Microbial or enzymatic biotransformation of nitriles result the conversion of these toxic compounds into industrially important compounds like acids and amides. Nitriles constitute a group of chemicals widely used in drugs, rubbers, and plastic industries. These compounds contain a cyano group in their backbone which is highly correlated with toxicity (Ramteke et al., 2013). Their high rate of manufacture and continuous usage make them an important source of environmental pollution and have been detected in different environmental samples including sediments of water-treatment plants, in marinas and beach areas (Baxter and Cummings, 2006). Clinically, nitrile toxicity has been associated with cancer and different health problems such as bronchial irritation, respiratory disorders, convulsions, coma, and skeletal deformities (Ramteke et al., 2013). Some researchers highlighted the link with psycho-behavioral abnormalities including learning, memory, motor nerve anomalies in rats treated with aliphatic nitriles such as acetonitrile, acrylonitrile and crotononitrile (Boadas-Vaello et al., 2007; Ramteke et al., 2013).

The removal of nitrile compounds from the environment is possible using microbial methods, due to their associated low cost and user-friendly approach (Fang et al., 2015). Microorganisms degrade nitrile compounds through the hydrolytic route, which comprises two enzymatic systems as indicated in Figure 1. In the first route, nitrile hydratase (NHase, EC 4.2.1.84) catalyzes the formation of amides from nitriles, which are later changed to ammonia and carboxylic acids by amidase (EC 3.5.1.4). Alternatively, nitrilase (EC 3.5.5.1) catalyzes for the direct conversion of nitriles into carboxylic acids and ammonia (Ramteke et al., 2013).

The production of acids and amides from nitriles is possible using chemical catalysts. However, this approach is only achieved under harsh conditions like extreme temperature, acidity or alkalinity (Nigam et al., 2009). Currently, many microorganisms having either of the two nitrile degradation enzymatic systems have been reported. These microorganisms can be categorized into two groups. The first is made up of bacteria like Mesorhizobium sp. F28 (Feng and Lee, 2009), Klebsiella oxytoca (Kao et al., 2006), Rhodococcus erythropolis A4 and Rhodococcus rhodochrous PA-34 (Vesela et al., 2012), which only contain a single enzyme system of NHase/amidase and Streptomyces sp. MTCC 7546 only contains nitrilase (Nigam et al., 2009). The other group contain bacteria such as Nocardia globerula NHB-2 (Bhalla and Kumar, 2005), Amycolatopsis sp. IITR215 (Babu et al., 2010), Bacillus subtilis ZJB-063 (Zheng et al., 2008) and R. rhodochrous BX2 (Fang et al., 2015), which have both NHase/amidase and nitrilase.

Variation among nitrile degrading microorganisms also exists in terms of the end products of nitrile degradation. Bacteria in the single enzyme system like R. rhodochrous PA-34 convert nitrile only into amides while K. oxytoca and Mesorhizobium sp. F28 convert nitriles into corresponding amides and carboxylic acids. Those bacteria which utilize the two enzyme system result in amides and carboxylic acids. Among these group of bacteria, R. rhodochrous BX2, B. subtilis AJB-063 and Paracoccus sp. SKG (Santoshkumar et al., 2011) displayed completed degradation of carboxylic acids with final end product of ammonia. As indicated above nitrile degradation systems vary among different bacterial genera as well as with in the same genus such as Rhodococcus. Unlike others, Streptomyces sp. MTCC 7546 in the immobilized as well as Free State biotransforms acrylonitrile into acrylic acid without the formation of amides. The authors suggested that due to several reasons such as operational stability (allow to reuse the system several times), and ease of production on a large scale, the conversion of acrylonitrile using immobilized cells is better than cells in the free state (Nigam et al., 2009).

Phthalate esters and phenols are the two most common chemicals used in industry for stabilization and modification of the characteristics and performance of polymers (He Z. et al., 2014). Di-n-butyl phthalate (DBP), a type of phthalate ester, is a component of different merchandises including pesticides, wrapping materials, makeups, wrappers, wears, and insulators in electric disposals (Dargnat et al., 2009). Similarly, phenol can be applied for the manufacturing of drugs, rubbers, polycarbonate resins, and nylon (Christen et al., 2011).

Phthalates are major environmental pollutants which come into contact with humans and animals through contaminated water systems (He Z. et al., 2014). The European community listed these compounds among the 33 dangerous substances to be controlled in surface water (Dargnat et al., 2009). As they are a constituent of plastics which are now ubiquitous in diverse environments phthalates are now present almost everywhere (Fang et al., 2010). Phthalate toxicity is associated with endocrine system disruption in different species of fish and mammals. These compounds were also observed to interfere with the reproductive system and in human and animal development (Lottrup et al., 2006; Li et al., 2010). Concurrent observation of phenols and phthalate esters has been reported in the Selangor River basin in Malaysia (Santhi and Mustafa, 2013) and induction of lactate dehydrogenase release from Sertoli cells, which is associated with infertility, coexist compared to individual chemical effects (Li et al., 2010).

Different approaches have been documented for removal of DBP from natural environments. These are hydrolysis, photo degradation and biodegradation (Lau et al., 2005; Jonsson et al., 2006; Chen et al., 2009). Two of the former approaches were not effective due to the structural nature of DBP and microbial mediated metabolic transformation of DBP is the current choice. Microbial mediated degradation of DBP involves initial conversion of DBP into phthalic acid and which is further transformed with the help of two dioxygenase pathways into 4,5-dihydroxyphalate and 4,5-dihydroxyphalate in gram negative and gram positive bacteria, respectively. Finally, these two compounds are transformed into a common intermediate protocatechuate under aerobic conditions (Wu et al., 2011). For the degradation of phenols the first step is hydroxylation of phenol to catechol followed by ring cleavage of catechol into 2-hydroxymuconic semialdehyde for the meta-pathway aided by catechol-2, 3-dioxygenase and into cis, cis-muconate with the help of catechol-1, 2-dioxygenase for ortho pathway. Finally 2-hydroxymuconic semialdehyde oxidized 4-oxalocrotonate or hydrolyzed it to 2-oxopent-4-enoate and cis, cis-muconate into muconolactone (Banerjee and Ghoshal, 2010).

Several bacteria strains have the ability to degrade DBP, such as Rhodococcus sp. (Yu et al., 2009; Jin et al., 2010), Gordonia sp. (Wu et al., 2011), Agrobacterium sp. (Wu et al., 2011) and Enterobacter sp. (Fang et al., 2010). Members of the Rhodococcus genus, have been demonstrated for degradation of Phenol in addition to DBP (Saa et al., 2010), individually as well as via simultaneously mineralization of DBP and phenol (Lu et al., 2009; He et al., 2013). Individual or synchronous biodegradation of DBP and phenol by Rhodococcus ruber strain DP-2 was also reported in similar study (He Y.C. et al., 2014).

Chlorophenols (CPs) are aromatic compounds which contain a minimum of one chlorine and a hydroxyl group on the benzene rings. Five types of CPs, as indicated in Figure 2 below, based on chemical structures include monochlorophenols, polychlorophenols, chloronitrophenols, chloroaminophenols, and chloromethylphenols (Arora and Bae, 2014). They are largely used as fungicidal, germicidal and wood preservatives agents. They are also important for synthesis of dyes and drugs (Arora and Bae, 2014). CPs and their derivatives rank among the top environmental pollutants where industrial wastes, pesticides, herbicides, and complex chlorinated hydrocarbons are major sources of contamination (Olaniran and Igbinosa, 2011). Direct skin contact and eating or drinking of contaminated substances are major sources of people exposure (Arora and Bae, 2014). Cellular exposure to CPs are associated with cytotoxic, mutagenic and carcinogenic properties, with several types of polychlorophenols labeled as potential human carcinogens by the World Health Organization and the International Agency for Research on Cancers (Igbinosa et al., 2013).

Different possible mechanisms are reported for bacterial degradation of CPs and their derivatives. In the first mechanism, hydroxylation of chlorophenolic rings at ortho-positions with the help of monooxygenases results in formation of chlorocatechols which are further degraded (Hollender et al., 1997; Solyanikova and Golovleva, 2011) or hydroxylated prior to ring cleavage (Nordin et al., 2005). In the second mechanism, with the same enzyme, hydroxylation of chlorophenolic rings at meta position results in chlorocatechols which degrade via hydroxylation (Nordin et al., 2005) or dehalogenation (Xun et al., 1992) prior to ring cleavage. The third mechanism applicable for degradation of chloronitrophenols where the degradation may be initiated by hydroxylation (Arora and Jain, 2012), reductive dehalogenation (Pandey et al., 2011), or reduction of the nitro group (Arora and Jain, 2012). Finally, in case of chloroaminophenols degradation, the pathway may start with removal of ammonium ions by the enzyme deaminase followed by the ring cleavage (Arora and Bae, 2014) or the dehalogenation (Arora and Bae, 2014). The detail mechanisms of different routes of biodegradation of chlrophenols and its derivatives with various bacteria, such as Pseudomonas knackmussii B-13, Rhodococcus opacus 1G, Arthrobacter chlorophenolicus A6, Streptomyces rochei 303, Pseudomonas sp. NCIB9340, Bacillus insolitus, Nocardioides sp. K44, Mycobacterium chlorophenolicum PCP1 and Mycobacterium fortuitum CG-2 are well documented in a recent review by Arora and Bae (2014); and the following Figure 3 presents degradation of 4-chlorophenol as an example of the process.

Hou and his colleagues reported for the first time magnetically immobilized R. rhodochrous cells for biodegradation of CPs. Their study demonstrated that R. rhodochrous DSM6263 depends on constitutively expressed enzymes for hydroxylation of CPs resulting in chlorocatechol formation and complete degradation. Their observation was consistent with another study where Rhodococcus sp. AN-22 (an aniline-assimilating bacterium) produced cis, cis-muconic acid from catechol (Matsumura et al., 2006) and highlighted how these compounds could also be metabolized with immobilized cells of R. rhodochrous DSM6263 (Hou et al., 2016). Researchers advocated the use of immobilized cells, over free cells, to degrade toxic chemicals due to many reasons such as long-term stability of the catalyst and the immobilization also protects cells from harmful effects of toxic pollutants (Jianlong et al., 2002). Immobilization of cells can be done using a number of techniques including surface adsorption, natural or artificial flocculation, covalent or electrostatic binding to carriers, and encapsulation in a polymer-gel (Hou et al., 2016).

Atrazine (2-chloro-4-(ethylamino)-6-(isopropylamino)-1, 3, 5-triazine) was first introduced in 1950s as an emergent herbicide and they are among widely used pesticides in different countries such as United States, Canada, Africa, and Asia Pacific region (Huang et al., 2003; Jablonowski et al., 2011). Usage of atrazine banned in European countries in 2004 as atrazine concentrations in water surpassed or were estimated to surpass allowable limits (Jablonowski et al., 2011). Due to factors like its widespread utilization as a herbicide and its persistence in the environment, it is common to observe traces of atrazine both in surface and ground water bodies (Gilliom et al., 2006). Traces of atrazine were detected in widely dispersed areas which are far from urban and agricultural areas such as in rainwater in different places (Brun et al., 2008) in fog, arctic ice and seawater.

Contact with atrazine is associated with a serious threat to human and ecosystem health. One of the most notable effects of exposure is endocrine (Solomon et al., 2008; Rohr and McCoy, 2010). Many studies link atrazine with harmful effects on the health of animals and humans, such as sexual abnormalities (demasculinization) in frogs, low testosterone production in rats and higher levels of prostate cancer in workers at an atrazine manufacturing factory (Hecker et al., 2005; Liu and Parales, 2009), and it is also categorized as a group 3 carcinogen according to the International Agency for research on cancer¹. The above observations indicated that there is cause for concern regarding atrazine residues in soil, groundwater, and surface waters.

Due to its persistence in the environment and being highly toxic, it is very important to develop approaches to degrade and remove atrazine deposits from the environment. Microbial-degradation is one of the methods for elimination of atrazine from soil (Tappe et al., 2002). Different species of microorganisms associated with degradation of atrazine with various degree of biodegradation where some undergo complete mineralization while others produce various intermediates including hydroxyatrazine, deethylatrazine, deisopropylatrazine, n-isopropylammilide, n-ethylammilide, and cyanuric acid (Mandelbaum et al., 1995; Ralebits et al., 2002; Ghosh and Philip, 2004; Getenga et al., 2009). Specifically, atrazine degradation was documented with the help of Rhodococcus sp. BCH2 (Kolekar et al., 2014), Arthrobacter sp. (Getenga et al., 2009), Nocardioides sp. (Topp et al., 2000). Pseudomonas sp. strain ADP (Rousseaux et al., 2001; Liu and Parales, 2009) was the first bacterium reported that could completely mineralize atrazine; with most of the degradation studies based on study this strain. As indicated in Figure 4 (below) atrazine degradation is achieved because of the presence of the genes, atzA, atzB and atzC, which code for enzymes such as chlorohydrolase (AtzA), hydroxyatrazine ethylaminohydrolase (AtzB) and N-isopropylammelide isopropylaminohydrolase (AtzC), respectively which convert atrazine sequentially to cyanuric acid (Neumann et al., 2004). Some strains of Pseudomonas can further degrade cyanuric acid into CO₂ and NH₃.

1, 4-Dioxane is a cyclic ether with many applications including components of deodorants, detergents and various types of paints. The process for manufacturing of polyesters also results 1,4-dioxane production. Various factors including illegal dumping of industrial wastes contribute to 1, 4-dioxane associated water contamination. High level of 1, 4-Dioxane can cause liver and nasal cancers in rats module (Dourson et al., 2014) and are listed as group 2B human carcinogen (Inoue et al., 2016). These compounds are soluble in water with low volatility and have a lower chance of absorbance in solids (Stepien et al., 2014). Therefore, once 1, 4-dioxane appears in the environment it can persist for many days and a high degree of 1, 4-dioxane pollution was observed in surface water, groundwater, and landfill leachatewa (Stepien et al., 2014).

Their removal from water bodies is an important public concern, especially as the routine physical and chemical methods for water decontamination are not effective for removal of 1, 4-dioxane (Adams et al., 1994). Advanced procedures such as the combination of ozone and hydrogen peroxide treatments are expensive to use (Adams et al., 1994; Kishimoto et al., 2008). Thus, cost effective as well as reliable methods for cleaning of 1, 4-dioxane from water lead to nocardioform actinobacteria such as Pseudonocardia (Matsui et al., 2016) and Rhodococcus (Deeb and Alvarez-Cohen, 1999) which account for the major portion of capable microorganisms. Inoue et al. (2016) tested various species of Pseudonocardia and Rhodococcus for their ability to degrade 1, 4-dioxane. Their findings indicated P. dioxanivorans JCM 13855T (also known as P. dioxanivorans CB1190) (Parales et al., 1994) was the only Pseudonocardia sp. tested that used 1,4-dioxane as a carbon source and degraded it. In contrast, they observed the inability of R. ruber JCM 3205T to degrade 1, 4-dioxane. However, there are reports on the ability of Rhodococcus spp. such as R. ruber 219 (Bernhardt and Diekmann, 1991) and R. ruber T1 and T5 (Oyama et al., 2013) to biodegrade 1,4-dioxane. This highlighted species variation among Rhodococcus in relation to effective degradation and utilization of 1, 4-dioxane.

Due to the continuing concern about climate change and depletion of fossil energy, expansion of biological processes using renewable biological resources to produce different chemical feed stocks and energy has become an attractive approach for the chemical industry. Ethylene glycol is a feedstock which serves as a starting material for the manufacture of several items including polymers, anti-freeze agents, and coolants (Zhang and Yu, 2013). Routinely, ethylene glycol is produced through a costly chemical process using ethylene derived from the petrochemical industry as a starting material. Therefore, there is a preference for biological synthesis of ethylene glycol over chemical methods as the former has a low impact on the environment and the reaction is selective (Mattam et al., 2013).

Fermentation of carbohydrates is an economical process for production of ethylene glycol from biofeed stock. Corynebacterium glutamicum, an actinobacterium, has been designed for manufacture of ethylene glycol directly from glucose via extension of the serine synthesis pathway. Serine is produced by most microorganisms from 3-phosphoglycerate (a glycolysis intermediate) using three enzymatic steps: 3-phosphoglycerate converted into P-hydroxypyruvate with the help of 3-phosphoglycerate dehydrogenase (PGDH; serA), followed by conversion of P-hydroxypyruvate to P-serine by phosphoserine aminotransferase (PSAT; serC) and P-serine to serine with the help of phosphoserine phosphatase (PSP; serB) (Peters-Wendisch et al., 2005). Chen et al. (2016) proposed two ways for ethylene glycol synthesis via the extension of the serine synthesis pathway where serine is the starting material for both systems with the end product glycoaldehyde (Chen et al., 2016). The first route involves deamination of serine with aminotransferase or amino acid dehydrogenase resulting in hydroxypyruvate. Finally, glycoaldehyde is produce from hydroxypyruvate with the help of α-ketoacid decarboxylase. Similarly, the other route also has two steps where ethanolamide produced from serine with decarboxylation and then converted to glycoaldehyde following oxidation by monoamine oxidase. Finally, glycoaldehyde can be reduce to ethylene glycol by alcohol dehydrogenase such as yqhD as indicated in Figure 5.

In addition to glucose, C. glutamicum uses several substrates such as sugars present in molasses (sucrose and fructose), pentose sugars present in lignocellulosic (Zahoor et al., 2012), n-acetyl-D-glucosamine and n-acetyl-D-muramic acid (Sgobba et al., 2018) for manufacturing of various amino acids (Bommareddy et al., 2014). Production of other compounds such as isobutanol, cadaverine, and succinate are possible with C. glutamicum (Blombach et al., 2011; Buschke et al., 2011; Litsanov et al., 2012).

Rhodococcus sp. are also associated with production of Ethylene glycol. Rhodococcus sp. CGMCC 4911 converted 1, 3-propanediol cyclic sulfate and its derivatives into corresponding diols. The growing cells of Rhodococcus successfully hydrolyzed ethylene sulfate, glycol sulfide, 1, 3-propanediol cyclic sulfate, and 1, 2-propanediol cyclic sulfate with different conversion rates (He et al., 2015).

Cyclic sulfates and its derivatives are very important compounds that serve as starting materials for the synthesis of various useful intermediates (Steinmann et al., 2001). Sulfatases hydrolyse organic sulfate esters into primary or secondary alkyl alcohols (Gadler and Faber, 2007). Microbial transformation of cyclic sulfates into diols can be achieved under mild reaction conditions. He et al. (2013) reported for the first time that Rhodococcus sp. CCZU10-1 can convert 1, 3-propanediol cyclic sulfate and its derivatives into diols, where factors like pH, temperature, and cells dose affect rate of biotransformation.

The absolute configuration of chiral centers in molecules determine the biological activities of the compounds as these molecules bind with receptors made of enantiomerically pure protein (Kato et al., 2003). Most of the available chiral carbon containing drugs are racemic which contain equal concentration of S (+) and R (+) enantiomers. Since only one of the two is active and the other associated with toxicity, the current drug development gives priority to compounds only with a single enantiomer.

Ibuprofen [(R,S)-2-(4′-isobutylphenyl)propionic acid] are a group of effective, orally active, nonsteroidal, anti-inflammatory agents which include drugs such as naproxen, fenoprofen, and flurbiprofen (Sen and Anliker, 1996). Even though the S-(+)- ibuprofen form is more than 100 times more effective than the R-(-)-ibuprofen, the racemic form of ibuprofen is widely used (Kumaresan, 2010). The (R)-ibuprofen form can be converted to its enantiomer in the livers and kidneys of pigs and rats, though this process is not free of toxicity (Liu et al., 2009). Figure 6 below highlighted the application of whole cells N. corallina to biocatalyse the enantioselective hydrolysis of racemic ibuprofen nitrile [2-(4-isobutylphenyl) propanenitrile, 1] to optically active ibuprofen amide [2-(4-isobutylphenyl) propanamide, 2], a prodrug using nitrile hydratase (NHase) and hydrolyse this amide into optically active ibuprofen (3) using amidase.

The results of the above observation indicated N. corallina B-276 displayed nitrile hydratase and amidase activities having low stereo-selectivity (Figure 6). This finding also revealed that N. corallina catalyzed deracemisation of racemic ibuprofen to (R)-ibuprofen enantiomer with an efficiency of >99%. This is the first observation where N. corallina B-276 catalyzed deracemisation processes. Currently, the (R)-enantiomer form of non-steroidal anti-inflammatory agents are the center of studies and research focus on resolving rac-ibuprofen (Pignatello et al., 2008); and due to this, recent efforts have been directed at resolving rac-ibuprofen (Trung et al., 2006).

A high percentage of optical active S-(+)-ibuprofen was reported from hydrolysis of ibuprofen amide and four related 2-phenylpropionamides using whole cell Rhodococcus AJ270 (Snell and Colby, 1999). This high purity was achieved through partial hydrolysis as complete hydrolysis of (R, S)-(+)-2-(4′-isobutylphenyl) propionamide (ibuprofen amide) results in a racemic mixture of ibuprofen enantiomers. The findings of this study suggested that prolonged hydrolysis of ibuprofen amide resulted in both ibuprofen amide and an optical purity of 90-94% of s-(+)-ibuprofen if the reaction is stopped before completion.

Due to its huge production and wide usage, ibuprofen is one of the most commonly detected compounds in wastewater (Buser et al., 1999). In addition to production of active ibuprofen, different species of actinobacteria including Patulibacter sp. strain I11 (Almeida et al., 2013) and Nocardia sp. NRRL 5646 (Chen and Rosazza, 1994; Cy et al., 2018) are involved in its biodegradation to the level where there is no more risk the community.

Carvedilol is a non-selective, β-adrenergic receptor antagonist and α1-adrenoceptor blocker, and it exists in two enantiomeric forms (Gagyi et al., 2008). The overall cardio-protective action of Carvedilol is due to its (S)-(-)-enantiomer, which is less hepatotoxic than the racemic mixture, and (R)-(+)- enantiomer (Hao and Kim, 2010). Ettireddy et al. (2017) tested Streptomyces halstedii and other bacteria for their ability to biotransform racemic carvedilol to its (S)-(-)-enantiomer. The result indicated some bacteria including S. halstedii exhibited incubation time dependence enantioselective conversion of carvedilol where the conversion rate increased up to 10 days of incubation and then after the rate is reduced and finally become zero.

Chiral amines have been widely used for manufacturing of several therapeutics such as codeine (pain relief), zoloft / sertraline (anti-depression), lariam (anti-malaria) and ethambuto (anti tuberculosis) and agrochemical intermediates including insecticides (imiprothrin, nornicotine), herbicides (imazapyr, imazapic) and fungicides (cyprofuram, fenbuconazole) (Ulrich et al., 2012). Several syntheses of optically active amines have been studied for many years. Asymmetric synthesis of chiral cyclic amine from cyclic imine achieved using whole-cell Streptomyces sp. GF3587 and 3546. As showed in Figure 7, these strains produced novel imine reductase enzymes which facilitate enantioselective redaction of cyclic 2-methyl-1-pyrroline into R-2-methylpyrrolidine and S-2-methylpyrrolidine in the presence of glucose.

In addition to enantioselective production of active drugs, regioselective addition or substitution of functional groups can also produce active drugs. Actinobacteria play a role in regioselective hydroxylation which mediates the conversion into more active forms. The hydroxylated form of isoflavones such as daidzein (4′, 7-dihydroxyisoflavone) and genistein (4′,5,7-trihydroxyisoflavone) are associated with lowering blood cholesterol and preventing cardiovascular diseases and cancer (Foti et al., 2005). As presented in Figure 8 below, some cytochrome P450 monooxygenases of Streptomyces avermitilis MA-4680 catalyze 3′-specific hydroxylation of daidzein and genistein to 3′,4′,7-trihydroxyisoflavone and 3′,4′,5,7-tetrahydroxyisoflavone, respectively (Roh et al., 2009).

Microbial biotransformation of natural steroids into pharmaceutically active intermediates has been practiced for many years (Fernandes et al., 2003). These active intermediates play a role in the manufacturing of all type I aromatase inhibitors (Lombardi, 2002) and several high-value steroidal drugs (Wadhwa and Smith, 2000).The bioconversion reaction proceeds with removal of the C-17 side chain of the steroid without any modification of the steroid nucleus (Szentirmai, 1990; Murohisa and Iida, 1993). As indicated in Figure 9, Mycobacterium sp. NRRL B-3683 and Mycobacterium sp. NRRL B-3805 facilitate a single step C-17 side chain cleavage of sitosterol, cholesterol, stigmasterol and ergosterol to produce C-19 steroids such as 1-androstene-3,17-dione and 1,4-androstadiene-3,17-dione (Sripalakit et al., 2006). The initial step of the side-chain oxidation of sterols is hydroxylation at C-17. The reaction is catalyzed by cytochrome P450 monooxygenase, then several dehydrogenase including 3-β-hydroxysterol dehydrogenase remove hydrogen to introduced a double bound at various points (Donova and Egorova, 2012).

Amino acids alone or in combination with other molecules can be used as drugs, food supplements, agriculture chemicals, and polymers. Ever year the annual production of amino acids has increased and is currently estimated to be about 3.7 million metric tons (Ikeda and Takeno, 2013). The global market for amino acids is estimated to be about US$6.6 billion and expected to increase by 8-10% each year (Leuchtenberger et al., 2005; Ikeda and Takeno, 2013). Amino acids including L-lysine, DL-methionine, L-threonine, and L-tryptophan account for the largest share in terms of production followed by L-glutamate, L-aspartate, and L-phenylalanine (Ikeda and Takeno, 2013).

Microbial fermentation is the primary source of most of the available amino acids in the market. The L-glutamate-producing bacterium, C. glutamicum, has been the cornerstone for the introduction of fermentation in industrial manufacturing of amino acids (Udaka, 1960). This bacterium is still the most widely utilized strain for manufacturing of important amino acids, mainly L-glutamate and L-lysine (Leuchtenberger et al., 2005). Manipulation of metabolic pathways of this bacterium allowed for the an improved product range including L-phenylalanine, L-aspartate, L-tryptophan, L-arginine, L-valine, nucleic acids such as purines, vitamins such as riboflavin and pantothenic acid (Burkovski, 2008; Gopinath et al., 2012) and significant amounts of organic acids such as acetic, lactic and succinic acid (Inui et al., 2004). Figure 10 highlights the biotechnological role of C. glutamicum. As indicated, C. glutamicum utilizes different starting materials for manufacturing of amino acids and other chemical commodities.

Apart from the above mentioned biologically active compounds, different species of actinobacteria are involved in the synthesis of active compounds. Table 2 below summarizes some of these compounds.