 (2010). DOI: 10.1186/1741-7007-8-123
192. Etebu E., Arikekpar I. Antibiotics: Classification and mechanisms of action with emphasis on molecular perspectives. J. Appl. Microbiol. Biotechnol. Res., 4, 90–101 (2016).
193. Ullah H., Ali S. Classification of Anti-Bacterial Agents and Their Functions. Agents (2017).
194. Loree J., Lappin S. L. Bacteriostatic Antibiotics. StatPearls Publishing: Treasure Island, FL, USA (2020). DOI: 10.5772/intechopen.68695
195. Cho H., Uehara T., Bernhardt T. G. Beta-lactam antibiotics induce a lethal malfunctioning of the bacterial cell wall synthesis machinery. Cell, 159 (6), 1300–1311 (2014). DOI: 10.1016/j.cell.2014.11.017
196. Heesemann J. Resistenzmechanismen gegen Betalaktamantibiotika [Mechanisms of resistance to beta-lactam antibiotics]. Infection, 21 (Suppl 1), S4–S9 (1993). DOI: 10.1007/BF01710336
197. James C. W., Gurk-Turner C. Cross-reactivity of beta-lactam antibiotics. (Baylor Univ. Med. Cent.), 14 (1), 106–107 (2001). DOI: 10.1080/08998280.2001.11927741
198. Aharonowitz Y., Cohen G., Martin J. F. Penicillin and cephalosporin biosynthetic genes: structure, organization, regulation, and evolution. Rev. Microbiol., 46, 461–495 (1992). DOI: 10.1146/annurev.mi.46.100192.002333.
199. van den Berg M. A., Albang R., Albermann K., et al. Genome sequencing and analysis of the filamentous fungus Penicillium chrysogenum. Biotechnol., 26 (10), 1161–1168 (2008). DOI: 10.1038/nbt.1498
200. Elander R. P. Industrial production of beta-lactam antibiotics. Appl. Microbiol. Biotechnol., 61 (5-6), 385–392 (2003). DOI: 10.1007/s00253-003-1274-y
201. Mateo C., Abian O., Grazu V., et al. Recent Advances in the Industrial Enzymatic Synthesis of Semi-Synthetic β-Lactam Antibiotics. Chem. Rev. Online, 2 (3), 207–218 (2005). DOI: 10.2174/1567203054065691
202. Barber M. S., Giesecke U., Reichert A., Minas W. Industrial enzymatic production of cephalosporin-based beta-lactams. Biochem. Eng. Biotechnol., 88, 179–215 (2004). DOI: 10.1007/b99261
203. DeModena J. A., Gutiérrez S., Velasco J., et al. The production of cephalosporin C by Acremonium chrysogenum is improved by the intracellular expression of a bacterial hemoglobin. Bio/Technology, 11 (8), 926–929 (1993). DOI: 10.1038/nbt0893-926
204. Rodríguez-Sáiz M., De La Fuente J. L., Barredo J. L. Cephalosporin Production by Fungal Metabolic Engineering. In: Encyclopedia of Industrial Biotechnology: Bioprocess, Bioseparation, and Cell Technology. Wiley (2009). DOI: 10.1002/9780470054581.eib331
205. Johnson D. H., Cunha B. A. Aztreonam. Med. Clin. North Am., 79 (4), 733–743 (1995). DOI: 10.1016/S0025-7125(16)30036-0
206. Leanza W. J., Wildonger K. J., Miller T. W., Christensen B. G. N-Acetimidoyl- and N-formimidoylthienamycin derivatives: antipseudomonal beta-lactam antibiotics. Med. Chem., 22 (12), 1435–1436 (1979). DOI: 10.1021/jm00198a001
207. Kahan F. M., Kropp H., Sundelof J. G., Birnbaum J. Thienamycin: development of imipenem-cilastatin. Antimicrob. Chemother., 12 (Suppl D), 1–35 (1983). DOI: 10.1093/jac/12.suppl_d.1
208. Retsema J., Fu W. Macrolides: structures and microbial targets. Int. J. Antimicrob. Agents, 18 (Suppl 1), S3–S10 (2001). DOI: 10.1016/S0924-8579(01)00401-0
209. Zhu Z. J., Krasnykh O., Pan D., Petukhova V., Yu G., Liu Y., Liu H., Hong S., Wang Y., Wan B., Liang W., Franzblau S. G. Structure-activity relationships of macrolides against Mycobacterium tuberculosis. Tuberculosis, 88 (Suppl 1), S49–S63 (2008). DOI: 10.1016/S1472-9792(08)70036-2
210. Omura S. Macrolide Antibiotics. Chemistry, Biology and Practice. Academic Press, San Diego, CA, USA (2002).
211. Jelić D., Antolović R. From Erythromycin to Azithromycin and New Potential Ribosome-Binding Antimicrobials. Antibiotics, 5 (3), 29 (2016). DOI: 10.3390/antibiotics5030029
212. Garrod L. P. The erythromycin group of antibiotics. Br. Med. J., 2 (5036), 57–63 (1957). DOI: 10.1136/bmj.2.5036.57
213. Fidaxomicin: Difimicin; Lipiarmycin; OPT 80; OPT-80; PAR 101; PAR-101. Drugs R&D, 10 (1), 37–45 (2010). DOI: 10.2165/11537730-000000000-00000
214. Kirst H. A. Macrolide Antibiotics. In: Marinelli F., Genilloud O. (eds) Antimicrobials. Springer, Berlin, Heidelberg (2014). DOI: 10.1007/978-3-642-39968-8_11.
215. Chopra I., Roberts M. Tetracycline antibiotics: mode of action, applications, molecular biology, and epidemiology of bacterial resistance. Mol. Biol. Rev., 65 (2), 232–260 (2001). DOI: 10.1128/MMBR.65.2.232-260.2001
216. Sobin, B. A., Finlay, A. C., Kane, J. H. Terramycin, a new antibiotic. Science, 112(2907), 423 (1950). DOI: 10.1126/science.112.2907.423
217. Goodman J. J. Fermentation and Mutational Development of the Tetracyclines. In: Hlavka J. J., Boothe J. H. (eds) The Tetracyclines. Handbook of Experimental Pharmacology, vol 78. Springer, Berlin, Heidelberg (1985). DOI: 10.1007/978-3-642-70304-1_2
218. Vardanyan R. S., Hruby V. J. Antibiotics. In: Synthesis of Essential Drugs, pp. 425–498 (2006). DOI: 10.1016/B978-044452166-8/50032-7
219. Fuoco D. Classification Framework and Chemical Biology of Tetracycline-Structure-Based Drugs. Antibiotics, 1 (1), 1–13 (2012). DOI: 10.3390/antibiotics1010001
220. Zhanel G., Critchley I., Lin L. Y., Alvandi N. Microbiological Profile of Sarecycline, a Novel Targeted Spectrum Tetracycline for the Treatment of Acne Vulgaris. Agents Chemother., 63 (1), e01297-18 (2018). DOI: 10.1128/AAC.01297-18
221. Ramachanderan R., Schaefer B. Tetracycline antibiotics. ChemTexts, 7, 18 (2021). DOI: 10.1007/s40828-021-00138-x
222. Davies J. E. Aminoglycosides: ancient and modern. Antibiot., 59 (9), 529–532 (2006). DOI: 10.1038/ja.2006.73
223. Kirst H. A., Marinelli F. Aminoglycoside Antibiotics. In: Marinelli F., Genilloud O. (eds) Antimicrobials. Springer, Berlin, Heidelberg (2014). DOI: 10.1007/978-3-642-39968-8_10
224. Mahajan G. B., Balachandran L. Antibacterial agents from actinomycetes - a review. Biosci. (Elite Ed.), 4 (1), 240–253 (2012). DOI: 10.2741/373
225. Bulik C. C., Nightingale C. H., Nicolau D. P. Aminoglycosides. In: Vinks A., Derendorf H., Mouton J. (eds) Fundamentals of Antimicrobial Pharmacokinetics and Pharmacodynamics. Springer, New York, NY (2014). DOI: 10.1007/978-0-387-75613-4_9.
226. Dutton C. J., Haxell M. A., McArthur H. A. I., Wax R. G. (eds.) Peptide Antibiotics: Discovery, Modes of Action and Applications. Marcel Dekker (2002). DOI: 10.1201/9780203910801
227. Jenssen H., Hamill P., Hancock R. E. Peptide antimicrobial agents. Microbiol. Rev., 19 (3), 491–511 (2006). DOI: 10.1128/CMR.00056-05
228. Magana M., Pushpanathan M., Santos A. L., et al. The value of antimicrobial peptides in the age of resistance. Lancet Infect. Dis., 20 (9), e216–e230 (2020). DOI: 10.1016/S1473-3099(20)30327-3
229. Van Epps H. L. René Dubos: unearthing antibiotics. J. Exp. Med., 203 (2), 259 (2006). DOI: 10.1084/jem.2032fta
230. Bricas E., Fromageot C. Naturally occurring peptides. Adv. Protein Chem., 8, 1–125 (1953). DOI: 10.1016/S0065-3233(08)60091-1
231. Satlin M. J., Jenkins S. G. Polymyxins. In: Infectious Diseases (4th Ed.), Cohen J., Powderly W. G., Opal S. M. (eds.), Elsevier, pp. 1285–1288.e2 (2017). DOI: 10.1016/B978-0-7020-6285-8.00151-9
232. Eliopoulos G. M., Thauvin C., Gerson B., Moellering R. C. Jr. In vitro activity and mechanism of action of A21978C1, a novel cyclic lipopeptide antibiotic. Agents Chemother., 27 (3), 357–362 (1985). DOI: 10.1128/AAC.27.3.357
233. Sauermann R., Rothenburger M., Graninger W., Joukhadar C. Daptomycin: a review 4 years after first approval. Pharmacology, 81 (2), 79–91 (2008). DOI: 10.1159/000109868
234. Kahne D., Leimkuhler C., Lu W., Walsh C. Glycopeptide and lipoglycopeptide antibiotics. Rev., 105 (2), 425–448 (2005). DOI: 10.1021/cr030103a
235. Allen N. E. From vancomycin to oritavancin: the discovery and development of a novel lipoglycopeptide antibiotic. Agents Med. Chem., 9, 23–47 (2010). DOI: 10.2174/187152110790886745
236. Rossolini G. M., Arena F., Pollini S. Novel Infectious Diseases and Emerging Gram-Positive Multi-Resistant Pathogens in Hospital and Community Acquired Infections. In: Marinelli F., Genilloud O. (eds) Antimicrobials. Springer, Berlin, Heidelberg (2014). DOI: 10.1007/978-3-642-39968-8_2
237. Van Bambeke F. Glycopeptides and glycodepsipeptides in clinical development: a comparative review of their antibacterial spectrum, pharmacokinetics and clinical efficacy. Opin. Investig. Drugs, 7 (8), 740–749 (2006).
238. van der Beek C. P., Roels J. A. Penicillin production: biotechnology at its best. Antonie van Leeuwenhoek, 50 (5-6), 625–639 (1984). DOI: 10.1007/BF02386230
239. Hook D. J. Production of antibiotics by fermentation. In: Ratledge C., Kristiansen B. (eds) Basic Biotechnology. Cambridge University Press, pp. 433–456 (2006). DOI: 10.1017/CBO9780511802409.020
240. Najafpour G. D. Industrial Microbiology. In: Biochemical Engineering and Biotechnology, pp. 1–13 (2007). DOI: 10.1016/B978-044452845-2/50001-X
241. Моисеев Д. В., Лукашов Р. И., Веремчук О. А., Моисеева А. М. Фармацевтическая биотехнология: пособие. Витебск: ВГМУ (2019).
242. Smith J. E. Concepts of Industrial Antibiotic Production. In: Alani D. I., Moo-Young M. (eds) Perspectives in Biotechnology and Applied Microbiology. Springer, Dordrecht (1986). DOI: 10.1007/978-94-009-4321-6_9
243. Fayerman J. New Developments in Gene Cloning in Antibiotic Producing Microorganisms. Biotechnol., 4, 786–789 (1986). DOI: 10.1038/nbt0986-786
244. Newbert R. W., Barton B., Greaves P., Harper J., Turner G. Analysis of a commercially improved Penicillium chrysogenum strain series: involvement of recombinogenic regions in amplification and deletion of the penicillin biosynthesis gene cluster. Ind. Microbiol. Biotechnol., 19 (1), 18–27 (1997). DOI: 10.1038/sj.jim.2900411
245. Dahlmann T. A., Böhm J., Becker K., Kück U. Sexual recombination as a tool for engineering industrial Penicillium chrysogenum strains. Genet., 61 (4), 679–683 (2015). DOI: 10.1007/s00294-015-0497-7
246. Bibb M. J. Regulation of secondary metabolism in streptomycetes. Opin. Microbiol., 8 (2), 208–215 (2005). DOI: 10.1016/j.mib.2005.02.016
247. Begunova A. V., Rozhkova I. V., Shirshova T. I., Glazunova O. A., Fedorova T. V. Optimization of Cultivation Conditions for the Lactobacillus reuteri LR1 Strain to Improve the Biosynthesis of Bacteriocin-Like Substances. Biochem. Microbiol., 56, 920–929 (2020). DOI: 10.1134/S0003683820090033
248. Bisht D., Iqbal Z. Lyophilization - Process and Optimization for Pharmaceuticals. J. Drug Regul. Aff., 3 (1), 30–40 (2018). DOI: 10.22270/ijdra.v3i1.156
249. Просеков А. Ю., Кригер О. В., Дышлюк Л. С., Асякина Л. К. Промышленное производство биологически активных веществ: учебное пособие. Кемерово: КемГУ (2020).
250. Mani I. Microbial Production of Vitamins. In: Singh V., Singh A., Bhargava P., Joshi M., Joshi C. (eds) Engineering of Microbial Biosynthetic Pathways. Springer, Singapore (2020). DOI: 10.1007/978-981-15-2604-6_9.
251. You J., Pan X., Yang C., Du Y., Osire T., Yang T., Zhang X., Xu M., Xu G., Rao Z. Microbial production of riboflavin: Biotechnological advances and perspectives. Eng., 68, 46–58 (2021). DOI: 10.1016/j.ymben.2021.08.009.
252. Averianova L. A., Balabanova L. A., Son O. M., Podvolotskaya A. B., Tekutyeva L. A. Production of Vitamin B₂ (Riboflavin) by Microorganisms: An Overview. Bioeng. Biotechnol., 8, 570828 (2020). DOI: 10.3389/fbioe.2020.570828
253. Yu S., Zheng B., Chen Z., Huo Y. X. Metabolic engineering of Corynebacterium glutamicum for producing branched chain amino acids. Cell Fact., 20 (1), 230 (2021). DOI: 10.1186/s12934-021-01721-0.
254. Martens J. H., Barg H., Warren M. J., Jahn D. Microbial production of vitamin B₁₂. Microbiol. Biotechnol., 58 (3), 275–285 (2002). DOI: 10.1007/s00253-001-0902-7
255. Yuan P., Cui S., Li J., Du G., Chen J., Liu L. Microbial Production of Vitamins. In: Liu L., Chen J. (eds) Systems and Synthetic Biotechnology for Production of Nutraceuticals. Springer, Singapore (2019). DOI: 10.1007/978-981-15-0446-4_7.
256. Leonardi R., Jackowski S. Biosynthesis of Pantothenic Acid and Coenzyme A. EcoSal Plus, 2 (2), 10.1128/ecosalplus.3.6.3.4 (2007). DOI: 10.1128/ecosalplus.3.6.3.4.
257. Barreiro C., Barredo J. L. Carotenoids Production: A Healthy and Profitable Industry. In: Barreiro C., Barredo J. L. (eds) Microbial Carotenoids. Methods in Molecular Biology, 1852. Humana Press, New York, NY (2018). DOI: 10.1007/978-1-4939-8742-9_2.
258. Agostoni C., Berni Canani R., Fairweather-Tait S., Heinonen M., Korhonen H., La Vieille S., Marchelli R., Martin A., Naska A., Neuhäuser-Berthold M., Nowicka G., Sanz Y., Siani A., Sjödin A., Stern M., Strain S. (J.J.), Tetens I., Tomé D., Turck D., Verhagen H. Scientific Opinion on the safety of vitamin D-enriched UV-treated baker's yeast. EFSA J., 12 (1), 3520 (2014). DOI: 10.2903/j.efsa.2014.3520.
259. Ilczuk Z. Studies on citric acid synthesis. 3. The synthesis of vitamins B by strains of Aspergillus niger of different acid production activity. Acta Microbiol. Pol., 14 (3), 337–339 (1965).
260. EFSA Panel on Additives and Products or Substances used in Animal Feed (FEEDAP); Bampidis V, Azimonti G, Bastos ML, Christensen H, Dusemund B, Durjava MF, Kouba M, López-Alonso M, López Puente S, Marcon F, Mayo B, Pechová A, Petkova M, Ramos F, Sanz Y, Villa RE, Woutersen R, Brantom PG, Cocconcelli PS, Glandorf B, Herman L, Maradona MP, Saarela M, Svensson K, Tosti L, Galobart J, Manini P, Pettenati E, Pizzo F, Tarrés-Call J, Anguita M. Safety and efficacy of the feed additive consisting of Vitamin B₂/Riboflavin produced by Eremothecium ashbyi CCTCCM 2019833 for all animal species (Hubei Guangji Pharmaceutical Co., Ltd). EFSA J., 19 (3), e06462 (2021). DOI: 10.2903/j.efsa.2021.6462
261. Kogan L. M., Obol'nikova E. A., Luka V. T., Vetsozola A. O., Samokhvalov G. I. Ubikhinon-9 i érgosterin iz drozhzheĭ Candida paralipolytica [Ubiquinone-9 and ergosterol from Candida paralipolytica yeasts]. Biokhim. Mikrobiol., 21 (1), 78–79 (1985).
262. Choudhari S., Singhal R. Media optimization for the production of beta-carotene by Blakeslea trispora: a statistical approach. Technol., 99 (4), 722–730 (2008). DOI: 10.1016/j.biortech.2007.01.044.
263. Zagoskina N. V., Nazarenko L. V., Kalashnikova E. A., Zhivukhina E. A. Biotechnology. Moscow: Onyx (2009)).
264. Del Mondo A., Smerilli A., Sané E., Sansone C., Brunet C. Challenging microalgal vitamins for human health. Cell Fact., 19 (1), 201 (2020). DOI: 10.1186/s12934-020-01459-1
265. Croft M. T., Warren M. J., Smith A. G. Algae need their vitamins. Cell, 5 (8), 1175–1183 (2006). DOI: 10.1128/EC.00097-06
266. Becker E. W. Micro-algae as a source of protein. Biotechnol. Adv., 25 (2), 207–210 (2007). DOI: 10.1016/j.biotechadv.2006.11.002.
267. Vicente A. R., Manganaris G. A., Sozzi G. O., Crisosto C. H. Nutritional Quality of Fruits and Vegetables. In: Postharvest Handling, pp. 57–106 (2009). DOI: 10.1016/B978-0-12-374112-7.00005-6
268. Biotechnology: textbook for students of higher educational institutions studying in agricultural, natural science, pedagogical specialties and master's programs / [I. V. Tikhonov et al.]; ed. by E. S. Voronin. - St. Petersburg: GIORD,, 2008. — 703 с.; ISBN 978-5-98879-072-3.
269. Slyunyaev V.P. Fundamentals of biotechnology. Fundamentals of industrial biotechnology: textbook for bachelors and masters of directions 240100 "Chemical technology" and 241000 "Energy- and resource-saving processes in chemical technology, petrochemistry and biotechnology" St. Petersburg: SPbLGTU,,, 2012
270. Tentsova A. I. Microbiological synthesis of steroids. Moscow: Medicine (2003). С. 112–148.
271. Nadyrova AI, Kosnyrev AS, Ulyanova VV, Dudkina EV, Vershinina VI, Ilyinskaya ON Efficiency of expression systems based on Escherichia coli and Bacillus subtilis to obtain mutants of ribonuclease binase // Molecular Biology.- - Т. 57. - №5. - C. 807-818. doi: 10.31857/S0026898423050154
272. Itakura K., Hirose T., Crea R., et al. Expression in Escherichia coli of a chemically synthesized gene for the hormone somatostatin. Science, 198 (4321), 1056–1063 (1977). DOI: 10.1126/science.412251
273. Goeddel D. V., Kleid D. G., Bolivar F., et al. Expression in Escherichia coli of chemically synthesized genes for human insulin. Natl. Acad. Sci. U.S.A., 76 (1), 106–110 (1979). DOI: 10.1073/pnas.76.1.106
274. Sereikaite J., Statkute A., Morkunas M., Radzevicius K., Borromeo V., Secchi C., Bumelis V. A. Production of recombinant mink growth hormone in E. coli. Microbiol. Biotechnol., 74 (2), 316–323 (2007). DOI: 10.1007/s00253-006-0673-2.
275. Donova M. V., Egorova O. V. Microbial steroid transformations: Current state and prospects. Microbiol. Biotechnol., 94 (6), 1423–1447 (2012). DOI: 10.1007/s00253-012-4078-0
276. Donova, M.V. Transformation of steroids by actinobacteria: A review. Appl Biochem Microbiol 43, 1–14 (2007). DOI: 10.1134/S0003683807010012
277. Mahato S. B., Mukherjee A. Microbial transformation of testosterone by Aspergillus fumigatus. Steroid Biochem., 21 (3), 341–342 (1984). DOI: 10.1016/0022-4731(84)90289-9.
278. Ríos L. O. L., Luengo J. M., Fernández-Cañón J. M. Steroid 11-Alpha-Hydroxylation by the Fungi Aspergillus nidulans and Aspergillus ochraceus. Methods Mol. Biol., 1645, 271–287 (2017). DOI: 10.1007/978-1-4939-7183-1_19.
279. Lenasi H., Breskvar K. Specific interactions of steroids, arylhydrocarbons and flavonoids with progesterone receptors from the cytosol of the fungus Rhizopus nigricans. Steroid Biochem. Mol. Biol., 91 (4-5), 273–284 (2004). DOI: 10.1016/j.jsbmb.2004.05.003
280. Warner S. A., Sovocool G. W., Domnas A. J. Ergosterol, the unusual dominant sterol of the pythiaceous fungus Zoophagus insidians. Phytochemistry, 21 (8), 2135–2136 (1982). DOI: 10.1016/0031-9422(82)83069-0.
281. Østergaard S., Olsson L., Nielsen J. Metabolic engineering of Saccharomyces cerevisiae. Mol. Biol. Rev., 64 (1), 34–50 (2000). DOI: 10.1128/MMBR.64.1.34-50.2000.
282. Villena G. K., Kitazono A. A., Hernández-Macedo M. L. Bioengineering Fungi and Yeast for the Production of Enzymes, Metabolites, and Value-Added Compounds. In: Hesham A. L., Upadhyay R., Sharma G., Manoharachary C., Gupta V. (eds) Fungal Biotechnology and Bioengineering. Fungal Biology. Springer, Cham (2020). DOI: 10.1007/978-3-030-41870-0_9.
283. Wu N., Wu X., Zhang M., Zhang C., Xu Q. Metabolic engineering of Aspergillus niger for accelerated malic acid biosynthesis by improving NADPH availability. J., 19 (5), e2400014 (2024). DOI: 10.1002/biot.202400014.
284. Bennett J.W., Lasure L.L. More Gene Manipulations in Fungi. 2nd ed. Academic Press, 1991.
285. Ward O. P. Production of recombinant proteins by filamentous fungi. Adv., 30 (5), 1119–1139 (2012). DOI: 10.1016/j.biotechadv.2011.09.012.
286. Nielsen J., Keasling J. D. Engineering Cellular Metabolism. Cell, 164 (6), 1185–1197 (2016). DOI: 10.1016/j.cell.2016.02.004
287. Bittencourt Fagundes M., Wagner R. Sterols Biosynthesis in Algae. IntechOpen (2021). DOI: 10.5772/intechopen.96719.
288. Tang H. F., Yi Y. H., Yao X. S., Xu Q. Z., Zhang S. Y., Lin H. W. Bioactive steroids from the brown alga Sargassum carpophyllum. Asian Nat. Prod. Res., 4 (2), 95–101 (2002). DOI: 10.1080/10286020290027362.
289. Stirk W. A., Van Staden J. Plant Growth Regulators in Seaweeds. In: Sea Plants, pp. 125–159 (2014). DOI: 10.1016/B978-0-12-408062-1.00005-6.