1 Introduction

Polymers are the most commonly exploited materials that are used to make human life more comfortable, and it is expected that the demand for polymers will exceed 400 million tons in 2020 (Zare 2015). Most polymers are fossil based, and their increasing demand enforces the overexploitation of these resources. The massive exploitation of these resources may lead to severe problems because their formation takes millions of years (Jiang and Loos 2016). The production of polymers also generates hazardous gases that are responsible for global warming. Biobased polymers (biopolymers) are a promising alternative because they are produced from renewable raw material (Ates 2015). Considering the abundance of raw material present in nature, researchers are focusing on the production of polymers through the biological routes. Microbes can synthesize and accumulate specific types of biopolymers inside their cells through enzymatically catalyzed reactions starting from various monomers produced during metabolic processes (Rehm 2010). The microbial production of biopolymers is limited as microbes require optimal conditions for growth and biopolymer accumulation. Additionally, it is difficult to engineer microbes for the synthesis of new biopolymers and to recover these from the fermentation broth. To increase the role of microbes in biopolymer production, researchers are attempting to engineer microbes for the production of monomer units that can be combined to produce different types of industrially important polymers (Bozell and Petersen 2010).

Polyesters and polyamides have immense potential due to their structural diversity, mechanical strength, and biodegradation and their accessibility due to the use of natural building blocks. Thus, many researchers are working on biobased polymer synthesis, typically using partial or complete biological methods. Researchers have succeeded in producing various monomers at the industrial scale through microbial fermentation, such as itaconic acid, succinic acid, 2,3-butanediol and 1,3-butanediol (Blazeck et al. 2015; Chen et al. 2014; Cho et al. 2015). These monomer units can be polymerized into various polymers through chemical- or enzyme-mediated condensation reactions. Various polymers, such as poly(hydroxybutyrate) (PHB), poly(butylene succinate) (PBS), poly(ethylene terephthalate) (PET), poly(butylene terephthalate) (PBT), nylon-5,4, nylon-4,6, that can be produced using biobased monomers (Houck et al. 2001; Kind et al. 2014). These polymers have applications in the textile, pharmaceutical and medical sectors (Hemalatha et al. 2016; Vardon et al. 2016). However, researchers have failed to develop an efficient and productive bioprocess for biopolymer production (Rehm 2010). The engineering of microbes for improved yield, higher product tolerance, and utilization of low-cost feed stock as a carbon source are major challenges in cost-effective bioprocess development. This review paper is focused on the advances in the production of polyester and polyamide building blocks, i.e., hydroxy acids, diacids, diols and diamines, using wild and engineered microbes in recent years (Fig. 1).

Fig. 1
figure 1

Schematic representation of the possible metabolic pathways involved in polyester and polyamide building-block (2-hydroxyisobutyric acid, 3-hydroxypropionic acid, itaconic acid, succinic acid, 1,3-propanediol, 1,3-butanediol, 1,4-butanediol, 2,3-butanediol, cadaverine and putrescine) synthesis in microbial systems. Substrates are presented in bold letters, and products are in colored square boxes. The central metabolic pathway is indicated by rounded-end colored square boxes. Multi-enzyme pathways are indicated by dotted lines, and single-enzyme-catalyzed reactions are indicated by simple arrowed lines. The enzymes coded by the genes are: glycerol dehydratase (dhaB), aldehyde dehydrogenase (ald), 1,3-PDO oxidoreductase (dhaT), glycerol dehydrogenase (dhaD), aconitase (acnA), cis-aconitic acid decarboxylase (cadA), ornithine carbamoyltransferase (argI), CoA-dependent succinate semialdehyde dehydrogenase (sucD), 4-hydroxybutyrate dehydrogenase (4hbd), 4-hydroxybutyryl-CoA transferase (cat2), 4-hydroxybutyryl-CoA reductase (ald), butyryl-CoA dehydrogenase (bld), alcohol dehydrogenase (adh), propionyl-CoA dehydrogenase (pacd), 3-hydroxypropionyl-CoA dehydratase (hpcd), propionate-CoA transferase (pct), methylmalonyl-CoA decarboxylase (ygfG), β-ketothiolase (phaA), acetoacetyl-CoA reductase (phaB), CoA-carbonyl mutase (mdpOR), 2-hydroxyisobutyryl-CoA mutase (hcm), α-acetolactate synthase (α-als), α-acetolactate decarboxylase (α-ald) and acetoin reductase (ar)

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2 Biological production of hydroxy acids

2.1 2-Hydroxyisobutyric acid (2-HIBA)

Various companies, such as Santa Cruz Biotechnology (California) and Tokyo industry Co. Ltd., are involved in 2-HIBA production. 2-HIBA has many applications, but researchers have not been able to produce it at large scale using microbial fermentation as it is not the main end product of the metabolic pathway. It can be produced using a partial biotechnology route, such as the hydrolysis of 2-hydroxyisobutyronitrile (acetone cyanohydrin) using nitrilase-mediated reactions. Nitrilase systems have been reported in various microorganisms, including Alcaligenes, Pseudomonas and Penicillium (Bhatia et al. 2014a; Duca et al. 2014; Kaplan et al. 2006). 2-HIBA is produced by acetone cyanohydrin using whole cells or pure enzyme reactions at the desired temperature. There are two pathways reported for 2-HIBA production starting from acetone cyanohydrin: (1) a single-step reaction mediated by nitrilase and (2) a two enzyme system, where the first step is catalyzed by nitrile hydratase and an intermediate amide is produced, which is further converted into the acid by amidase (Fig. 2a) (Bhatia et al. 2013). Alcaligenes sp. MTCC 10674 whole cells induced by isobutyronitrile can produce 25 g/L 2-HIBA by biotransformation of acetone cyanohydrin (Table 1) (Bhatia et al. 2013). The main disadvantage of this process is the instability of acetone cyanohydrin. At pH 7, it degrades into acetone and cyanide. The nitrile hydrolyzing enzyme requires acidic pH for optimum activity and loses its activity in the presence of acetone and cyanide.

Fig. 2
figure 2

Biotechnological route for 2-hydroxyisobutyric acid, mandelic acid and adipic acid: a common reaction for acid production starting from the corresponding nitriles and b adipic acid production from glucose, benzoate, long-chain dicarboxylic acid and adiponitrile. The dotted arrows represent multiple enzyme reactions

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Table 1 Summary of biopolymer building block produced using microbial systems
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Another method for 2-HIBA production is the oxidation of tert-butanol (Lopes Ferreira et al. 2006). Mycobacterium austroafricanum IFP 2012 has been reported for monooxygenase activity on tert-butanol to produce diol. Diol is further oxidized by two dehydrogenases, MpdB and MpdC, to 2-HIBA (Lopes Ferreira et al. 2006). Aquincola tertiaricarbonis L108 possesses a mutase enzyme that can act on 3-hydroxybutyryl-CoA to produce 2-HIBA (Rohwerder et al. 2006). Muller et al. engineered the Citrobacter necator H16 strain deficient in PHA synthase (phaC) for autotrophic production of 2-HIBA by overexpressing 2-hydroxyisobutyryl-CoA mutase (2-HIB-CoA) from A. tertiaricarbonis L108 that can produce 3.2 g/L 2-HIBA from hydrogen and carbon dioxide (Przybylski et al. 2015).

2.2 3-Hydroxypropionic acid (3-HP)

3-HP is one of the top twelve chemical compounds that can be produced from biomass and was initially reported in Chloroflexus aurantiacus (Holo 1989). Many microbes have been found to store 3-HP as a storage material, e.g., Alcaligenes latus, Metallosphaera sedula, Pseudomonas oleovorans and Ralstonia eutropha (Berg et al. 2007; Holo 1989; Saito and Doi 1994). 3-HP is an intermediate of autotrophic carbon dioxide fixation pathway, which is self-sufficient in CO2 fixation. C. aurantiacus can grow phototrophically and can secrete 3-HP (Holo 1989). Various pathways are involved in 3-HP production using glucose and glycerol as the carbon sources. Byssochlamys sp. can utilize glucose to produce 4.8 % v/v 3-HP in the presence of 7 % v/v acrylic acid. Lee et al. isolated Rhodococcus erythropolis LG12, which can use acrylic acid as the sole carbon source to produce 17.5 g/L 3-HP (Table 1) (Lee et al. 2009). Many microbes have been reported to naturally utilize glycerol to produce 3-HP, e.g., Klebsiella pneumonia and Citrobacter freundii. Glycerol dehydratase (dhaB) and aldehyde dehydrogenase (ald) are enzymes involved in the conversion of glycerol to 3-HP. dhaB catalyzes the conversion of glycerol to 3-hydroxypropionaldehyde (3-HPA) in the presence of coenzyme B12 (Honda et al. 1980; Seyfried et al. 1996). ald uses the 3-HPA intermediate as a substrate and oxidizes it to 3-HP using NAD+ or NADP+ as cofactors (Jo et al. 2008).

Most of the organisms can produce 3-HP as an intermediate, but no organism can produce 3-HP as a major metabolic end product. Genetic engineering can be applied to produce 3-HP as a major product using E. coli and K. pneumoniae as host strains. Both strains have their own advantages and disadvantages, such as E. coli is easy to engineer and possesses abundant ald genes, but it lacks B12 coenzyme production, which leads to high production cost (Luo et al. 2012). K. pneumoniae has its own dha operon and can synthesize the B12 coenzyme, but it is not easy to transform due to a lack of genetic diversity tools (Huang et al. 2012). 1,3-PDO is a side product produced by oxidoreductase (dhaT and yqhD) using the 3-HPA intermediate as the substrate, which reduces 3-HP production (Kwak et al. 2013). Kwangwook et al. engineered E. coli by disrupting the glycerol metabolizing gene (glycerol kinase (glpk), glycerol dehydrogenase (gldA) and alcohol dehydrogenase (yqhD) and overexpressing genes involved in the 3-HP pathway. The engineered E. coli strain can produce 57.3 g/L 3-HP (Kim et al. 2014). There is another route using engineered E. coli to produce 3-HP through the propionyl-CoA pathway. Recombinant E. coli overexpressing propionyl-CoA dehydrogenase (pacd), 3-hydroxypropionyl-CoA dehydratase (hpcd), and propionate-CoA transferase (pct) produce sixfold 3-HP than the parental strain (Luo et al. 2012). Researchers have proposed additional pathways for 3-HP production using various intermediates, e.g., alanine, lactate and malonyl Co-A, but there is not method to produce 3-HP in good quantity using a microbial system (Borodina et al. 2015; Rathnasingh et al. 2012). Glucose and glycerol are typically used as the carbon sources for 3-HP production; there is a need to find a cheaper carbon source for microbial fermentation.

2.3 Mandelic acid (MA)

Mandelic acid (MA) is an industrially important precursor molecule used for the production of a wide range of pharmaceutically important molecules, e.g., antibiotics, antitumor agents and antiobesity drugs (Bhatia et al. 2014a, b; Wang et al. 2015). Recently, MA has received attention because a polymer produced through the condensation of MA showed antiviral activity against human immunodeficiency virus (HIV) and herpes simplex virus (HSV) (Herold et al. 2002). There are multiple chemical and biotechnological routes available for MA production. MA can be produced using arylacetonitrilase, a nitrilase that can hydrolyze mandelonitrile (Fig. 2a) (Zhang et al. 2010). Bhatia et al. (2014b) isolated Alcaligenes sp. MTCC 10675 from soil samples and produced MA (0.78 g/L/h) from mandelonitrile using the arylacetonitrilase activity of this organism (Table 1). The arylacetonitrilase-mediated synthesis of MA has advantages over other routes because it does not require a cofactor and inexpensive mandelonitrile acts as the starting material (Gong et al. 2012).

Substrate inhibition is the main limitation in nitrilase-mediated reactions. Wang et al. (2015) engineered E. coli by overexpressing nitrilase from Burkholderia cenocepacia J2315, and the recombinant strain showed high substrate tolerance and produced 14.5 g/L/h MA in fed-batch reaction mode. To solve the stability and reusability problem, Kaul et al. (2006) developed a bioprocess by immobilizing whole cells of Alcaligenes faecalis on an alginate matrix and produced 35.5 g/L/h MA. Nitriles are toxic compounds, so researchers are focused on finding an alternate route for MA production using renewable resources. Sun et al. (2011) constructed a phenylalanine pathway in E. coli by overexpressing hydroxymandelate synthase (hmaS) from Amycolatopsis orientalis and produced 0.03 g/L/h MA.

3 Biological production of diacids

3.1 Itaconic acid (IA)

Worldwide production of IA has exceeded 88,000 tons, and its selling price is approximately US$ 2/kg. Pfizer Co. Inc. (New York) started IA production using fermentation, and other countries, such as Japan, France, England and Russia, also became involved in this business. Aspergillus species, e.g., A. itaconicus and A. terreus, have the ability to secrete IA acid at >80 g/L. However, this productivity is lower than the rate at which researchers are already able to produce citrate (up to 200 g/L), which an intermediate for IA production. Kobayashi and Nakamura reported that low IA production was due to the accumulation of IA because it suppressed the growth of A. terreus. To overcome the product inhibition problem, Yahiro et al. (1995) generated the mutant A. terreus IFO 6365 using N-methyl N′-nitro-N-nitrosoguanidine treatment and achieved 82 g/L IA in a shake flask (Table 1).

Glucose is used as a carbon source for the commercial production of IA, which makes the production process costly. Therefore, alternative cheaper raw materials, such as starch, cellulose molasses, and other abundantly available biomass, are needed. IA production using sago starch is almost equivalent to production using glucose, but the purity is slightly lower (Dwiarti et al. 2007). cis-Aconitate decarboxylase (CadA), encoded by cadA, is the key enzyme in IA production from citrate. Scale up of IA production is problematic because a short (20 min) interruption of the oxygen supply decreases the IA yield up to 51.2 % (Lin et al. 2004). To reduce the sensitivity to oxygen, a hemoglobin gene (vgb) from Vitreoscilla was expressed in A. terreus, and 17 % higher IA production was recorded (Lin et al. 2004). IA production by stirred-tank fermentation is a challenging because fungal culture growth is affected by the stress of the agitator, which results into low titer. The air lift reactor (ALR) has been widely studied, and almost double IA production was achieved compared to STR (Yahiro et al. 1997). The fungal strain Yarrowia lipolytica, which can accumulate citrate pathway intermediates, was engineered by heterologously expressing IA synthesis genes from A. terreus and produced 4.6 g/L IA using glucose as the carbon source (Blazeck et al. 2015). Rafi et al. investigated Ustilago maydis using orange pulp as a carbon source in solid-state fermentation and produced 28.9 g/kg IA (RAFI 2014). E. coli BW25113 (DE3), a strain deficient in phosphate acetyltransferase (pta) and lactate dehydrogenase (ldhA), was engineered by overexpressing citrate synthase (gltA) and aconitase (acnA) from Corynebacterium glutamicum and can produce 0.086 g/L IA (Vuoristo et al. 2015). Many research groups are searching for cheap raw materials for IA production with enhanced efficiency and purity.

3.2 Adipic acid (AA)

AA is an important polymer precursor with a global market of $6.3 billion per year. The major companies involved in AA production are Invista, PetroChina, Ascend and Honeywell. The production of AA has reached approximately 2.85 million tons using petrochemical sources (cyclohexanol and cyclohexanone) as the raw material (Polen et al. 2013). These methods lead to excessive production of carbon dioxide, nitrous oxide and nitrogen oxide, which cause environmental problems. Researchers are working on more ecofriendly and renewable routes for AA production using a biological system, but no biobased method has been developed because there is no natural metabolic pathway present in microbes (Kircher 2006; Niu et al. 2002).

AA can be produced using chemo-catalytic conversion of the biobased precursors cis-muconic acid and d-glucaric acid, which are produced by fermentation (Fig. 2b) (Jung et al. 2015; Moon et al. 2009; Weber et al. 2012; Yu et al. 2011). Cis-muconic acid and d-glucaric are intermediates of the benzoate and ascorbate metabolic pathways (Myers et al. 2001; Yu et al. 2011). Frost and coworkers developed an E. coli strain that can produce cis-muconic acid from 3-dehydroshikimic acid (DHS), an intermediate of the aromatic amino acid pathway (Draths and Frost 1994). d-Glucaric acid production was initially reported by Moon and coworkers using microbial fermentation. They constructed an E. coli strain by overexpressing the genes myo-inositol-1-phosphate synthase (ino1) from Saccharomyces cerevisiae and myo-inositol oxygenase (miox) from mice and produced glucaric acid from glucose-6-phosphate (Moon et al. 2009). AA can also be produced by α- or β-oxidation of long-chain n-alkanes, alcohols or fatty acid substrates using engineered yeast (Smit et al. 2005). An engineered strain of Candida tropicalis can produce AA by α- or β-oxidation of fatty acids (Yang et al. 2010). Enzymatically, AA is produced from adiponitrile using nitrilase, but researchers have not been developed a commercial process due to the low activity of nitrilase (Fig. 2b) (Bayer et al. 2011).

3.3 Terephthalic acid (TPA)

TPA is a commodity chemical produced by the aerobic catalytic oxidation of para-xylene (pX). Its global consumption was approximately 47 million tons in 2012, and it is expected to increase to 65 million tons by 2018 (Collias et al. 2014). Purified TPA is used as a raw material for high-performance, multi-purpose plastics, e.g., polybutyl terephthalate (PBT), polyethylene terephthalate (PET), and polytrimethylene terephthalate (PTT) (Tachibana et al. 2015). Biobased methods can be used to produce TPA and its precursors para-xylene (pX), para-toluic acid (pTA), para-tolualdehyde (pTALD), and 4-carboxybenzaldehyde (4-CBA), which can be converted to TPA (Collias et al. 2014). Microbes do not naturally possess a pathway to produce these intermediates, but genetic engineering can be used to construct new pathways to convert carbohydrates into TPA and its precursors. Most microbes can degrade TPA and its precursors (toluene and aromatic compounds), which restricts their use in TPA production. The degradation pathways typically involve monooxygenases, which operate irreversibly to degrade aromatic compounds rather than to synthesize compounds.

A research group from the DuPont Company isolated a group of bacteria from a wastewater treatment bioreactor that could grow on pX, pTA, and TPA (Bramucci et al. 2002). These bacteria degraded pX by oxidizing the methyl group to form pTA and then cleaving the aromatic ring to produce TPA. Biomass can also be used to produce TPA, but microbes cannot utilize biomass directly as a carbon source. The biomass is first converted to free sugars through hydrolysis using acid and base pretreatment followed by fermentation. Depolymerization of lignin leads to different aromatic compounds, including benzene, toluene and xylene (BTX) (Fan et al. 2014). Most of the xylene produced from lignin is meta-xylene which can be isomerized into para-xylene for use in TPA production. During the pretreatment of biomass, xylose is degraded into furfural (Li et al. 2015). A research group from Japan reported a synthetic pathway for TPA production from furfural. In this process, furfural was oxidized and dehydrated to give maleic anhydride, which yielded the Diels–Alder (DA) adduct upon reaction with furan. Upon dehydration, the DA adduct gave phthalic anhydride, which was converted via phthalic acid and dipotassium phthalate to TPA (Tachibana et al. 2015). Many novel pathways have been explored by researchers, but there remains a need for a complete biological route for TPA production (Miller et al. 2014).

3.4 Succinic acid (SA)

SA is an important feedstock for many industrial products. SA has a current global production of approximately 30,000–50,000 tons annually. The market price is US$ 2400–3000 per ton and is expected to grow at a rate of 18.7 % from 2011 to 2016 (Cao et al. 2013; Cheng et al. 2012). Europe and North America are the two largest consumers, followed by China and India, and the increasing demand is prompting researchers to develop more efficient ways to produce SA. SA is traditionally produced by catalytic oxidation of paraffin, but the yield and purity are low, and the production method causes pollution problems (Cao et al. 2013). The SA metabolic pathway occurs in a natural biological system so it can be produced through microbial fermentation (Fig. 1). A number of natural and engineered microbes have been reported to produce succinate. Most natural producers are facultative anaerobic bacteria and fungi, i.e., Actinobacillus succinogenes, A. niger, Anaerobiospirillum succiniciproducens, Mannheimia succiniciproducens and Paecilomyces varioti (Carvalho et al. 2016; Lee et al. 2001, 2002; Salvachúa et al. 2016). A. succinogenes can utilize a broad range of carbon sources, including arabinose, fructose, glucose, galactose, cellobiose, maltose, lactose, sucrose, mannose, mannitol and xylose (Guettler et al. 1999; Van der Werf et al. 1997). A. succinogenes FZ53 is the only well-characterized natural strain and can produce 105.8 g/L SA (Table 1).

In addition to natural producers, other genetically engineered microbes, such as E. coli, Corynebacterium glutamicum and Saccharomyces cerevisiae, also produce SA. C. glutamicum overexpressing pyruvate carboxylase (pyc) can produce up to 146 g/L SA using sodium bicarbonate and glucose (Table 1) (Okino et al. 2008). The production of SA using fermentation is currently at the demonstration level due to the high production costs. Downstream processing is a major factor in determining the production cost of succinate, and greater focus is needed on this research area. The quality of the fermentation process also leads to difficulty in downstream processing because several side products (organic acids) are produced during fermentation. Microbial engineering for improved SA production with minimal side products may result in simpler and less expensive succinate production. Different polyesters of SA, i.e., poly(ethylene succinate) (PES) and poly(trimethylene succinate) (PTS), can be produced by direct polycondensation with titanium tetraisopropoxide as the catalyst (Tsai et al. 2008).

4 Biological production of dialcohols

4.1 1,3-Propanediol (1,3-PDO)

1,3-PDO is one of the oldest chemicals identified during the glycerol fermentation of Clostridium pasteurianum, with a current market demand of 100 million pounds annually (Kraus 2008). DuPont and Shell are the major producers of 1,3-PDO, using acrolein and ethylene oxide as the raw material, respectively (Saxena et al. 2009). These chemical methods require expensive catalysts and release toxic intermediates, which makes a new fermentation method desirable. Glycerol is a major byproduct of the biodiesel industry, and more than 66 × 105 tons are produced by Europe alone (Papanikolaou and Aggelis 2009). 1,3-PDO production through fermentation is restricted to bacteria (Biebl et al. 1999). Many microbes ferment glucose into glycerol or glycerol then into 1,3-PDO (Fig. 1), but no organism can ferment sugar directly into 1,3-PDO (Cameron et al. 1998).

Metabolic engineering can be applied to combine the sugar fermentation pathway with 1,3-PDO production (Tong and Cameron 1992). Citrobacter, Clostridia, Klebsiella, and Lactobacillus can produce 1,3-PDO naturally from glycerol (da Silva et al. 2015; Drozdzynska et al. 2014; Ricci et al. 2015; Szymanowska-Powałowska 2014). Glycerol dehydratase (gdhB) and 1,3-propanediol oxidoreductase (dhaT) are the main enzymes involved in 1,3-PDO production from glycerol (Fig. 1). gdhB converts glycerol to 1,3-hydroxypropanaldehyde (1,3-HPA), which dhaT further oxidizes to 1,3-PDO (Jiang et al. 2016). Glycerol phosphatase catalyzes the conversion of sugar into glycerol, and has been reported in many microbes, such as Bacillus licheniformis, Pichia farinose and Saccharomyces cerevisiae. Multiple fermentation strategies have been used for 1,3-PDO production, such as batch, fed batch, continuous, aerobic fed batch, micro-aerobic fed batch and immobilization using various microbes (Cheng et al. 2007; Hao et al. 2008; Papanikolaou et al. 2008; Yang et al. 2007; Zhao et al. 2006). Cheng et al. (2007) scaled up 1,3-PDO production to 5000 L using K. pneumoniae and produced 58.8 g/L 1,3-PDO (Table 1). To further increase 1,3-PDO productivity, Zhao et al. (2006) used immobilized cells of K. pneumonia in batch culture and produced 63.1 g/L 1,3-PDO from glycerol. The substrate and products inhibit the productivity of 1,3-PDO. To overcome this inhibition, Petitdemange et al. (1995) isolated a new strain, Clostridium butyricum E5, that was resistant to high levels of glycerol and 1,3-PDO and produced 58 g/L 1,3-PDO using 109 g/L glycerol (Table 1).

Despite the development of various fermentation strategies 1,3-PDO productivity is still low, so metabolic engineering is a good approach to further increase the productivity. Klebsiella oxytoca M5al is an excellent producer of 1,3-PDO, but the coproduction of lactic acid results in decreased productivity. Yang et al. engineered K. oxytoca M5al by deleting lactate dehydrogenase (ldhA), and the mutant strain produced 83.56 g/L 1,3-PDO in fed-batch fermentation (Yang et al. 2007). C. freundii and K. pneumoniae are the most widely studied bacterial strains for 1,3-PDO production, but their pathogenic nature restricts their use at the industrial level. To solve this problem, Przystałowska et al. engineered E. coli by heterologously overexpressing glycerol dehydratase (dhaBCE) from C. freundii and 1,3-PDO oxidoreductase from K. pneumoniae. The engineered E. coli strain can produce 10.6 g/L 1,3-PDO in batch fermentation (Przystałowska et al. 2015). Tang et al. performed two-stage fermentation using recombinant E. coli. During the first phase, a high cell density was achieved. In the second stage, fresh glycerol was fed into the reactor and 104.4 g/L 1,3-PDO was produced (Table 1). To further decrease the production cost, new routes are needed to produce 1,3-PDO using an abundant carbon source.

4.2 2,3-Butanediol (2,3-BDO)

2,3-BDO is a commodity chemical used for the production of biopolymers and number of industrially important compounds, such as methyl ketone and 1,3-butadiene (Jing et al. 2016; Tran and Chambers 1987). 2,3-BDO has a global market of approximately 32 million tons. After World War II, 2,3-BDO gained attention because it can be used for the production of synthetic rubber. 2,3-BDO is produced by microbial fermentation through α-acetolactate, acetoin and diacetyl, with the intermediate steps catalyzed by α-acetolactate synthase, α-acetolactate decarboxylase, and acetoin reductase (Fig. 1) (Mallonee and Speckman 1988). Acetate acts as an inducer for these enzymes, and their expression helps to reduce intracellular acidification by diverting acid production to neutral compounds (Blomqvist et al. 1993). Many bacterial species produce 2,3-BDO through a pyruvate intermediate, e.g., Aeromonas, Bacillus, Klebsiella, Lactobacillus and Synechocystis (de Oliveira and Nicholson 2016; Gaspar et al. 2011; Jung et al. 2014; Savakis et al. 2013; Willetts 1985). Klebsiella sp. is the best reported microbe because it can utilize a broad range of carbon sources, such as arabinose, glucose, galactose, mannose, xylose, cellobiose, lactose and glycerol. Glycerol is a good substrate for K. pneumoniae G31 and can produce 49.2 g/L 2,3-BDO in a fed-batch process (Table 1) (Petrov and Petrova 2009). 2,3-BDO can also be produced using other carbon sources, such as lignocellulose and noncellulosic biomass (Jerusalem artichoke tubers). Sun et al. used Jerusalem artichoke tuber hydrolysate as a carbon source for K. pneumoniae fermentation and produced 91.63 g/L 2,3-BDO (Sun et al. 2009).

The concentration of CO2 -in the atmosphere has increased by 25 % during the last 150 years; researchers are now suggesting that it could be assimilated by photosynthetic microorganisms to produce chemicals. Oliver et al. engineered a 2,3-BDO pathway in Synechococcus elongates PCC7942 and produced 2.38 g/L 2,3-BDO using CO2 (Oliver et al. 2013). Abubackar et al. developed a method for carbon dioxide utilization using Clostridium autoethanogenum that could produce 2,3-butanediol and other alcohols (Abubackar et al. 2015). K. pneumoniae is not suitable for the industrial production of 2,3-BDO due to its pathogenic nature. C. glutamicum, which is generally regarded as safe, was engineered by Rados et al. with the 2,3-BDO synthesis pathway genes (Rados et al. 2015). The engineered C. glutamicum can grow aerobically on acetate and can convert glucose into 2,3-BDO (6.3 g/L) under non-growing oxygen-limiting conditions. Bacillus amyloliquefaciens B10-127 showed excellent glycerol metabolizing potential, but its 2,3-BDO productivity was low. Yang et al. engineered B. amyloliquefaciens B10-127 with a cofactor (NADH/NAD+) regeneration system and overexpression of glycerol dehydrogenase and acetoin reductase (ar) to produce 102.3 g/L 2,3-BDO (Fig. 1) (Yang et al. 2015a). The highest 2,3-BDO productivity (131.5 g/L) was achieved in Klebsiella oxytoca M3, a strain deficient in pduC (encoding glycerol dehydratase large subunit) and ldhA (encoding lactate dehydrogenase), which produce 1,3-PDO and lactic acid side products (Cho et al. 2015).

4.3 1,4-Butanediol (1,4-BDO)

1,4-BDO has an annual global production of approximately 2 million tons and is mainly used for the production of automotive plastics and electronics (Burgard et al. 2016). Fossil feedstocks (coal and oil) are typically used for diol production, and these methods lead to greenhouse gas emission. There is no natural pathway for 1,4-BDO synthesis in microbes. The biological production of 1,4-BDO requires four enzymatic steps, starting from a succinyl-CoA intermediate catalyzed by CoA-dependent succinate semialdehyde dehydrogenase (sucD), 4-hydroxybutyrate dehydrogenase (4hbd), CoA-acyl transferase (cat2), 4-hydroxybutyryl-CoA reductase (ald), butyryl-CoA dehydrogenase (bld) and alcohol dehydrogenase (adh) (Fig. 1). The production of 1,4-BDO from the succinyl-CoA intermediate requires several reduction steps and a net energy input, so there is need for a strain that can balance both redox and ATP (Barton et al. 2015). Barton designed a new pathway using SimPheny™ Biopathway Predictor software to screen several enzymes and engineered E. coli with the best 1,4-BDO synthesis genes. Several genes that produce side products, such as ethanol, formate and lactate, were knocked out to conserve the redox activity for use in 1,4-BDO production. The engineered strain can produce 18 g/L 1,4-BDO in fed-batch fermentation (Table 1) (Barton et al. 2015).

Biomass is an abundantly available carbon source that is composed of glucose, xylose, arabinose and galacturonate as the main component. A de novo pathway for 1,4-BDO production was constructed in E. coli by combining synthetic biology and metabolic engineering. 1,4-BDO production and carbon utilization were separated into two parts. 1,4-BDO production was separated into 1,4-BDO synthesis and genetic control, where the former was performed by the de novo enzymatic pathway and the latter using a synthetic circuit that acts as a genetic controller. Carbon utilization was broken into two routes, where xylose is used for 1,4-BDO production and the other carbon sources are utilized for cellular growth. The final engineered strain produced 0.44 g/L 1,4-BDO (Liu and Lu 2015). Tai et al. engineered E. coli for the nonphosphorylative assimilation of sugars (xylose, arabinose and galacturonate) to produce TCA cycle derivatives. The engineered strain could produce 16.5 g/L 1,4-BDO in fed-batch fermentation (Tai et al. 2016). 1,4-BDO production is challenging because it requires a balance between the pathway intermediates, redox cofactors and ATP.

4.4 1,3-Butanediol (1,3-BDO)

1,3-BDO is an important industrial chemical with applications in the production of polymers, antibiotics, pheromones and insecticides (Matsuyama et al. 2001). 1,3-BDO is generally produced through a multistep chemical reaction starting from acetaldehyde. Despite its immense demand, researchers have not developed a bioprocess for the production of 1,3-BDO using a renewable raw material, and no natural pathway exists. Therefore, its production strictly depends on the genetic engineering of a new strain. Kataoka et al. constructed an E. coli strain by heterologously overexpressing beta-ketothiolase (phaA) and acetoacetyl-CoA reductase (phaB) from Ralstonia eutropha and butyryl-CoA dehydrogenase (bld) from Clostridium saccharoperbutylacetonicum (Fig. 1). Under the optimized conditions, this recombinant strain can produce 9.05 g/L 1,3-BDO (Table 1) (Kataoka et al. 2013). A semi-biotechnological route through enzymatic asymmetric reduction of 4-hydroxy-2-butanone to 1,3-BDO was also reported by research groups using different microbial strains, e.g., Candida krusei ZJB-09162 and Pichia jadinii (Yang et al. 2014; Zheng et al. 2012). To produce 1,3-BDO at the commercial scale, there is a need to develop an integrated technology platform through engineering microbe metabolic pathways and improving the fermentation and downstream recovery process.

5 Biological production of diamines

5.1 Cadaverine

Cadaverine was discovered more than 100 years ago and is found in few microbes as a minor polyamine. It plays a role in the growth and function of cells (Tabor and Tabor 1985; Wallace et al. 2003). Cadaverine is used for bio-nylon production and has many other applications in medicine and agriculture (Becker et al. 2011; Schneider and Wendisch 2011). Biotechnologically, cadaverine can be produced by direct microbial fermentation or by biotransformation of lysine using whole cells or free enzyme (Fig. 1) (Bhatia et al. 2015a; Kim et al. 2015; Qian et al. 2011). Lysine conversion is catalyzed by lysine decarboxylase (Fig. 1). E. coli has two types of lysine decarboxylase, inducible lysine decarboxylase (cadA) and constitutive enzyme (ldcC) (Kikuchi et al. 1997). C. glutamicum is a promising strain for the production of lysine but it lacks lysine decarboxylase activity. Mimitsuka et al. engineered C. glutamicum by introducing cadA from E. coli and deleting homoserine dehydrogenase (hom) to produce 2.6 g/L cadaverine through glucose fermentation (Table 1) (Mimitsuka et al. 2007). To further increase cadaverine production, Kind et al. (2010a) engineered a central supporting pathway for lysine production in C. glutamicum by introducing point mutations in aspartokinase and pyruvate carboxylase to avoid feedback regulation. To ensure the supply of the precursor oxaloacetate, pyruvate carboxylase was overexpressed and phosphoenolpyruvate carboxykinase, which removes oxaloacetate, was deleted. Many other genes were overexpressed (aspartokinase, dihydrodipicolinate reductase, diaminopimelate dehydrogenase and diaminopimelate decarboxylase), and the engineered C. glutamicum strain resulted in markedly increased cadaverine production (Kind et al. 2010a). Lysine decarboxylase (cadA) is a bottleneck because it is inhibited by the end product inside the cell. Kind et al. further engineered C. glutamicum by expressing a permease encoded by cg2893, which enhanced the export of cadaverine by 20 % (Kind et al. 2011). The highest production of cadaverine (88 g/L) was reported for a C. glutamicum strain in which the native promoter of the genes involved in lysine production was replaced with a sod promoter (superoxide dismutase) and the ldc gene promoter was replaced with a tuf promoter (Kind et al. 2014).

C. glutamicum possesses a cadaverine degradation pathway that leads to the formation of N-acetylcadaverine, which is an undesired product that causes problems in product purification (Kind et al. 2010b). E. coli is a suitable alternative because it lacks the cadaverine metabolizing pathway. Qian et al. (2011) engineered E. coli by overexpressing dihydrodipicolinate synthase (dapA) to increase the l-lysine pool and cadA to convert lysine into cadaverine. The final engineered strain produced 9.6 g/L cadaverine (Table 1). The production of l-lysine has reached approximately 2,200,000 tons/year using a mutant strain of C. glutamicum (Eggeling and Bott 2015). Simple biotransformation is another approach to produce cadaverine using radially available l-lysine (Bhatia et al. 2015a). Kim et al. reported that E. coli overexpressing cadA can directly convert crude lysine into cadaverine with 80 % conversion in the presence of the cofactor pyridoxal-5′-phosphate (PLP) (Kim et al. 2015). To further improve cadaverine production, Bhatia et al. immobilized E. coli cells using a barium alginate matrix and used the E. coli for the biotransformation of lysine to cadaverine. The immobilized cells produced 75.8 g/L cadaverine with 84 % conversion and could be reused for several cycles (56 % residual activity after the 18th cycle) (Bhatia et al. 2015a). The addition of the cofactor PLP increases the cost of cadaverine production. Ma et al. engineered an E. coli strain for PLP synthesis by heterologously expressing pdxS and pdxT of Bacillus subtilis and increased the PLP pool by 1144 nmol/g DCW, which resulted in 2.9-fold cadaverine production (Ma et al. 2015).

To extend the substrate range for cadaverine production, researchers engineered an E. coli strain to display beta-glucosidase (BGL) on its surface and to overexpress cadA to enable it to utilize cellobiose as a carbon source (Ikeda et al. 2013). Tateno et al. (2009) reported the one-step production of cadaverine from soluble starch using an engineered strain of C. glutamicum overexpressing α-amylase from Streptococcus bovis 148 and cadA from E. coli. Cadaverine production was extended to additional microbes, such as Bacillus methanolicus, which can utilize methanol as a carbon source. A thermophilic B. methanolicus overexpressing cadA and ldcC from E. coli produced 11.3 g/L cadaverine in fed-batch methanol fermentation (Naerdal et al. 2015). The cadaverine produced by these strains can be combined with various diacids to produce different types of nylon.

5.2 Putrescine

Putrescine is a biogenic amine of industrial interest that can be used as a building block for PA-4,6 and PA-4,4 (Scott et al. 2007). Putrescine is synthesized commercially by a chemical process, starting from succinonitrile, which is produced via the addition of hydrogen cyanide to acrylonitrile (Lammens et al. 2011). Biotechnologically, putrescine can be produced from l-ornithine or l-arginine decarboxylation catalyzed by ornithine decarboxylase and arginine decarboxylase (Schneider and Wendisch 2011). C. glutamicum is a promising host for putrescine production as it can tolerate a higher concentration of putrescine than E. coli (Schneider and Wendisch 2010). Schneider et al. engineered C. glutamicum heterologously overexpressing ornithine and arginine decarboxylase from E. coli and reported the ornithine pathway is 40 times more efficient than that of arginine (Fig. 1). Further improvement of the strain by deletion of the arginine repressor (ArgR) and the ornithine carbamoyltransferase (ArgF) resulted in 6 g/L putrescine production in shake flask batch culture fermentation (Table 1) (Schneider and Wendisch 2010).

To further improve the purity of putrescine, a research group deleted the N-acyltransferase gene responsible for the acylation of putrescine and other amines from the genome of C. glutamicum. The engineered strain accumulated 41 % more putrescine because there was no accumulation of acetylputrescine (Nguyen et al. 2015). Nguyen et al. reported that the expression of the putrescine transporter (CgmA), which was identified as a cadaverine transporter by Kind et al., can increase putrescine production by up to 24 % compared to the parental strain (Nguyen et al. 2015). C. glutamicum expressing the ornithine decarboxylase gene (speC) using the argF plasmid addiction system produced 19 g/L putrescine in fed-batch fermentation (Schneider et al. 2012). The highest production of putrescine (24.2 g/L) was recorded in E. coli engineered by deleting the ornithine carbamoyltransferase (argI) and overexpressing ornithine synthesizing genes and ornithine decarboxylase under a tac or trc promoter. To increase the substrate range for putrescine production, Meiswinkel et al. engineered C. glutamicum for glycerol utilization by expressing the glycerol transporter (glpF), glycerol kinase (glpK) and glycerol-P dehydrogenase (glpD) from E. coli (Meiswinkel et al. 2013b), and the utilization of xylose as the carbon source was achieved by overexpressing xylose isomerase from Xanthomonas campestris (Meiswinkel et al. 2013a). Putrescine production was also reported for an E. coli strain engineered by inactivating putrescine degradation genes and deleting the ornithine carbamoyltransferse gene. The ornithine pool was increased by expressing ornithine synthesis genes under the control of the trc promoter. The final engineered strain produced 1.68 g/L putrescine using glucose media (Qian et al. 2011).

6 Biopolymer synthesis

Condensation is the most common method for polymer synthesis, and different building blocks, such as diacids, diols, and diamines, can be used to produce polyester and polyamide biopolymers (Fig. 3). Various biopolymers, such as poly(hydroxybutyrate) (PHB), poly(butylene succinate) (PBS), poly(ethylene terephthalate) (PET), and polybutylene terephthalate (PBT), are produced commercially (Chuah 2004). The polymerization of polyesters and polyamides is performed using common basic steps: (1) step-growth polycondensation of diacids/diamines and (2) ring-opening polymerization (Jiang and Loos 2016).

Fig. 3
figure 3

General scheme for polyester and polyamide polymerization

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Step-growth polycondensation requires bifunctional monomers, and polymerization occurs in both directions in a stepwise manner when a functional group from one monomer reacts with a functional group of another monomer (Billiet et al. 2009). The byproduct produced is water or methanol, depending on the reactant used, namely, dicarboxylic acid or its dimethylester. The polycondensation reaction is performed in the presence of a catalyst, such as antimony, germanium, titanium or aluminium. (Haldimann et al. 2013).

Ring-opening polymerization is a form of chain-growth where the terminal end of a polymer chain acts as a reactive center and other cyclic monomers react by opening its ring system to form a long polymer chain (Nuyken and Pask 2013). In contrast to the other polycondensation reaction, this method does not require the removal of end byproducts, and polymers of high molecular weight can be achieved in a few minutes. Various polymer syntheses have been reported using this method, including PET and PBT. The main disadvantage of this method is the requirement that one of the monomers (diol or diacid) has to be converted to its cyclic form, which requires additional solvents and a catalyst. The chemical methods traditionally used for polymerization require high temperature (150–280 °C) and metal catalysts that are toxic to the environment (Gross et al. 2010).

Biotechnologically, various enzymes can be used for polyester and polyamide synthesis. Enzymatic methods are advantageous because they can be performed under mild conditions and do not require protection deprotection reactions. Hydrolase (lipase and esterase)-catalyzed esterification and transesterification reactions are used for polyester polycondensation. Lipase is the most dominant enzyme used for polymerization; in nature, it generally catalyzes the hydrolysis of fatty acid glycerol esters. It can be used to perform polymerization in non-aqueous media and to remove the side products of the reaction. Candida antarctica lipase B (CALB) is the most widely studied and used enzymatic system due to its high regioselectivity, chemoselectivity and enantioselectivity and high thermal stability and activity (Kobayashi 2009). Ren et al. (2015) reported an enzymatic process for high-molecular-weight poly(butylene succinate) synthesis from succinic anhydride and butane-1,4-diol using Novozym 435 (an immobilized CALB).

In addition to lipase, other enzymes, such as cutinase and pha synthase, can catalyze polymerization (Hunsen et al. 2007; Jia et al. 2016). Hunsen et al. (2007) used cutinase from Humicola insolens and produced polyesters of AA with diols of various carbon chain lengths. PHA synthase catalyzes the esterification of (R)-hydroxyalkanoate-CoA monomers subunits to produce poly(hydroxyalkonates) (Bhatia et al. 2015b). Lipase can also be used for hydroxyalkonoate polymerization, but it requires a longer reaction time and results in low molecular weight polymers (Feder and Gross 2010). Polyamide biopolymers (nylon) are widely used due to their resistance and good mechanical properties. Nylon is the most common and abundantly used polymer for various purposes, including furniture, packaging and electrical devices (Kind et al. 2014). Lipase-mediated polymerization reactions are not well studied for polyamide synthesis due to the poor solubility of polyamides (Garcia Linares and Baldessari 2013). Cheng et al. (2004) first reported the lipase-mediated polymerization of polyamides using different dialkyl esters (adipate, fumarate, phenylmalonate) and amines (triethylenetetramine, diethylenetriamine).

7 Downstream processing and applications

The downstream processing of biopolymers (recovery/purification) is a huge part of the total production cost (Samori et al. 2015). Various methods have been developed for biopolymer recovery, i.e., chemical methods, enzymatic methods and the use of an engineered strain capable of self-lysis (Riedel et al. 2013; Yang et al. 2015b). Chemical methods use organic solvents and sodium hypochlorite, which are toxic to the environment. Enzymatic methods are easy and ecofriendly, but cost is a major issue preventing the further use and development of these methods. Researchers have developed a method of automatic cell lysis for biopolymer liberation, which is currently gaining attention. Initially, this system was developed using the E. coli phage lysis genes of the bacteriophage PhiX174 combined with phaCAB along with a thermoactive control system for lysis gene expression (Resch et al. 1998). Recently, a new method using the predatory bacterium Bdellovibrio bacteriovorus HD100, which attacks Gram-negative bacteria and helps to release the stored polymer, was reported (Martínez et al. 2016).

Many of the described processes are too costly and inefficient for large-scale use. Heinrich et al. developed a PHA extraction method from Ralsotonia eutropha at 50 L scale using sodium hypochlorite and recovered PHA with 91 % efficiency (Heinrich et al. 2012). Other physical methods, such as homogenization followed by centrifugation coupled with organic solvents, have also been investigated, but no method has been developed with high purity and recovery (Li et al. 2007). Biopolymer extraction is still a challenging task because the use of hazardous chemicals is an unacceptable option to produce ecofriendly material. Biopolymers have many applications in the textile, pharmaceutical, and medical sectors, such as packaging, dental implants, drug delivery, dendritic material, and electrical engineering (Table 3) (Chuah 2004; Hemalatha et al. 2016). Various building blocks, their possible polyesters and polyamides and their application are listed in Table 2.

Table 2 Various polymers produced using biobased building blocks and their potential applications
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8 Discussion

The biopolymer production cost mainly depends on the yield of the biopolymer relative to the amount of carbon source used and the efficiency of the downstream process. Microbial systems are an area of interest for the production of various building blocks due to the availability of genetic engineering techniques, which provide advantages for the production of biopolymers with the desired properties. Most biotechnological methods developed for the production of building blocks depend on the utilization of pure carbohydrates as the carbon source, e.g., sugars, which are called first-generation carbon sources (Bai et al. 2008). The annual production of glucose is approximately 30 million tons with a selling price of $0.39/kg, and the annual production of glycerol is approximately 37 billion gallons with a selling price of $0.66/kg (Song and Lee 2006; Yang et al. 2012). The raw material cost is approximately 60 % of the total production cost (Song and Lee 2006). Recently, researchers have started to focus on the use of second-generation carbon sources, e.g., lignocellulose and cellulose, for microbial fermentation (Naik et al. 2010). The utilization of biomass also presents several challenges because it consists of 10–35 % lignin, which is recalcitrant for breakdown by microbes (Wang et al. 2011). This organic waste can be converted into a mixture of gas, i.e., syngas (CO2, CO and H2) by gasification (Richardson et al. 2012). Syngas is a third-generation raw material and can be metabolized by certain carbon fixing microorganisms into multi-carbon compounds, such as acetate, ethanol, butyrate, 2,3-butanediol and PHA (Choi et al. 2010; Latif et al. 2014). The use of mixed culture integrated with dark fermentation can further reduce production costs by up to 30–40 % (Du et al. 2012). Mixed culture fermentation is advantageous over pure culture as there is no need to maintain aseptic conditions, and it can use a wide range of materials as the carbon source, which may further decrease production costs (Bastidas-Oyanedel et al. 2015). Jiang et al. (2012) developed a mixed culture process for PHA production using paper mill waste.

Even with the development in biotechnology, microbial fermentation routes cannot compete with organic synthesis. To achieve the industrial production of building blocks, metabolic engineering is an important tool, and E. coli is the predominant strain due to recent developments in systems and synthetic biology. To develop a sustainable process, several issues must be addressed, e.g., utilization of an inexpensive raw material, elimination of bottlenecks, improvement of the biocatalyst and understanding of the regulatory mechanism for better productivity. Further advancements and the introduction of new technology, such as genome sequencing, genome engineering, and functional genomics, will lead to the development of new metabolic routes for building blocks with increased resistance to inhibitors and environmental factors. For the production of biobased chemicals from an industrial perspective, in addition to these advancements, improvement of the downstream processing is necessary to develop an efficient bioprocess. The introduction of new technology and the development of new biocatalytic bioprocesses will reduce the dependency on petroleum-based chemicals and open provide new opportunities for a biobased economy. Currently, few biobased polymers (polyesters/polyamides) are commercially available, e.g., PHB, PBS, PET, PTT, nylon-4,6 and nylon-4,10 (Jacquel et al. 2015; Jiang et al. 2016). Various polymers produced at the commercial scale by different companies and with their trademarks are given in Table 3.

Table 3 Polyester and polyamide biopolymers produced at the commercial scale
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9 Perspectives and conclusion

This review provides broad information about the development of microbiological routes for the production of biopolymer building blocks. Few microbes have the natural ability to produce biopolymers, and multiple heterologous enzymes are required to construct a new pathway. Advancements in metabolic engineering and fermentation technology have enabled microbial cells to produce various biopolymer building blocks. Biopolymer production using biological processes is not feasible due to the high cost of the raw materials, the low productivity, the coproduction of side products and the costly recovery process. To make the biological process more competitive, the following issues must be considered because the biopolymer building blocks should be produced in large scale with reduced prices.

  • Due to the higher price of cofactors and the lower productivity of the final product, the development of cofactor regeneration systems, such as PLP, FAD and NAD(P)H are essential to reduce the production cost of the desired products.

  • The engineering of a microbial strain to tolerate various inhibitors (e.g., hydroxymethylfurfural, furfural, vanillin, and acetate), which are produced during biomass pretreatment and affect microbial fermentation (Bhatia et al. 2016), and the utilization of biomass hydrolysate as a carbon source are needed.

  • The polymerization of building-block monomers is a more difficult process and depends on traditional chemical reactions. To make the process more ecofriendly and economic, there is need to find new enzymes and to improve the activity and selectivity of the current enzymes.

  • The development of biotransformation reactions for one- or two-step processes could be a more successful method than the construction of a long metabolic pathway.

The biological method of biopolymer production can reduce the dependence on non-renewable resources and help to decrease greenhouse gases. With the continuous and joint efforts of researchers, industry and government, biological processes can become a green alternative for polymer production in the near future.