Introduction

Plant biomass potentially provides a renewable source for the production of different fuels and chemicals. One promising route involves the use of native or engineered microorganisms, which can convert the sugars present in plant biomass into a wide range of value-added products (Isikgor and Becer 2015). Much of this work has focused on the conversion of glucose, the most abundant sugar in plant biomass. However, many other sugars are also present in plant biomass (Isikgor and Becer 2015; Jagtap et al. 2012, 2013; Rubin 2008; Tai et al. 2016). Key among them is D-xylose, a pentose (five-carbon) sugar, hereafter referred to simply as xylose. It represents 30–40% of the sugars recoverable from plant biomass (Dhiman et al. 2012, 2013; Hamacher et al. 2002; Himmel et al. 2007; Jagtap et al. 2014a, b; Jeffries 1983; Jeffries and Shi 1999; Rubin 2008). Not surprising, many different microorganisms have been explored for the conversion of xylose to different products (Jeffries 1983, 2006; Jeffries et al. 2007; Johnsen et al. 2009; Moysés et al. 2016; Nunn et al. 2010; Stephens et al. 2007). Much of this work has focused on the conversion of xylose to ethanol (Gong et al. 1981; Jeffries 2006; Millati et al. 2005; Mohd Azhar et al. 2017; Toivola et al. 1984; Zhang et al. 1995; Zhou et al. 2016). However, more recently, a number of studies have shown that microbes can convert xylose to other valuable fuels and chemicals—in some cases with better results than those obtained from glucose (Brat and Boles 2013; Feng et al. 2015; Guo et al. 2016b; Jordan et al. 2016; Kim et al. 2014; Kwak et al. 2017; Lee et al. 2015; Lian and Zhao 2015; Mohd Azhar et al. 2017; Moysés et al. 2016; Tippmann et al. 2013; Turner et al. 2015). Moreover, some of these chemicals, such as sugar alcohols, are natural by-products of xylose metabolism, indicating that xylose may provide the ideal source for their production (Jagtap and Rao 2018; Jeffries 2006; Jeppsson et al. 2006; Kwon et al. 2006; Lin et al. 2005; Moysés et al. 2016; Rafiqul and Sakinah 2013).

While xylose represents an abundant and renewable feedstock for many microbial conversion processes, its metabolism is not native to many production strains such as Saccharomyces cerevisiae and Zymomonas mobilis (Jeffries 2006; Toivari et al. 2004; van Zyl et al. 1989; Zhang et al. 1995). As a consequence, these microorganisms need to be genetically engineered to utilize xylose, a process that involves many genetic modifications in addition to expressing the heterologous enzymes. Even in microorganisms where xylose metabolism is present, it is often inefficient as compared to glucose metabolism and thus often requires further optimization.

In addition, developing efficient and economical processes for the conversion of plant biomass into different fuels and chemicals will require microorganisms capable of utilizing both glucose and xylose, the two principle sugars in plant biomass. However, xylose metabolism is often subject to glucose catabolite repression, where the cells will first consume glucose before they consume the xylose (Farwick et al. 2014; Gancedo 1998; Görke and Stülke 2008; Meijer et al. 1998). This phenomenon reduces the productivity of fermentation processes involving mixtures of glucose and xylose, because it lengthens the fermentation times. Ideally, microorganisms could be identified or engineered to simultaneously consume these two sugars (Kim et al. 2012).

Not surprisingly, the development of microorganisms capable of efficiently converting xylose to different fuels and chemicals has attracted significant attention over the years and still is a vibrant area of research (Jagtap and Rao 2018; Ledesma-Amaro et al. 2016; Liu et al. 2015; Park et al. 2017; Zhang et al. 2016a, b). Much of this effort, as noted above, has focused on engineering microorganisms capable of converting xylose into ethanol. A number of excellent reviews have already been written on this topic (Hahn-Hagerdal et al. 2007a; Jordan et al. 2012; Lin and Tanaka 2006; Matsushika et al. 2009; Mohd Azhar et al. 2017; Zaldivar et al. 2001). As a consequence, we limit our discussion to the conversion of xylose to products other than ethanol. Our review mostly focuses on yeast and fungi for brevity’s sake, aside from discussing the different pathways involved in xylose metabolism. Of course, bacteria also provide promising hosts for xylose conversion, particularly in case of the production host and native xylose-utilizer Escherichia coli. In addition, multiple strains of the bacterium Zymomonas mobilis—which are capable of producing ethanol at high rates, yields, and titers—have been engineered to efficiently utilize xylose (Agrawal et al. 2011; Dunn and Rao 2014, 2015; Jeon et al. 2005).

Xylose metabolism

Microorganisms primarily catabolize xylose through three different pathways (Fig. 1). Bacteria most commonly employ the isomerase pathway for conversion of xylose to D-xylulose using xylose isomerase (XI). D-xylulose is then phosphorylated by xylulose kinase (XK), yielding D-xylulose-5-phosphate, which then enters central metabolism through the pentose phosphate pathway. The XI pathway is found in diverse prokaryotes (Jeffries 1983; Lajoie et al. 2016; Schellenberg et al. 1984; Umemoto et al. 2012). In addition, it is also found in a few anaerobic fungi such as Piromyces and Orpinomyces (Brat et al. 2009; Gárdonyi and Hahn-Hägerdal 2003; Harhangi et al. 2003; Madhavan et al. 2008; Nierman et al. 2001; Sarthy et al. 1987; Walfridsson et al. 1996).

Fig. 1
figure 1

Xylose metabolic pathways in microorganisms. XR, D-xylose reductase; XDH, xylitol dehydrogenase; XI, D-xylose isomerase; XK, xylulokinase; PK, phosphoketolase; PTA, phosphotransacetylase; ACK, acetate kinase; ACS, acetyl-CoA synthetase; XDH, D-xylose dehydrogenase; XL, xylonolactonase; XAD, xylonate dehydratase; KDXA, 2-oxo-3-deoxy xylonate aldolase; KDXD, 2-oxo-3-deoxy xylonate dehydratase; and αKGSADH, α-oxoglutaric semialdehyde dehydrogenase

Full size image

Archaea, along with some bacteria such as Caulobacter crescentus, possess oxidative xylose metabolic pathways called the Weimberg and Dahms pathways (Johnsen et al. 2009; Nunn et al. 2010; Stephen Dahms 1974; Weimberg 1961). In the Weimberg pathway, xylose is oxidized to D-xylonolactone by D-xylose dehydrogenase, followed by a lactonase to hydrolyze the lactone to D-xylonate. Xylonate dehydratase then acts on D-xylonate to produce 2-keto-3-deoxy-D-xylonate and a second dehydratase forms the α-keto-semialdehyde. The α-keto-semialdehyde is subsequently oxidized to alpha-ketoglutarate, an intermediate of the tricarboxylic acid cycle (TCA) (Fig. 1) (Weimberg 1961). The Dahms pathway is similar to the Weimberg pathway except that the 2-keto 3-deoxy-D-xylonate is cleaved by an aldolase to form pyruvate and glycoaldehyde (Stephen Dahms 1974).

Xylose-utilizing yeasts and some filamentous fungi employ an oxido-reductive pathway consisting of two enzymatic reactions for xylose utilization (Fig. 1) (Jeffries 2006; Moysés et al. 2016). First, xylose is reduced to D-xylitol by xylose reductase (XR) using NADH or NADPH as a cofactor, depending on the enzyme. Some XRs solely utilize NADPH, whereas others can utilize both NADPH and NADH, typically with higher specificity for the former (Lee 1998; Schneider et al. 1989). The ability of some yeasts to utilize xylose anaerobically has been attributed to this dual cofactor specificity (Bruinenberg et al. 1984). Next, D-xylitol is oxidized to D-xylulose by the strictly NAD+-dependent xylitol dehydrogenase (XDH) (Kotter et al. 1990). D-xylulose is then phosphorylated to D-xylulose-5-phosphate by xylulokinase (XK) (Jin et al. 2002). Xylulose-5-phosphate is a metabolic intermediate of the nonoxidative pentose phosphate pathway (Stincone et al. 2015). Yeasts metabolize D-xylulose-5-phosphate to various phosphorylated sugars such as fructose-6-phosphate and glyceraldehyde-3-phosphate, which enters into glycolysis pathway (Jeffries 2006; Moysés et al. 2016). The difference in cofactor specificity in the first two steps of xylose metabolism can result in cofactor imbalance when oxygen or respiration is limiting (Dijken and Scheffers 1986). The redox imbalances during conversion of xylose into xylulose form a bottleneck in pentose fermentation for many yeasts.

Some oleaginous yeasts such as Rhodotorula graminis and Rhodotorula glutinis use the phosphoketolase (PK) pathway to convert xylulose-5-phosphate to glyceraldehyde-3-phosphate and acetyl phosphate (Whitworth and Ratledge 1977). Glyceraldehyde-3-phosphate further enters in glycolysis pathway. Acetate kinase (ACK) and acetyl-CoA synthase (ACS) convert acetyl phosphate into acetyl-CoA (Ingram-Smith et al. 2006).

Xylitol production

Xylitol is a five-carbon sugar alcohol used as a natural sweetener and sugar substitute. In addition, it has been shown to prevent tooth decay and, as a consequence, is often used as a sugar substitute for chewing gum (Saha et al. 2007). It is also one of the Department of Energy’s top 12 bio-based building block chemicals (Werpy et al. 2004). Xylitol can be produced chemically from xylose (Rafiqul and Sakinah 2012). It can also be produced using biological methods (Albuquerque et al. 2014; Jordan et al. 2012; Kwak and Jin 2017; Lane et al. 2018; Rafiqul and Sakinah 2013; Saha and Bothast 1997). A number of microorganisms naturally produce xylitol during growth on xylose (Table 1). In particular, many yeasts employ the XR-XDH pathway for xylose metabolism. Xylitol is a metabolic intermediate in this pathway, the product of the NADPH-dependent xylose reductase (XR) that catalyzes the conversion of xylose to xylitol. Xylitol is then oxidized to xylulose by the NAD+-dependent xylitol dehydrogenase (Jeffries 2006; Moysés et al. 2016). Xylitol production in these yeasts is known to result from a redox imbalance associated with the different cofactor specificities for these two enzymes. These imbalances tend to be magnified during growth under oxygen-limited conditions due to the inability of the cells to regenerate NAD. Many studies have focused on five main engineering strategies for improving xylitol production from xylose: overexpression of heterologous or native genes involved in xylitol production pathway, altering the cofactor specificity by deleting or overexpressing relevant genes, disruption of XDH gene, expressing heterologous xylose transporters, and optimizing culture conditions (Pal et al. 2016).

Table 1 List of xylitol production during growth on xylose by wild-type and engineered microorganisms
Full size table

Fungi

Multiple filamentous fungi produce xylitol during aerobic growth on xylose (Table 1). In one study, the authors screened 11 different fungi for their ability to produce xylitol during growth on 11.5 g/L xylose (Sampaio et al. 2003). All were found to produce xylitol with titers ranging from 0.14 to 0.52 g/L. While these titers are relatively low, other fungi have been found to produce xylitol at far higher titers. For example, Petromyces albertensis produces 36.8 g/L xylitol from 100 g/L xylose (Dahiya 1991). The addition of methanol was found to further increase xylitol production to 39.8 g/L xylitol. The oxidation of methanol yields NADH, which is utilized for the reduction of xylose to xylitol by the NADH-dependent xylose reductase (Dahiya 1991).

In addition to native production, filamentous fungi have also been engineered to produce xylitol from xylose. For example, Ashbya gossypii was engineered to produce 22.6 g/L xylitol from 80 g/L xylose by overexpressing the native genes encoding for xylose reductase (GRE3), xylitol dehydrogenase (XYL2), and xylulose kinase (XKS1) (Díaz-Fernández et al. 2017). Trichomderma reesei has also been engineered for xylitol production by reducing the activity of XDH. In one study, RNA interference was used to reduce the expression of XDH in T. reesei, which increased xylitol production 5-fold and yielded titers of 2.37 g/L (Wang et al. 2005). In a second study, T. reesei was engineered to produce xylitol by deleting XDH and overexpressing XR, which increased xylitol production from 0 to 3.72 g/L (Hong et al. 2014).

Yeast

Multiple yeasts from diverse genera such as Candida, Clavispora, Hansenula, Issatchenkia, Kluyveromyces, and Pichia have been have shown to produce xylitol during aerobic growth on xylose in defined media (Barbosa et al. 1988; Gong et al. 1981; Kim et al. 2004; Oh and Kim 1998; Rao et al. 2007; Sampaio et al. 2008; Suryadi et al. 2000; Vandeska et al. 1995; Wang et al. 2005). Candida tropicalis and Candida guilliermondi exhibit some of the best yields, with reported values of 0.80 and 0.74 g/g, respectively (Table 1) (Barbosa et al. 1988; Gong et al. 1981). The highest xylitol productivity (12 g/L/h) from xylose using a submerged membrane bioreactor was reported in Candida tropicalis with glucose as co-substrate (Kwon et al. 2006).

In addition to native production, yeasts have also been engineered to produce xylitol. For example, the expression of heterologous XR, disruption of XDH, and overexpression of heterologous transporters in Candida tropicalis and Kluyveromyces marxianus produced higher xylitol as compared to wild-type strains (Jeon et al. 2012, 2013; Lee et al. 2003b; Zha et al. 2013; Zhang et al. 2013, 2014, 2015). A codon-optimized XR from Neurospora crassa was expressed in C. tropicalis LNG2 where the gene encoding xylitol dehydrogenase (XYL2) was disrupted (Jeon et al. 2012). The resulting strain produced 48 g/L of xylitol during fed-batch growth (Jeon et al. 2012). In another study, the NADH preferring XR from C. parapsilosis was expressed in C. tropicalis BN1. The resulting strain produced 280 g/L xylitol with productivity of 5.09 g/L/h during fed-batch growth (Lee et al. 2003b). In another study, the xylose transporter gene CoXT2 from Arabidopsis thaliana was integrated into the genome of a C. tropicalis strain where XYL2 was disrupted and the xylose reductase from N. crassa expressed. This strain produced xylitol with productivity of 1.14 g/L/h (Jeon et al. 2013). The three transporter genes KmFPS1 encoding aquaglyceroporin from Kluyveromyces marxianus, CiGXF1 encoding Candida intermedia glucose/xylose facilitator, and CiGXS1 encoding glucose/xylose symporter gene were expressed in K. marxianus YZJ074 (Zhang et al. 2014). This strain produced 71.5 g/L xylitol from 100 g/L xylose with productivity of 1.49 g/L/h at 42 °C (Zhang et al. 2014).

Native strains of S. cerevisiae cannot grow on xylose (Jeffries 1983). S. cerevisiae has been engineered for xylose utilization and transport, yielding strains capable of producing high levels of xylitol (Hahn-Hagerdal et al. 2007b; Jeppsson et al. 2006; Saloheimo et al. 2007). The δ-integration vector for chromosome integration and YRp-based episomal plasmid vector were used to introduce XR in S. cerevisae (Chung et al. 2002). Chromosomal integration showed better mitotic stability of the XR gene along with a high expression level in S. cerevisae and yielded 1.7-fold improvement in xylitol productivity in fed-batch culture (Chung et al. 2002). S. cerevisiae was also engineered to produce xylitol by overexpressing XR and XDH from Pichia stipitis and overexpressing the native transaldolase (TAL) and transketolase (TKL). This strain produced 2 g/L xylitol with a yield of 0.82 g/g xylose (Walfridsson et al. 1997). Expression of xylose reductase from P. stipitis in a S. cerevisiae strain produced 19.2 g/L xylitol (Zha et al. 2013). S. cerevisiae has also been engineered to produce xylitol by expressing CDT1, GH1-1, a β-glucosidase from N. crassa, and two copies of XYL1 from P. stipitis. This strain is able to consume both xylose and cellobiose to produce xylitol with a yield of 1 g/g xylose (Oh et al. 2013; Zha et al. 2013).

Bacteria

A few bacteria possess the XR-XDH pathway for producing of xylitol from xylose (Rangaswamy and Agblevor 2002; Yoshitake et al. 1973). For example, Enterobacter liquefaciens produced 28 g/L xylitol during growth on 100 g/L xylose as the sole carbon source (Yoshitake et al. 1973). Corynebacterium sp. B424 has been shown to produce 10 g/L xylitol, the highest among 17 screened cultures (Rangaswamy and Agblevor 2002).

Escherichia coli has also been engineered to produce xylitol from xylose (Häcker et al. 1999; Suzuki et al. 1999). Replacement of the native cyclic AMP receptor protein (CRP) with a cyclic AMP-independent mutant (CRP*) facilitated xylose utilization in mixtures of xylose and glucose, where glucose serves as source of cell growth and reducing equivalents (Cirino et al. 2006). In addition, E. coli expressing XR genes from Candida tenuis and C. tropicalis produced 13.3 g/L xylitol from 50 g/L xylose (Häcker et al. 1999; Suzuki et al. 1999).

A number of other bacteria have been engineered to produce xylitol from xylose (Kim et al. 2010; Nyyssölä et al. 2005; Sasaki et al. 2010; Suzuki et al. 1999; Yoshitake et al. 1973). Corynebacterium glutamicum was engineered to produce xylitol by integrating the pentose transporter gene (araE), disrupting the native lactate dehydrogenase gene (ldhA) and expressing a mutant XR (K274R) (Sasaki et al. 2010). The resulting strain produced 26.5 g/L xylitol. The genes encoding xylulokinase (XylB) and the phosphoenolpyruvate-dependent fructose phosphotransferase (PTSfru) were also expressed in this strain to eliminate intracellular xylitol phosphate formation. The resulting strain produced 166 g/L xylitol with productivity of 7.9 g/L/h from a mixture of xylose and glucose during fed-batch growth (Sasaki et al. 2010). Lactococcus was also engineered to produce 160 g/L xylitol with productivity of 1.75 g/L/h during fed-batch growth from a mixture of xylose and glucose by overexpressing the native xylose transporter and XYL1 from P. stipitis (Nyyssölä et al. 2005).

Arabitol production

D-arabitol is a five-carbon sugar alcohol, derived from xylose, that can be also used as a natural sweetener and sugar substitute (Werpy et al. 2004). It is also listed, along with xylitol, as one of the Department of Energy’s top 12 bio-based building block chemicals. Multiple yeasts are known to produce arabitol during growth on glucose (Bernard et al. 1981; Bisping et al. 1996; Blakley and Spencer 1962; Escalante et al. 1990; Hajny 1964; Moran and Witter 1979; Van Eck et al. 1993; Wilson and Mortlock 1973). Far fewer are reported to produce arabitol from xylose (Lin et al. 2005; Saha et al. 2007). However, arabitol is linked to xylose metabolism through arabitol dehydrogenase, which reduces xylulose to arabitol using NADH as the cofactor. In these regards, arabitol can potentially be produced as an overflow metabolite associated with redox imbalances during growth on xylose, where the production of arabitol by arabitol dehydrogenase regenerates NAD. In support of this mechanism, a recent study demonstrated that the oleaginous yeast Rhodosporidium toruloides produces 49 g/L arabitol during growth on 150 g/L xylose in rich media; however, during growth on low-nitrogen media, which is commonly used to induce lipid production in oleaginous yeast, no arabitol was produced (Table 2) (Jagtap and Rao 2018). These differences were associated with the rates of xylose utilization. R. toruloides consumes xylose at far greater rate in rich medium than in low-nitrogen medium. Likely, these high rates of xylose utilization induce a redox imbalance that results in the production of arabitol. This would also suggest that additional yeasts produce arabitol, assuming they have arabitol dehydrogenase.

Table 2 List of erythritol and arabitol production from xylose by yeast
Full size table

Erythritol production

Erythritol is a four-carbon polyol produced by microorganisms as an osmoprotectant (Blomberg and Adler 1992; Carly and Fickers 2018; Spencer and Sallans 1956). Erythritol is a noncaloric sweetener, about 60–70% as sweet as sucrose with similar taste and texture (Carly and Fickers 2018). The human body cannot metabolize erythritol, despite being quickly absorbed by the small intestine, and is rapidly excreted in urine. Unlike other polyols, erythritol is not a laxative (Hiele et al. 1993; Livesey 2001). Erythritol is also noncariogenic and shown to prevent dental plaque on a long-term basis (Falony et al. 2016; Mäkinen et al. 2005).

Erythritol can be produced chemically from xylose (Nabors and Gelardi 2001). It can also be produced by bacteria, yeasts, and fungi during growth on xylose (Guo et al. 2016a; Jovanović et al. 2014; Veiga-da-Cunha et al. 1993). In bacteria, xylose is converted to xylulose-5-phosphate using the XI-XK pathway. Xylulose-5-phosphate enters into pentose phosphate pathway to generate fructose-6-phosphate. The phosphoketolase enzyme splits fructose-6-phosphate into erythrose-4-phosphate and acetyl phosphate. An erythrose-4-phosphatase converts erythrose-4-phosphate to erythrose, which is further converted into erythritol by erythrose reductase (ER) (Moon et al. 2010; Veiga-da-Cunha et al. 1993). Acetyl phosphate is either converted into acetate by ACK or into acetyl-CoA by phosphate acetyltransferase (PTA).

In yeasts, xylose is converted to xylulose-5-phosphate by an action of XR, XDH, and XK. Xylulose-5-phosphate enters pentose phosphate pathway to synthesize erythrose-4-phosphate. Erythrose-4-phosphatase dephosphorylates erythrose-4-phosphate into erythrose. An erythrose reductase converts erythrose into erythritol with NADPH as the cofactor (Mirończuk et al. 2017; Rzechonek et al. 2018). The genes encoding erythritol reductase have been identified in multiple yeast (Janek et al. 2017; Kobayashi et al. 2013; ; Lee et al. 2002, a).

Four different yeast strains were screened for erythritol production from xylose (Guo et al. 2016a). Aureobasidium pullulans CGMCC3.0837 showed the highest erythritol production (11.5 g/L) and the most efficient xylose consumption among the screened strains (Table 2) (Guo et al. 2016a). A. pullulans CGMCC3.0837 was subjected to UV mutagenesis, and erythritol production increased by 51% in the A. pullulans mutant ER35. Furthermore, response surface methods were used for medium composition optimization to increase erythritol synthesis by A. pullulans ER35. The erythritol production respectively increased 2.6-fold (29.6 g/L) and 2.7-fold (31.8 g/L) as compared to the parent strain during growth in a shake flask and a fermentor (Table 2).

Filamentous fungi produce very low concentrations of erythritol during aerobic growth on xylose (Jovanović et al. 2014). The err1 gene encoding erythrose reductase from Trichoderma reesei has been overexpressed in T. reesei QM6a and T. reesei Rut-C30 (Jovanović et al. 2014). The overexpression strains QPEC1 and RPEC1 produced 3.2-fold and 1.4-fold higher erythritol as compared to wild-type T. reesei QM6a and T. reesei Rut-C30, respectively. Erythritol production was less than 5 mg/L, which is very low as compared to bacteria and yeasts (Table 2).

2,3-Butanediol production

2,3-Butanediol (2,3-BDO) can be used to produce cosmetics, foods, fumigants, antifreeze agent, transport fuels, medicines, and polymers (Celińska and Grajek 2009). Wild-type S. cerevisiae has an endogenous 2,3-BDO biosynthetic pathway (Kim and Hahn 2015; Kim et al. 2017a). Pyruvate is first converted into α-acetolactate by α-acetolactate synthase (ALS) in the mitochondria. The α-acetolactate then spontaneously decaboxylates into diacetyl in the presence of oxygen. The diacetyl is reduced to 2,3-BDO via acetoin by 2,3-butanediol dehydrogenase (BDH) (Fig. 2) (Kim and Hahn 2015; Kim et al. 2013, 2017a, b). In comparison to the bacterial 2,3-BDO pathway, S. cerevisiae does not possess the α-acetolactate decarboxylase (ALDC), which is responsible for direct production of acetoin from α-acetolactate. Native acetoin production is very low in S. cerevisiae, and pyruvate is preferentially used for ethanol synthesis instead of 2,3-BDO. Therefore, S. cerevisiae produces a trace amount of 2,3-BDO.

Fig. 2
figure 2

Xylose pathway engineering for biofuel and chemical production. XR, D-xylose reductase; XDH, xylitol dehydrogenase; XI, D-xylose isomerase; ADH, D-arabitol dehydrogenase; XK, xylulokinase; RPE, ribulose-5-phosphate epimerase; PRK, phosphoribulokinase; RuBisCO, ribulose-1,5-bisphosphate carboxylase/oxygenase; PDC, pyruvate decarboxylase; ALD, alcohol dehydrogenase; ADH, aldehyde dehydrogenase; ACS, acetyl-CoA synthetase; LDH, lactate dehydrogenase; ALS, acetolactate synthase; ALDC, acetolactate decarboxylase; BDH, 2,3-butanediol dehydrogenase; KARI, ketoacid reductoisomerase; DHAD, dihydroxyacid dehydratase; KIVD, α-ketoisovalerate decarboxylase, Adh, alcohol dehydrogenase, ACC, acetyl-CoA carboxylase; FAS, fatty acid synthase; FAR, fatty acyl-CoA reductase FAR; ACAT, acetyl-CoA C-acetyltransferase; HMGCS, hydroxymethylglutaryl-CoA synthase; HMGR, hydroxymethylglutaryl-CoA reductase; MVK, mevalonate kinase; PMVK, phosphomevalonate kinase; MVD, diphosphomevalonate decarboxylase; IIPI, isopentenyl-diphosphate delta-isomerase; FDPS, farnesyl diphosphate synthase; SQS, squalene synthase; ADS, amorphadiene synthase

Full size image

Pyruvate is also a precursor for ethanol synthesis; hence, it is important to redirect flux from ethanol to 2,3-BDO. The deletion of ADH and PDC genes eliminated ethanol production and increased 2,3-BDO production (Kim et al. 2017b). The ALS from B. subtilis has higher affinity for pyruvate than the endogenous ALS encoded by IIV2 in yeast. Acetolactate decarboxylase (ALSD) from B. subtilis catalyzes α-acetolactate to acetoin (Kim et al. 2013). The overexpression of endogenous BDH, ALS, and ALSD from B. subtilis has increased 2,3-BDO production from xylose in S. cerevisiae. The engineered S. cerevisiae BD4X strain produced 20.7 g/L 2,3-BDO with yield 0.27 g/g xylose. The BD4X strain produced 43.6 g/L 2,3-BDO during fed-batch fermentation (Kim et al. 2017b). In addition, overexpressing transaldolase (TAL), xylose reductase, and an NADH oxidase in a PDC-deficient yeast strain enhanced the yield of 2,3-BDO. The engineered BD5X-TXmNP strain produced 69.2 g/L 2,3-BD in batch fermentation and 96.8 g/L 2,3-BDO with productivity of 0.58 g/L/h in fed-batch fermentation (Table 3) (Kim et al. 2017b). Xylose appears to be a better substrate than glucose to produce 2,3-BDO, because the 2,3-BDO pathway is regulated by glucose repression. The deregulation of glucose-dependent repression redirects the metabolic flow from ethanol to 2,3-BD (Alff-Tuomala et al. 2016; Kwak and Jin 2017).

Table 3 List of nonethanol products and chemical production from xylose by engineered S. cerevisiae
Full size table

Isobutanol production

Isobutanol is a branched-chain alcohol that can be used to make transportation fuels, and its derivatives are used in tires, plastic bottles, carpets, and clothing (Felpeto-Santero et al. 2015). Isobutanol is naturally produced in low quantities by S. cerevisiae as a degradation product of valine (Chen et al. 2011).

Overexpression of the genes involved in valine metabolism has been used to increase the isobutanol production in S. cerevisiae (Chen et al. 2011). Mitochondrial valine biosynthesis enzymes such as ALS, keto-acid reductoisomerase (KARI), and dihydroxyacid dehydratase (DHAD) have been truncated and expressed in the cytosol to increase isobutanol production. The cytosolic isobutanol pathway was overexpressed in S. cerevisiae, which was previously engineered for xylose consumption. The resulting S. cerevisiae strain utilized 12 g/L xylose and produced 1.36 mg/L isobutanol with a yield of 0.16 mg/g xylose (Table 3) (Brat and Boles 2013). The consumption of nonfermentable sugars deregulates glucose-dependent repression on the development of mitochondria. The reverse localization of cytosolic pathways into mitochondria would be a better strategy to increase isobutanol production from xylose (DeRisi et al. 1997; Egner et al. 2002). Optimization of the xylose metabolic pathway and eliminating the competing pathways could enhance isobutanol production from xylose (Brat and Boles 2013).

Citric acid production

Citric acid is a widely used organic acid in the food industry (Berovic and Legisa 2007). For example, citric acid is used in carbonated beverages to provide taste and complement fruit flavors. The chelating and pH-adjusting properties of citric acid have been used to increase the stability of frozen food products by enhancing the activity of antioxidants. It is also used to inhibit color and flavor deterioration in frozen fruits. In addition, citric acid is used in oral pharmaceutical liquids and to maintain the activity of preservatives (Berovic and Legisa 2007).

The filamentous fungus Aspergillus niger has been used for decades to produce citric acid (Berovic and Legisa 2007). Traditionally, citric acid is produced from starch or sucrose. However, A. niger can also naturally produce citric acid from xylose. After 14 days of fermentation, A. niger produced 22 g/L citric acid from 71 g/L xylose with a yield of 0.31 g/g (Maddox et al. 1985). A. niger iMA871 produced 22 g/L citric acid from 160 g/L xylose within 8 days, and production was commenced upon a switch to phosphate-limited growth (Upton et al. 2017). The wild-type Aspergillus carbonarius produced citric acid at yield of 3 g/g dry cell mass and the engineered strain A. carbonarius SXR1, in which an NADH-dependent xylose reductase was expressed, produced citric acid at 13.8 g/g dry cell mass (Weyda et al. 2014).

The oleaginous yeast Yarrowia lipolytica naturally produces high concentrations of citric acid during growth on glucose. However, it does not grow well on xylose (Ryu et al. 2015). Y. lipolytica strain Po1g was engineered and adapted to grow on xylose by overexpressing XR and XDH from P. stipitis (Ledesma-Amaro et al. 2016; Ryu et al. 2015). This strain efficiently produced 80 g/L citric acid with a yield 0.53 g/g xylose (Ledesma-Amaro et al. 2016). Moreover, the production titer was similar to previously reported citric acid overproducing strains growing in glucose and glycerol (Lazar et al. 2011; Rywińska et al. 2010).

Lactic acid production

Lactate has a wide range of applications in the production of polylactic acid, biodegradable plastics, textile fibers, and additives in cosmetic and pharmaceutical industries (Yang et al. 2013). Lactic acid bacteria are generally used for lactate production (Papagianni 2012). The low pH during lactic acid fermentation, however, is inhibitory for bacteria, whereas S. cerevisiae is more tolerant to acidic pH.

L-lactate dehydrogenase (LDH) has been overexpressed in yeasts for the production of lactic acid from pyruvate (Dequin and Barre 1994; Porro et al. 1995). The LDH reaction is reversible, and LDH is allosterically inhibited by lactate (Papagianni 2012). Pyruvate is the branch metabolite between lactate and ethanol pathways. LDH has higher affinity for pyruvate and it competes with PDC for pyruvate. LDH could direct more metabolic flux to lactate from pyruvate than PDC because of weak glycolytic metabolism on xylose.

In ethanol-fermenting yeasts, pyruvate is converted into ethanol via acetaldehyde by PDC and alcohol dehydrogenase (ADH), respectively. Ethanol is still a major by-product produced with lactate in engineered S. cerevisiae. The deletion of aldehyde dehydrogenase (ALDH) and PDC reduced the metabolic flux toward ethanol and increased metabolic flux toward lactate (Baek et al. 2016).

The expression of LDH in xylose-fermenting S. cerevisiae produced lactic acid 60 g/L with a yield of 0.67 g/g xylose with low ethanol accumulation (0.01 g/g xylose) (Table 3) (Turner et al. 2015, 2017). The expression of lactate transporter gene JEN1 in S. cerevisiae has increased lactate transport outside the cell and led to increased lactate productivity (Branduardi et al. 2006).

LDH from Lactobacillus helveticus has been expressed in P. stipitis and the resulting strain produced 58 g/L lactate with a yield of 0.58 g/g xylose. This engineered strain cultured on xylose was better for lactate production than the best reported S. cerevisiae strains on glucose (Colombié et al. 2003; Ishida et al. 2005; Skory 2003).

Isoprenoid production

Isoprenoids are a class of natural products and main constituents of essential oils from plants and flowers (Chang and Keasling 2006). Extraction methods of isoprenoids from natural sources are inefficient and require a large quantity of natural resources (Nevoigt 2008). The isoprenoid squalene is linked to reduce serum cholesterol levels and is used in cosmetic, pharmaceutical, and food industries (Zhang et al. 2002). The isoprenoid amorphadiene is a plant derived isoprenoid and a precursor of antimalarial drug artemisinin (Chang and Keasling 2006). Metabolic engineering approaches have been used for the overproduction of the isoprenoids squalene and amorphadiene from xylose by S. cerevisiae. Xylose is preferred over glucose because of increased oxidative metabolism and reduced fermentative metabolism (Alff-Tuomala et al. 2016).

The pyruvate dehydrogenase bypass in S. cerevisiae has been engineered by overexpressing ALDH and ACS from Salmonella enterica to increase the supply of acetyl-CoA to the mevalonate pathway resulting in enhanced amorphadiene production (Shiba et al. 2007). The overexpression of truncated HMG-CoA reductase isozyme 1 (tHMG1) in an engineered S. cerevisiae capable of xylose utilization has also been an effective strategy for isoprenoid overproduction. TAL1, acetyl-CoA C-acetyltransferase (ACAT), and hydroxymethylglutaryl-CoA reductase (HMGR) were overexpressed using constitutive promoters in S. cerevisiae to increase xylose consumption and increase the metabolic fluxes from acetyl-CoA to mevalonate (MEV) pathway (Fig. 2). The co-expression of truncated ACAT and HMGR in S. cerevisiae has produced higher squalene from xylose than glucose, because of ethanol reassimilation with xylose utilization and metabolic flux transfer to cytosolic acetyl-CoA synthesis. Finally, 532 mg/L squalene was produced with a productivity of 11.3 mg/L/h from xylose in engineered S. cerevisiae (Table 3) (Kwak et al. 2017). Squalene synthase (SQS) downregulation and overexpression of ACAT, HMGR, and amorphadiene synthase (ADS) further increased amorphadiene titers. The HX4HEA-ES strain produced 254 mg/L amorphadiene under xylose culture conditions (Table 3) (Kwak et al. 2017).

Carotenoid production

Carotenoids are naturally occurring lipid soluble pigments displaying yellow, orange, and red color synthesized via the mevalonate pathway. Phytoene acts a precursor for the biosynthesis of specific carotenoids such as β-carotene, torulene, astaxanthin, and torularhodin in pigmented yeasts (Moliné et al. 2012). These pigments act as vitamin A precursors and have coloring and antioxidant properties. Rhodotorula spp. can synthesize carotenoids such as β-carotene, torulene, and torularhodin (Liu et al. 2018; Moliné et al. 2012).

The carotenogenic genes from Erwinia uredovora, Agrobacterium aurantiacum, and Xanthophyllomyces dendrorhus have been introduced into S. cerevisiae, Candida utilis, and Y. lipolytica for the production of carotenoids such as β-carotene, lycopene, or astaxanthin (Lange and Steinbuchel 2011; Matthaus et al. 2014; Misawa and Shimada 1997). The red yeast Phaffia rhodozyma NRRL Y-17268 has produced 5.2 mg/L astaxanthin from xylose-grown media (Parajo et al. 1997; Parajó et al. 1998).

Lipid production

Microbial lipid production has several advantages over plant oils, such as faster growth in defined media, lower land usage, better adaptability to market demands, and the ability to be produced from multiple substrates (Park et al. 2017). These lipids can be used to produce fuels, lubricants, and surfactants (Beopoulos et al. 2014; Blazeck et al. 2014; Matthaus et al. 2014; Ratledge 2004; Xue et al. 2013). Multiple microbes can accumulate large amounts of lipids, ranging from 20 to 70% of dry cell weight, during growth on many different sugars (Díaz et al. 2018; Huang et al. 2013; Kosa and Ragauskas 2011; Kurosawa et al. 2013; Shi and Zhao 2017; Yaguchi et al. 2018; Zhang et al. 2016a). Typically, these microbes accumulate lipid when cultured in excess sugar relative to other essential nutrients such nitrogen or phosphorous. As they cannot use the sugar for growth, they instead store it as lipid (Wang et al. 2018).

The theoretical yield of lipid production from xylose is 0.34 g/g, which is higher than glucose 0.32 g/g (Papanikolaou and Aggelis 2011). However, the practical yield of lipid is lower (Jin et al. 2015; Li et al. 2008; Weete 2012). Multiple strategies have been used to enhance lipid production such as overexpressing native or heterologous genes involved in sugar uptake and assimilation, overexpressing genes of fatty acid biosynthesis and TAG pathway, increasing availability of precursors for lipid synthesis, and deleting lipolysis pathway and β-oxidation pathway (Ferreira et al. 2018; Ledesma-Amaro et al. 2016; Li and Alper 2016; Niehus et al. 2018; Xu et al. 2016; Zhang et al. 2016a).

Bacteria

A few bacteria are able to produce high quantities of lipids during growth on xylose (Kurosawa et al. 2013; Xiong et al. 2012). For example, the actinomycetes genera of Rhodococcus, Streptomyces, and Gordonia accumulate lipids up to 20–30% of dry cell weight during growth on xylose (Castro et al. 2016; Waltermann and Steinbuchel 2005).

Bacteria such as Rhodococcus opacus PD630 and Rhodococcus jostii RHA1 can efficiently convert glucose into lipids but are unable to consume xylose naturally (Alvarez 2006; Hernández et al. 2008). XylA encoding xylose isomerase and xylB encoding xylulokinase from Streptomyces lividans TK23 were overexpressed into both R. opacus PD630 and R. jostii RHA1. Under nitrogen-limiting conditions, R. opacus PD630 XYLB and R. jostii RHA1 XYLB strains produced lipid up to 52.5 and 68.3% of dry cell weight, respectively. In another study, the xylose metabolic pathway genes from Streptomyces padanus MITKK-103 were expressed in R. opacus PD630 (Kurosawa et al. 2013). The resulting strain has produced 12 g/L lipid during growth on 120 g/L xylose.

Fungi

Oleaginous fungi can accumulate lipids up to 70% of dry cell weight (Chatzifragkou et al. 2010; Dey et al. 2011; Fakas et al. 2009; Mamatha et al. 2008; Ruan et al. 2012; Zheng et al. 2012). In one study, 11 filamentous fungi were investigated for lipid production during growth on xylose (Zheng et al. 2012) (Table 4). The lipid production ranged from 0.15 to 3.12 g/L, and lipid content ranged from 4 to 51% of dry cell weight (Gao et al. 2013; Zheng et al. 2012). The highest 50.9% lipid content was achieved by Mortierella isabellina during growth on xylose (Zheng et al. 2012). M. isabellina produced 18.5 g/L lipids with 64.3% lipid content during growth on 100 g/L xylose (Gao et al. 2013).

Table 4 List of lipid production from xylose by wild-type and engineered microorganisms
Full size table

Yeast

Oleaginous yeasts such as R. toruloides, Lipomyces starkeyi, Cryptococcus curvatus, and Cutaneotrichosporon oleaginosus can accumulate intracellular lipids in the form of triacylglycerol (TAGs) as high as 40–60% of their cell dry weight from xylose under nitrogen-limited condition (Díaz et al. 2018; Shi and Zhao 2017; Yaguchi et al. 2018; Zhang et al. 2016b). R. toruloides IFO0880 produced 7.1 g/L lipid from xylose in low nitrogen medium (Zhang et al. 2016b). L. starkeyi AS 2.1560 and R. toruloides AS 2.138 produced 7.4 g/L and 5.3 g/L lipid, respectively, from pure xylose solution without any other nutrients in a two-stage fermentation process (Table 4) (Lin et al. 2014).

In R. toruloides strains, UV mutagenesis combined with a selection using cerulenin was used to increase lipid productivity (Yamada et al. 2017). A high-lipid-producing (1.86 g/L lipid) mutant of R. toruloides 87663-11C 2-53C was selected, and transcriptional levels of genes related to xylose utilization and lipid production were profiled (Yamada et al. 2017). The transcription levels of three (XYL1, XYL2, and XKS1) xylose metabolic genes in the mutant were the same as parental strain. Meanwhile, the transcription six genes (ACL1, FAS1, FAS2, GDH1, MAE1, and PYC1) related to lipid production in mutant strain were 4-fold upregulated (Yamada et al. 2017). In another study, overexpressing the native ACC1 and diacylglycerol acyltransferase (DGA1) enzymes involved in lipid production in R. toruloides IFO0880 increased lipid production from 7.1 to 9.5 g/L from xylose (Table 4) (Zhang et al. 2016a).

The overexpression of the native XDH, XR, and XK genes in Y. lipolytica allowed for growth on xylose as a sole carbon source for lipid production (Ledesma-Amaro et al. 2016; Li and Alper 2016; Niehus et al. 2018; Rodriguez et al. 2016). The resulting ylXYL+ strain produced 2.35 g/L lipid during growth on 60 g/L xylose (Table 4) (Niehus et al. 2018). The ylXYL+ strain was further engineered to obtain ylXYL-Obese-XA strain by overexpressing and deleting a number of genes involved in lipid production. The resulting ylXYL-Obese-XA strain produced 16.5 g/L lipid, 8.3 times higher than wild type in fed-batch fermentation. The maximum lipid content and productivity were 67% and 1.85 g/L/h, respectively.

Fatty alcohol production

Fatty alcohols are used in the production of cosmetics, detergents, lubricants, esters, and emulsifiers (Feng et al. 2015). Fatty alcohols can be extracted from natural plant oils or chemically synthesized from petrochemical sources. Both methods have limitations due to environmental concern or competition with food supply (Rupilius and Ahmad 2007). Microbial production of fatty alcohols from xylose may provide a better alternative than traditional methods.

As compared to glucose, xylose metabolism in engineered strains of S. cerevisiae appears to be more amendable for fatty alcohol production. The cytosolic acetyl-CoA is more efficiently produced under xylose compared to glucose (Jin et al. 2004; Salusjärvi et al. 2008). Many genes encoding the TCA cycle and respiratory enzymes in mitochondria are upregulated during growth on xylose and enhance ATP and citrate production. As a consequence, acetyl-CoA generation is enhanced by the action of ACS and ATP citrate lyase (CSL) (Jin et al. 2004; Salusjärvi et al. 2008). In addition, xylose culture conditions prevent the inhibition of fatty acid synthesis and deregulation of acyl-CoA because xylose does not induce SNF1, a protein that downregulates the expression of acetyl-CoA carboxylase (ACC1) and upregulates β-oxidation (Brink et al. 2016; Feng et al. 2015; Lee et al. 2015).

The overexpression of structural genes and deletion of negative regulators involved in downregulation of lipid synthesis in yeasts have been demonstrated to increase acetyl-CoA availability to improve 1-hexadecanol production. Two families of transcriptional factors, AMP-activated serine/threonine protein kinase (SNF1) and regulators of inositol-3-phosphate synthase (INO1), have been knocked-out to improve fatty acid synthesis in yeasts (Seip et al. 2013; Shi et al. 2014).

The overexpression of fatty acyl-CoA reductase (FAR), ACC1, ACL, and deletion of this histone deacetylase RPD3 improved 1-hexadecanol production from 45 mg/L to 1.1 g/L through fed-batch fermentation of glucose (Feng et al. 2015). Furthermore, when a xylose utilization pathway was introduced into the engineered S. cerevisiae strain, it produced 0.4 g/L 1-hexadecanol from xylose. Promoter engineering of xylose pathway increased 1-hexadecanol titer by 1.7-fold. The evolved strain produced over 1.2 g/L 1-hexadecanol using xylose during fed-batch fermentation (Table 3) (Guo et al. 2016b).

Conclusions

In this review, we discussed how different microorganisms are being used to produce diverse chemicals and fuels from xylose, an abundant sugar derived from plant biomass. Some of these products, such as sugar alcohols, are directly linked to xylose metabolism and, as such, represent ideal products derived from xylose. Others, such as lipid-based chemicals, are derived from core metabolic intermediates. The viability of making these products from xylose depends on the efficiency of xylose utilization, which is often less efficient than glucose utilization. In these regards, multiple opportunities exist for increasing the efficiency of xylose utilization such as the discovery and engineering of improved transporters and enzymes involved in xylose utilization. In addition, engineering microorganisms capable of simultaneously utilizing multiple sugars found in plant biomass would greatly increase the potential of plant biomass as a renewable feedstock for the production of diverse chemicals and fuels. While different strategies to address these challenges were not discussed in this review due to space limitations, they nonetheless represent a fertile area of active and future research.

We conclude by noting that xylose, despite limitations in utilization efficiency, may be a better feedstock than glucose, particularly in the case of yeast. Many pathways associated with nonfermentative metabolism are repressed during growth on glucose but not on xylose in engineered yeast. This means that it is often easier to redirect metabolic flux away from ethanol and toward other products during growth on xylose than it is on glucose (Alff-Tuomala et al. 2016; Kwak and Jin 2017). Thus, many nonethanol products are easier to produce from xylose than from glucose (Alff-Tuomala et al. 2016; Brat et al. 2009; Feng et al. 2015; Jin et al. 2004; Kim et al. 2017b; Kwak et al. 2017; Niehus et al. 2018; Salusjärvi et al. 2008; Shiba et al. 2007). In these regards, xylose may represent the ideal sugar for producing novel bioproducts from renewable biomass.