**The Xylose Metabolizing Yeast** *Spathaspora passalidarum* **is a Promising Genetic Treasure for Improving Bioethanol Production**

#### **Khaled A. Selim 1,\*, Saadia M. Easa <sup>2</sup> and Ahmed I. El-Diwany <sup>1</sup>**


Received: 15 February 2020; Accepted: 16 March 2020; Published: 18 March 2020

**Abstract:** Currently, the fermentation technology for recycling agriculture waste for generation of alternative renewable biofuels is getting more and more attention because of the environmental merits of biofuels for decreasing the rapid rise of greenhouse gas effects compared to petrochemical, keeping in mind the increase of petrol cost and the exhaustion of limited petroleum resources. One of widely used biofuels is bioethanol, and the use of yeasts for commercial fermentation of cellulosic and hemicellulosic agricultural biomasses is one of the growing biotechnological trends for bioethanol production. Effective fermentation and assimilation of xylose, the major pentose sugar element of plant cell walls and the second most abundant carbohydrate, is a bottleneck step towards a robust biofuel production from agricultural waste materials. Hence, several attempts were implemented to engineer the conventional *Saccharomyces cerevisiae* yeast to transport and ferment xylose because naturally it does not use xylose, using genetic materials of *Pichia stipitis*, the pioneer native xylose fermenting yeast. Recently, the nonconventional yeast *Spathaspora passalidarum* appeared as a founder member of a new small group of yeasts that, like *Pichia stipitis*, can utilize and ferment xylose. Therefore, the understanding of the molecular mechanisms regulating the xylose assimilation in such pentose fermenting yeasts will enable us to eliminate the obstacles in the biofuels pipeline, and to develop industrial strains by means of genetic engineering to increase the availability of renewable biofuel products from agricultural biomass. In this review, we will highlight the recent advances in the field of native xylose metabolizing yeasts, with special emphasis on *S. passalidarum* for improving bioethanol production.

**Keywords:** fermentation; xylose metabolism; genetic engineering; biofuel; *Spathaspora passalidarum*; *Pichia stipitis*

#### **1. Fermentation Technology and Challenges**

The modern biotechnological applications for generation of alternative and renewable sources of biofuels are receiving more attention due to global worries over the climate change, rapid global warming, and the rising of fossil fuel costs. One of such growing biotechnological trends is the fermentation technology to convert the sugar-rich agriculture waste into bioethanol by conventional or non-conventional yeasts [1–4]. In general, yeasts have advantages over bacteria for commercial fermentation due to the thickness of their cell walls, less stringent nutritional requirements, large sizes, utmost resistance to contamination, and better growth at acidic pH of bioreactor fermenters.

In nature, the second most abundant hemicellulosic sugar in fast-growing hardwoods and agricultural biomass is xylose. Xylose sugar forms up to 15–25% of all angiosperm biomass, and it could supply an alternative fuel source for its ability to be commercially fermented into ethanol. Several approaches have been employed to engineer xylose assimilation metabolism into conventional fermenting yeasts, such as *Saccharomyces cerevisiae*[4–6]. Therefore, efficient hemicellulosic sugar fermentation is crucial for the economic conversion of lignocellulose biomass to renewable biofuels [4,6–8]. The discovery of xylose-fermenting yeasts in new niches and genetic engineering of yeasts to be capable of rapid fermentation of xylose and other sugars to recoverable concentrations of bioethanol could provide alternative biofuel sources for the future (Figure 1) [4,9].

**Figure 1.** Model of yeast fermentation machinery for bioethanol production using the agriculture waste to feed yeasts [4]. The metabolic pathways for xylose and glucose assimilation and fermentation are indicated including the pentose phosphate pathway and glycolysis. The agriculture waste is treated through the enzymatic and chemical simultaneous saccharification and fermentation (SSF) processes to release the cellulosic and hemicellulosic sugars. The hexose and pentose sugars are transported by specific hexose and pentose sugar transporters into yeast for further metabolizing processes.

As a rule of thumb for metabolizing the xylose in most of xylose-fermenting yeasts [4], firstly the xylose is reduced by xylose reductase (XR) to xylitol. In the second step, the xylitol is oxidized by xylitol dehydrogenase (XDH) to xylulose. Afterward, the xylulose passes into the pentose phosphate pathway being metabolized into glyceraldehyde-3-P which is further reduced to pyruvate. Finally, the pyruvate is decarboxylated to acetaldehyde which is further reduced to ethanol by alcohol dehydrogenase (Figure 1). Notably, most xylose reductase enzymes have dual cofactor specificity, using both NADH and NADPH, but typically favor NADPH. However, xylitol dehydrogenase enzymes use NAD<sup>+</sup> specifically as a cofactor, which could cause imbalance between the cofactor's source for the XR-XDH pathway and xylitol accumulation under uncontrolled oxygen conditions (Figure 2) [10].

**Figure 2.** Schematic model of the central metabolism for bioethanol production from xylose indicating the rate limiting steps (XR: xylose reductase; XDH: xylitol dehydrogenase, and ADH: alcohol dehydrogenase) and cofactors demand/balance in most of native xylose metabolizing yeasts. Xylose assimilation reactions starts with XR to produce xylitol. The xylitol is further metabolized by XDH to produce xylulose, which further metabolized to xylulose-phosphate to enter the glycolysis (indicate by black dotted arrow and summarized in Figure 1) to produce acetaldehyde. The acetaldehyde finally converted to ethanol by ADH.

One of pioneer xylose fermenting yeasts is *Pichia stipites*. *P. stipitis* is heterothallic ascomycetous yeast, predominantly haploid and related to pentose fermenting yeasts, such as *Candida shehatae* [1,4,11–15]. *P. stipitis* was recently renamed to be *Sche*ff*ersomyces stipitis* [15] and it is natively one of the highest xylose-utilizing and fermenting yeasts. In type culture collections, the *P. stipitis* strains are among the best xylose-metabolizing microbes [16]. Under controlled low O2 conditions, *P. stipitis* is able to consume xylose and produce up to 57 g/L of bioethanol at 30◦C [4,13,14,17]. *Pichia* uses an alternative nuclear genetic code (ANGC) in which CUG encodes for Ser rather than Leu [17], which makes the genetic manipulation of *Pichia* with the commercial drug resistance markers unusually problematic because essentially all of these markers are derived from bacteria that use the universal codon system. Moreover, one of classical challenges in fermentation technology is that some of key enzymes of bioethanol production pathway are expressed relatively in low levels [1,4,13,14]. Therefore, the metabolic engineering of the bioethanol pathway in yeasts, which can ferment the sugars of the agriculture biomass with considerable and recoverable bioethanol concentrations, could enhance the productivity and sustainability of renewable biofuel sources [1,13,14]. To improve bioethanol production and xylose metabolism, a stable and manipulatable genetic system that enables overexpression or deletion of one or more of key enzymes and sugar transporters in xylose-fermenting yeast *P. stipitis*, was developed [1,3,18]. This approach comprises modelling, metabolic and flux analysis, quantitative metabolomics and transcriptomics followed by the targeted overexpression or deletion of genes of the rate-limiting steps [1,3,4,13,14]. Since there is reasonable information about the metabolic capacity of *P. stipitis* to ferment xylose on various omics levels, this makes it an attractive model system for metabolic engineering.

Recently, a new xylose-fermenting yeast *Spathaspora passalidarum* was discovered, which naturally co-ferments xylose, glucose, and cellobiose and demonstrates potentials in the effective conversion of mixed sugars from hemicellulosic hydrolysates into ethanol [9]. *S. passalidarum* was initially isolated from extremely O2-limited and hemicellulosic sugar rich environments from the gut of a wood-boring beetle (as will be discussed below). Although the anaerobic fermentation of glucose is broadly known, the xylose fermentation typically needs a controlled oxygen condition. Uncontrolled oxygen conditions in another xylose-fermenting yeast, such as *P. stipitis*, leads to accumulation of xylitol due to insufficient amounts of NAD+, and as consequence the xylose metabolism will be blocked (Figure 2) [9,13,14]. To solve this problem, precise controlled O2 (very low O2 concentrations) during the xylose fermentation is required in *P. stipitis* to generate NAD<sup>+</sup> from NADH.

The bioethanol production from the bioconversion of lignocellulose biomass must be achieved at high rates and yields for economically recoverable concentrations. The achieving of such targets for efficient bioethanol production are more difficult with cellulose and hemicellulose. The major barrier for cellulose utilization is enzymatic saccharification, while for hemicellulose it is the utilization of mixed sugars (hexose sugars: glucose, galactose, mannose, and rhamnose; and pentose sugars: xylose and arabinose) in the presence of ferulic and acetic acids along with other byproducts of the thermochemical pretreatment of the hydrolysates [13,14]. However, *S. passalidarum* and *P. stipitis* yeasts possess a set of unique physiological merits that make them very useful biodegradable organisms for bioconversion of lignocellulosic biomass [4,9,13,14]. *Pichia* can utilize and ferment effectively cellobiose, glucose, galactose, and mannose along with xylan high oligomeric sugars xylan and mannan, in addition to its extensively studied ability to metabolize and ferment the xylose [4,13,14]. The primary sugar released in enzymatic hydrolysis is cellobiose and, remarkably, *P. stipitis* and *S. passalidarum* have the capability to utilize the cellobiose, which make such yeasts potent organisms for simultaneous saccharification and fermentation (SSF) or hydrolysate, because most commercially available cellulase products are often deficient in β-glucosidase enzyme so the accumulation of cellobiose inhibits cellulose activities. Since *P. stipitis* and *S. passalidarum* can directly metabolize the cellobiose, they have the potential to improve SSF processes [4,9,10,13,14,19,20].

Collectively, the native ability of *P. stipitis* and *S. passalidarum* to metabolize the oligomeric sugars is of high importance as the mild acidic pretreatments of agriculture waste biomass can prevent the formation of the sugar degradation byproducts, which could inhibit significantly the fermentation process, but can release about 15–55% of soluble oligomeric sugars. Therefore, with low cost and high yield, the hemicellulosic sugars can be more readily recovered and underutilized from cellulose biomass than glucose. Although such easy recoverable sugars can be utilized for formation of a number of useful products such as xylitol, butanol, lactic acid, and other chemicals, bioethanol is still the major product with the largest potential market. Hence, bioethanol production from the lignocellulosic biomasses is receiving a lot of attention as a consequence of agriculture policies and energy demands to improve the production of alternative renewable biofuels and to reduce CO2 emissions [2,4,13,14].

#### **2.** *Spathaspora passalidarum* **a Promising Genetic Source**

The Spathaspora clade contains many bioethanol producer yeasts, including *Spathaspora arborariae*, *Spathaspora brasiliensis*, *Spathaspora gorwiae*, *Spathaspora hagerdaliae*, *Spathaspora passalidarum*, *Spathaspora roraimanensis*, *Spathaspora suhii*, *Spathaspora xylofermentans*. They are usually endosymbioticly associated with wood-boring-beetles that occupy rotting wood. *Spathaspora passalidarum* (Figure 3), the first identified species of genus Spathaspora, was isolated from the gut of passalid beetle *Odontotaenius disjunctus* [9,21–24]. Notably, *S. arborariae*, *S. gorwiae*, *S. hagerdaliae*, and *S. passalidarum* ferment xylose to produce bioethanol, whereas the rest within the Spathaspora clade are thought to be xylitol producers [9].

**Figure 3.** *Spathaspora passalidarum* budding cells with characteristic curved and elongated ascospore.

*S. passalidarum* was firstly described in 2006 by Nguyen et al. [21]. The authors speculated that *Spathaspora* mainly exists in the beetle's biosphere rather than the beetle's gut microbiota, and it may be only by coincidence that *O. disjunctus* beetles ingested decaying wood contaminated by yeasts. Later in 2012 and 2017, another 12 strains were described in two independent studies from wood-boring beetles and wood samples of Amazonian forest in Brazil [23,25]. Among these strains, only one isolate was obtained from the gut of *Popilus marginatus* beetle, while the rest of the strains were obtained from the woody samples inhabited by the beetles [23,25]. In 2014, two more strains were isolated from rotted wood in China [26]. Additionally, Rodrussamee and colleges in 2018 reported a new thermotolerant strain, named *S. passalidarum* CMUWF1–2, which was isolated from Thailand soil [27]. The frequency of finding *S. passalidarum* mainly among the woody samples supports the notion that those yeasts are probably associated with decaying wood niches rather than with the gut microbiota of wood-boring beetles. However, the fact of the low frequency of finding *S. passalidarum* among other yeast species keeps an open possibility that they inhabit mainly the wood-related beetles [9].

#### **3. Fermentation Capability of** *Spathaspora passalidarum*

It is believed that the beetle's gut is truly anaerobic or microaerobic, therefore it was speculated that *S. passalidarum* possess a unique adaptation capability to survive under oxygen-depleted conditions on mixtures of hemicellulosic sugars in the midgut of wood-boring beetles [10,19]. Currently, *S. passalidarum* is among the best xylose-utilizing and fermenting yeasts. Under anaerobic or microaerobic conditions, *S. passalidarum* possess rapid utilization and consumption rates for xylose and produces up to 0.48 g/g bioethanol (near to the maximum theoretical bioethanol production of 0.51 g/g), in contrast to *P. stipitis* which can hardly metabolize xylose anaerobically, accumulating xylitol and a very low yield of bioethanol [10,19,20,28,29]. Under anaerobic conditions, Hou in 2012 showed that *S. passalidarum* has a high growth rate with rapid consumption rate of sugars and can ferment xylose into a high yield of bioethanol with higher production efficiency than *P. stipitis* [10]. Similarly, Veras and colleges in 2017 showed that under anaerobic conditions, *S. passalidarum* accumulates 1.5 times more bioethanol than *S. stipitis*, while both stains accumulate around 0.44 g/g under O2 limiting conditions [30]. The previous work by Hou (2012) defined strictly that *S. passalidarum* can metabolize and ferment xylose in tightly capped flasks [10]. In contrast to the previous report by Hou (2012) [10], under stringent O2 limiting conditions, the *S. passalidarum* was not able virtually to utilize the sugars, indicating that native wild-type *S. passalidarum* does not ferment sugars under truly anaerobic conditions [19,20]. Therefore, it is still under debate whether *S. passalidarum* can ferment xylose truly anaerobically or whether it requires a controlled microoxygenic condition similar to *P. stipitis*.

One of the major challenges in fermentation technology is the inability of the majority of known microbes to co-ferment xylose and glucose, since glucose usually inhibits the metabolization of the other sugars in lignocellulose hydrolysate, as in the case of *P. stipitis* [13,14]. Astonishingly, in a recent study to address the metabolic profiling and fermentation capacity of *S. passalidarum*, *S. passalidarum* was found to co-ferment xylose, cellobiose, and glucose simultaneously with high bioethanol yields ranging from 0.31 to 0.42 g/g [19,20]. Moreover, an adapted *S. passalidarum* strain was found to accumulate up to 39 g/L bioethanol with a 0.37 g/g yield from a lignocellulosic hydrolysate. The specific production rate of bioethanol on xylose as a carbon source was superior with three times more than the corresponding rate on glucose, where the flux of glycolytic intermediates was meaningfully lower on glucose than on xylose and its xylose reductase enzyme had a higher affinity for NADH than NADPH [19,20]. Thus, the allosteric activation of glycolytic routes associated with the xylose utilization and the NADH-dependent xylose reductase are most likely the causes for such unique ability of *S. passalidarum* to co-ferment mixed sugars [19,23]. Later, such results were confirmed in a metabolic flux study, where *S. passalidarum* showed about 1.5–2 times high flux rate in the NADH-dependent xylose reductase reaction [31], which caused continuous recycling and reduction of xylitol levels. Such directed high flux rates to glycolytic routes and pentose phosphate pathway was the cause for high levels of bioethanol production in *S. passalidarum* [31]. In large scale fed-batch fermentation study, *S.* *passalidarum* was able metabolize around 90% of xylose sugar and all of glucose of sugarcane bagasse hydrolysate, even so glucose had approximately three-fold higher xylose content; and produced a high ethanol yield of 0.46 g/g with volumetric productivity of 0.81 g/L/h in contrast to *P. stipitis* which produced 0.32 g/g ethanol with productivity of 0.36 g/L/h [32]. In follow up study, *S. passalidarum* UFMG-CM-Y473 strain was able to simultaneously utilize and co-ferment about 78% of the released sugars (xylose, glucose, and cellobiose) of pretreated sugarcane bagasse hydrolysate (delignified and enzymatically hydrolyzed) to yield up to 0.32 g/g bioethanol with productivity of 0.34 g/L/h without any nutritional supplementation [33]. Moreover, the new thermotolerant strain, *S. passalidarum* CMUWF1–2, was able to co-ferment various sugars (mannose, galactose, xylose, and arabinose) of lignocellulosic biomass, even in presence of glucose, to accumulate considerable amounts of bioethanol and low amounts of xylitol at higher temperatures. For example, it was able to accumulate 0.43, 0.40, and 0.20 g/g ethanol per xylose at 30, 37, and 40 ◦C, respectively [27]. Constant with absence of the glucose repression effect on the utilization of other sugars, *S. passalidarum* CMUWF1–2 exhibited a resistance to 2-deoxy glucose, the nonmetabolizable glucose analog, and tolerance to elevated levels of glucose (35.0% of *w*/*v*) and ethanol (8.0% of *v*/*v*) [27]. In contrast, the first discovered *S. passalidarum* NRRL Y-27907 strain was sensitive to 2-deoxy glucose, as 2-deoxy glucose suppressed the xylose consumption under anaerobic conditions. While under aerobic conditions, the 2-deoxy glucose inhibited, only partially, *S. passalidarum* NRRL Y-27907 [10]. Therefore, the author speculated that xylose uptake in *S. passalidarum* NRRL Y-27907 may take place by different xylose transport systems under aerobic and anaerobic conditions. Under aerobic conditions, xylose is taken up by means of ATP-dependent/high affinity xylose-proton symporter and low affinity transporter via facilitated diffusion driven only by the sugar gradient. While, under anaerobic condition, the yeasts are most likely to use only the low affinity xylose pump, as the active transport via xylose-proton symporter will deplete the ATP levels. The inhibitory effect of 2-deoxy glucose on *S. passalidarum* NRRL Y-27907 can be therefore explained by (1) blocking of the low-affinity-facilitated diffusion transporters which are occupied with transporting 2-deoxy glucose, and (2) the inhibition xylose active transport due to the depletion of the intracellular ATP levels to actively phosphorate the 2-deoxy glucose into the non-metabolizable phospho-2-deoxy glucose [10].

#### **4. Genetic and Physiological Features of** *Spathaspora passalidarum* **Emphasis Special Roles for Xylose Reductase and Xylitol Dehydrogenase**

These unusual unique traits of *S. passalidarum* are very attractive for studying on a molecular level. The complete genome sequence of xylose-fermenting yeast *S. passalidarum* was therefore necessary and it was accomplished and published for first time in 2011 [34]. The comparative genomic, transcriptomic, and metabolomic analysis between two of the native xylose-fermenting yeasts, the relatively newly discovered *S. passalidarum*, and the deeply studied *P. stipitis*, allowed a better understanding of the regulatory mechanisms of lignocellulose utilization, and identified the target key genes involved in xylose metabolism [9,10,19,31,34]. The comparative genomic and phylogenetic analysis clearly revealed that *S. passalidarum* is one of the CUG yeast clades, similar to *P. stipites* [34]. In addition, the transcriptome analysis indicated upregulation of the genes implicated in transporting carbohydrate and xylose- and carbohydrate-metabolisms under xylose growth. Several of genes, which are involved in regulation of redox balance and recycling of NAD(P)H/ <sup>+</sup>, were upregulated to probably keep the redox balance during xylose utilization. Additionally, the genes encoding for cellulases and β-glucosidases were also upregulated, which suggests a positive feedback of xylose on the upstream genes to activate its own liberation from the higher oligomeric sugars of hemicelluloses by means of the catalytic activities of cellulases and β-glucosidases [34].

The previously mentioned capabilities of *S. passalidarum* to co-ferment different sugars and accumulate high levels of bioethanol with very low concentrations of xylitol, can be explained by the presence of a set of physiological characters encoded by unique set of genes [10,19,34]. Thus, the high capacity of xylose fermentation and low levels of xylitol accumulation by *S. passalidarum* was speculated to be due to the cofactor's equilibrium between the intracellular demand and supply of the cofactors via NADH-favored xylose reductase enzyme and NAD+-specific xylitol dehydrogenase enzyme, the key enzymes of the xylose utilization pathway [10,28]. Normally, the xylose metabolization occurs through the reduction of xylose to xylitol with xylose reductase, which requires NADPH or NADH as a cofactor with preference for NADPH. Only few NADH-favored xylose reductase enzymes have been described so far [10,35–37]. Then the xylitol is metabolized further by xylitol dehydrogenase which is strictly NAD<sup>+</sup> dependent (Figure 2). The unbalance between NAD<sup>+</sup> supplement and requirement can block the xylose metabolization and leads to accumulation of xylitol. Later, *S. passalidarum* was found to harbor two genes encoding for xylose reductase (*SpXYL1.1* and *SpXYL1.2*) [28]. The *SpXYL1.1* gene product is more equivalent to *XYL1* found in other yeasts. The expression levels of *SpXYL1.2* were found to be higher than *SpXYL1.1* and bioethanol production in *S. passalidarum* was attributed to higher xylose reductase activity with NADH than with NADPH [28]. The *SpXYL1.2* was found to use both NADH and NADPH with preference for NADH, while *SpXYL1.1* was stringently NADPH-dependent. Furthermore, the transformation of *S. cerevisiae* with *SpXYL1.2* of *S. passalidarum* enabled the overexpressing *S. cerevisiae*::*Sp*XYL1.*2* strain to grow anaerobically on xylose and to ferment it to higher ethanol yield than the isogenic *S. cerevisiae* TMB 3422 strain, which overexpresses *P. stipitis XYL1*. While, the *S. cerevisiae*::*SpXYL1.1* overexpressing strain was not able to grow on xylose [28]. Similarly, in the yeast-like fungus *Aureobasidium pullulans*, the overexpression of *SpXYL1.2* xylose reductase along with *S. passalidarum* xylitol dehydrogenase encoded by *SpXYL2.2* enhanced the xylose metabolization by 17.76% and improved the fermentation capability and the pullulan production by 97.72% of the overexpressing mutants compared with the parental strain [38].

Finally, a metabolic analysis of *S. passalidarum* speculated that NADH-preferred xylose reductase and NAD+-dependent xylitol dehydrogenase would tend to drive both of xylose assimilation via the oxidoreductase pathway and the acetaldehyde reduction to ethanol by the alcohol dehydrogenase enzyme [19]. Recently, a metabolic flux analysis of different xylose-fermenting yeasts confirmed a better cofactors balance within *S. passalidarum* cells during xylose catabolism to bioethanol production than within *P. stipitis* cells [31], which further supports the growth characteristics of *S. passalidarum*.

Collectively, those unique and unusual traits of *S. passalidarum* encourage using it as a source for genes to improve xylose utilization and bioethanol production from lignocellulosic biomass in the current xylose fermenting yeasts, such as *P. stipites*, or to introduce xylose metabolism genes to develop industrial strains of *Saccharomyces cerevisiae* capable of co-fermentation of pentose and hexose sugars. Or alternatively, to domesticate it given the excellent results already accomplished by wild-type representatives of that species for co-fermentation of mixed sugars. To facilitate that purpose, Li et al. (2017) developed a stable genetic expression system compatible with the CUG yeasts clade for genomic integration of Gene Of Interest (GOI) into several yeasts [39]. The developed multi-host integrative system was functional in several of the xylose-fermenting yeasts including *S. passalidarum*, *P. stipitis*, and *Candida je*ff*riesii* and *Candida amazonensis*, as well as in a hexose metabolizing yeast *Saccharomyces cerevisiae*, for heterologous expression of green fluorescent protein (GFP) or lactate dehydrogenase. For lactate dehydrogenase overexpressing strains, all the engineered yeast strains were able to metabolize either glucose (in case of *S. cerevisiae*) or xylose (in case of xylose-fermenting yeasts) to produce lactate [39].

#### **5. New Adaptive Strains of** *Spathaspora passalidarum* **for Potential Industrial Applications**

One of the unique features of those xylose metabolizing yeasts, is the ability to use not only the monomeric hexose and pentose sugars but also the high oligomeric disaccharide sugar in mixed co-fermentation [9,19], which can be an advantage for large scale industrial applications. The mild acid pretreatment of agriculture waste biomasses is relatively cheap and prevents the accumulation of harmful compounds, which inhibits the fermentation processes, but releases the sugars in higher oligomeric stats. Therefore, the ability of such native xylose fermenting strains, *P.*

*stipitis*, and *S. passalidarum*, to use the high oligomeric sugar can be a great advantage for various biotechnological applications.

One of the major problems that hinders the use of *S. passalidarum* for industrial bioethanol production, even with its remarkable ability for bioethanol production, is the high sensitivity of *S. passalidarum* to the chemical inhibitors, such as ferulic and acetic acids, which are released in preparation of hemicellulosic hydrolysates [9]. Several elaborative studies have focused mainly on improving the tolerability of *S. passalidarum* to the hydrolysates inhibitors with keeping in mind the bioethanol productivity of the strains [19,40–43]. Hou and Yao in (2012) reported a strong strain [40], which is able to grow on furfurals and many other inhibitors of wheat straw hydrolysate (75%) and able to accumulate up to 0.40 g/g ethanol. Such strain was generated through hybridization of a *S. cerevisiae* and a UV-mutagenized *S. passalidarum* [40]. In 2012 also, another resistant strain was developed under O2 limiting conditions through several passage of the wild-type *S. passalidarum* NRRL Y-27907 on wood hydrolysate, followed by adaptive growth of the strain on corn stover AFEX (ammonia fiber expansion) hydrolysate [19]. Even with such efforts, the strain was not able to accumulate significant amounts of ethanol during the fermentation of the AFEX hydrolysate, despite its ability to grow in AFEX hydrolysate media. When the acetic acid was depleted from AFEX hydrolysate media, ethanol production was surprisingly observed with a yield of 0.45 g/g and most of the xylose content was consumed [19]. Later in 2017, Morales and colleagues developed an evolutionary adapted strain [41] with high tolerance toward the classical inhibitor of the fermentation processes, acetic acid, and that produces ethanol with a yield of 0.48 g/g. In a non-detoxified hydrolysate of *Eucalyptus globulus*, the authors reported also the ability of this strain to co-utilize mixed sugars of xylose, glucose, and cellobiose under microaerobic conditions [41]. This strain was generated by UV irradiation followed by successive growing of the strain under elevated acetic acid concentrations [41]. In similar way, another group also obtained a mutated *S. passalidarum* strain but via plasma mutagenesis and continuous cultivation in alkaline liquor pretreated corncob [42]. Under a simultaneous saccharification and co-fermentation, the obtained strain produced bioethanol with efficiency of 75% [42]. Finally, Su et al. in 2018 developed an adaptive *S. passalidarum* strain (named YK208-E11) [43], which is designated for resistance to AFEX hydrolysate inhibitors, from the wild-type NRRL Y-27907 through high-throughput screen via combining several approaches of batch adaptation, cell recycling, and cell mating [43]. The *S. passalidarum* YK208-E11 strain produced less biomass (about 40% compared to the wild-type), co-metabolized mixed sugars of xylose, glucose, and cellobiose, and exhibited a three-fold improvement in the ethanol production rate with a yield of 0.45 g/g. The whole genome sequence of *S. passalidarum* YK208-E11 strain revealed a deletion of about 11 kb in this strain. The ORF, which was deleted in *S. passalidarum* YK208-E11, is encoding for proteins predicted to be involved in cell division and respiration. Therefore, the authors speculated that this deletion may account for those unique adaptive/physiological features of this AFEX-acclimatized *S. passalidarum* YK208-E11 strain [43].

#### **6. Future Perspective for Engineering New Strains for Better Bioethanol Production**

The metabolic engineering approaches involve targeted overexpression and/or deletion of fermentative key genes that facilities quick and efficient conversion of sugars into bioethanol with high recoverable yields [1,3]. As we discussed above, *S. passalidarum* xylose reductase and xylitol dehydrogenase are among the promising candidates for targeted overexpression. The cumulative knowledge of the transcriptomics, metabolomics, and comparative genomics studies for *P. stipitis* and *S. passalidarum*, identified other key enzymes controlling the xylose assimilation, rather than XDH and XR (Figure 2). One of such promising key genes is *adh* that encodes for fermentative isozyme alcohol dehydrogenase (ADH), which is vital for production and/or assimilation of ethanol (Figure 2). Generally, ADH catalyzes the final (rate limiting) step in the yeast glycolytic pathway, the reduction of acetaldehyde to ethanol and NAD+, and therefore it accepts NADH as a co-factor [44,45]. However, ADH enzymes are also able to perform the reverse reaction from ethanol to acetaldehyde, enabling the yeasts to oxidize and grow on ethanol as a carbon source. In *P. stipitis*, the ADH fermentative activities

is crucial not only for ethanol production and/or consumption but also for maintenance redox balance within the yeast cell, so it is considered to be a part of the cofactor balance system in *P. stipitis* [44].

The sequencing projects of *S. passalidarum* NRRL Y-27907 and *P. stipitis* CBS6054 (JGI-MycoCosm) revealed the presence of several/different alcohol dehydrogenase (ADH) encoding genes. For example, in *P. stipitis*, sevem genes were predicted to encode for alcohol dehydrogenase (*Ps*ADH1 to *Ps*ADH7) enzymes [13,17]. Among them *Ps*ADH1 and *Ps*ADH2 were found to be essential for xylose assimilation and ethanol production [44,46]. Each of the ADH proteins in *S. passalidarum* and *P. stipitis* are supposed to have different kinetic properties. Some of the enzymes could be mainly responsible for producing ethanol while others might be responsible for oxidizing it. In *S. passalidarum*, the gene encoding for *Sp*ADH1 was found to be expressed at a very high level during xylose metabolization [34]. In addition, metabolic analysis and metabolic flux analysis revealed that alcohol dehydrogenase is one of key enzymes driving ethanol production in *S. passalidarum* [19,31]. Notably, owing to relative *Sp*ADH1 abundance, the *Sp*ADH1 promotor was used to develop a multi-host integrative system for xylose-fermenting yeast [39].

While in *P. stipitis*, the function of some ADH enzymes are better understood, in particular *Ps*ADH1 and *Ps*ADH2 [13,17,44,46–49]. Transcriptomic studies of the *P. stipites adh* system indicated that the *Ps*ADH activities are correlated with and induced under O2 limited/microaerobic conditions [46,48]. Under xylose fermentation, the *Ps*ADH1 was found to be the primary key enzyme among the *Ps*ADH system. The deletion of *Ps*ADH1 caused a reduction in *P. stipites* growth rate and a notable increase in xylitol accumulation accompanied with a dramatic decrease in ethanol production, due to intracellular cofactors imbalance [44]. The *Ps*ADH2 is not expressed under microaerobic or aerobic conditions unless *Ps*ADH1 is deleted [44,46], which further confirms that the significant role of *PsADH1* is in sugar assimilation and ethanol production. The levels of *Ps*ADH1 and *Ps*ADH2 transcripts were observed, however, to be low through xylose metabolism relative to the transcript levels of other fermentative and glycolytic enzymes [13,17]. In addition, *Ps*ADH1 and *Ps*ADH2 were able to complement the growth of the *S. cerevisiae* Δ*adh* mutant on ethanol as a sole carbon source [47]. Moreover, *Ps*ADH3 to *Ps*ADH7 were speculated to keep the balance between the cofactors NADPH and NADH [17]. However, the expression patterns of the other *Ps*ADHs on xylose and glucose under microaerobic conditions, in particular, for *Ps*ADH7 and *Ps*ADH4 are not fully understood [13]. *Ps*ADH5 was found in proximity to NADPH dehydrogenase, implying a function in maintenance the intracellular cofactors balance, however, it is not proven yet. Notably, *Ps*ADH7 was found to be upregulated under aerobic growth on xylose [50]. *Ps*ADH7 was described as a strictly NADP(H) dependant enzyme with broad spectrum for substrates-specificity, including variety of aromatic and linear aldehydes (e.g., acetaldehyde, butanal, propanal, and furfural) and alcohols (e.g., ethanol, butanol, pentanol, hexanol, and octanol) for forward and reverse reactions, respectively [50]. Surprisingly, *Ps*ADH7 was able to utilize xylitol as a substrate too with moderate activity. In the same context, the overexpression of *Ps*ADH7 into a *P. stipites* xylitol dehydrogenase mutant (Δ*Ps*XDH) [18], which cannot metabolize xylitol and therefore cannot grow on xylose as a sole carbon source, was able exclusively to complement the growth of Δ*Ps*XDH on xylose, in contrast to *Ps*ADH1, 2, 4, and 5 [50]. Hence, there is a need to understand the kinetic characteristics of each of *Ps*ADH and *Sp*ADH enzymes in order to target the correct genes for overexpression and/or deletion. Finally, we would like to state that genes encoding for *adh* isozymes are worth studying, especially of *S. passalidarum*, owning to their significant functions in bioethanol production/consumption and/or intracellular cofactor balance.

#### **7. Conclusions**

Taken together, the advances in fermentation performance by *S. passalidarum* pave the way for engineering the conventional and the nonconventional fermenting yeasts, such as *S. cerevisiae* and *P. stipites*, for economical fermentation of hexose and pentose sugars in hemicellulosic hydrolysates on industrial scales. Keeping in mind that the efficient metabolization and fermentation of xylose is essential for the bioconversion of lignocellulosic biomasses into biofuels and chemicals, but the conventional wildtype strains like *S. cerevisiae* cannot use the xylose. Therefore, researchers keep trying to engineer the xylose utilization pathway into the conventional yeast. The genomes of the natural xylose-fermenting yeasts, in particular of *P. stipitis* and *S. passalidarum,* are of huge importance, as their genomics features and regulatory patterns can serve as guides and genomic resources for further genetic engineering development in those native xylose-metabolizing yeasts or to engineer non-xylose fermenting yeasts. Therefore, *S. passalidarum* and *P. stipitis* can be considered as genomic treasure sources for various genes to engineer the xylose metabolism and to improve the bioethanol production [1,24,34].

**Author Contributions:** K.A.S. undertook extensive literature review and wrote the manuscript. S.M.E. and A.I.E.-D. supervised the research. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Acknowledgments:** The authors would like to thank Thomas Jeffries (University of Wisconsin Madison), Laura Willis (Forest Products Laboratory, USDA), Amal rabeá, Ali Selim, and Katerina Peros for the invaluable and unlimited support to complete this project. K.A.S. received the Parwon scholarship at Institute for Microbial and Biochemical Technology, FPL, USDA.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## *Article Ustilago Rabenhorstiana***—An Alternative Natural Itaconic Acid Producer**

#### **Susan Krull, Malin Lünsmann, Ulf Prüße and Anja Kuenz \***

Thünen-Institute of Agricultural Technology, Bundesallee 47, 38116 Braunschweig, Germany; susan.krull@thuenen.de (S.K.); malinluensmann@ymail.com (M.L.); ulf.pruesse@thuenen.de (U.P.)

**\*** Correspondence: anja.kuenz@thuenen.de; Tel.: +49-531-596-4265

Received: 22 November 2019; Accepted: 24 December 2019; Published: 2 January 2020

**Abstract:** Itaconic acid is an industrial produced chemical by the sensitive filamentous fungus *Aspergillus terreus* and can replace petrochemical-based monomers for polymer industry. To produce itaconic acid with alternative renewable substrates, such as lignocellulosic based hydrolysates, a robust microorganism is needed due to varying compositions and impurities. Itaconic acid producing basidiomycetous yeasts of the family *Ustilaginaceae* provide this required characteristic and the species *Ustilago rabenhorstiana* was examined in this study. By an optimization of media components, process parameters, and a fed-batch mode with glucose the final titer increased from maximum 33.3 g·L−<sup>1</sup> in shake flasks to 50.3 g·L−<sup>1</sup> in a bioreactor. Moreover, itaconic acid was produced from different sugar monomers based on renewable feedstocks by *U. rabenhorstiana* and the robustness against weak acids as sugar degradation products was confirmed. Based on these findings, *U. rabenhorstiana* has a high potential as alternative natural itaconic acid producer besides the well-known *U. maydis* and *A. terreus*.

**Keywords:** *Ustilago*; itaconic acid; process improvement; lignocellulosic feedstock

#### **1. Introduction**

Itaconic acid is an interesting chemical for the polymer industry, which is produced in a biotechnological process based on renewable substrates [1]. Petrochemical-based substances, like methacrylic or acrylic acid, can be replaced by this single unsaturated dicarbonic acid and its derivatives. Therefore, the field of products and applications is widespread, e.g., synthetic latex, styrene-butadiene rubber, superabsorbent polymers, or unsaturated polyester resins [2–6].

Since the 1960s, the filamentous fungus *Aspergillus terreus* is industrially used with a titer of 85–100 g·L−1, whereas in laboratory scale, final titers of 160 g·L−<sup>1</sup> itaconic acid are described [4,7–9]. *A. terreus* achieves a productivity up to 1.15 g (L·h)−<sup>1</sup> and a yield of 0.64 (*w*/*w*), whereby the theoretical yield with glucose is 0.72 (*w*/*w*) [7,8]. Besides pure glucose, itaconic acid was successfully produced by *A. terreus* with glycerol, starch hydrolysates, molasses, and different monosaccharides, like xylose, arabinose, galactose, and rhamnose [10]. A great cultivation challenge is caused by sugar degradation products or other impurities in lignocellulosic hydrolysates, which influence the morphology and itaconic acid production of the fungus. Due to the sensitivity of the fungus, complex purification processes are used for such hydrolysates or more resistant strains are generated by mutagenesis [11–13]. Another alternative is itaconic acid producing yeasts of the species *Candida*, *Pseudozyma,* or *Ustilago,* which are more robust and not as sensitive to metal ions as *A. terreus* [14–18]. For wildtype strains of the species *Ustilago,* low final titers of 44.5 g·L−<sup>1</sup> itaconic acid, low yields up to 0.24 (*w*/*w*), and a low productivity of maximum 0.31 g (L·h)−<sup>1</sup> are disadvantageous [19]. This is due to a variety of by-products like other organic acids, glycolipids, and intracellular triacylglycerols, which are produced in parallel to itaconic acid [16,18,20,21]. Nevertheless, in addition to the robustness of the yeasts,

the formation of haploid yeast-like cells is an advantage compared to the filamentous growth or formation of pellets of *A. terreus* with a decreased oxygen supply or increased viscosity [5,22].

In recent years, the research interest in itaconic acid production with the phytopathogenic basidiomycete *Ustilago maydis*increased. It was found, that an ammonium limitation triggers the itaconic acid overproduction in *U. maydis*[5,19] and itaconic acid is synthesized in the cytosol via the intermediate cis-aconitate and trans-aconitate and can be further converted to 2-hydroxyparaconic acid [23–25]. The itaconic acid gene cluster was also characterized and relevant enzymes, transporters, and promoters were found [23,25,26], whereby a summary of metabolic aspects is given by Wierckx et al. [27]. Based on these findings, metabolic engineering strategies and process optimization of *U. maydis* resulted in a reduction of by-product concentrations of malic acid and 2-hydroxyparaconic acid with a significant increased itaconic acid titer of 63.2 g·L−<sup>1</sup> and a yield of 0.48 (*w*/*w)* [23]. All in all, detailed examinations are available for itaconic acid production of *U. maydis*, but also other wildtype strains of the family *Ustilaginaceae* could offer advantages of less sensibility or a yeast-like morphology for using second-generation feedstocks. This family is well-known for organic acid production [16,18,28,29], but the level of knowledge about alternative itaconic acid producer, like *U. cynodontis* or *U. rabenhorstiana,* are low.

This study considers the cultivation of *Ustilago rabenhorstiana* for itaconic acid production and its potential as alternative natural producer. Although the used strain is known as natural itaconic acid producer [18,30], the microorganism was not examined in literature more precisely. Only the growth of the organism with glycerol as substrate was described, whereby non-formation of organic acids was detected [28]. Concerning itaconic acid production based on renewable resources, usability of different sugar monomers and robustness towards influence of sugar degradation products were examined in this study. Moreover, the influences of media and fermentation parameters on the production of itaconic acid and by-products as well as the morphology of the yeast were investigated.

#### **2. Materials and Methods**

#### *2.1. Microorganism*

The basidiomycete *Ustilago rabenhorstiana* NBRC 8995 was purchased from the National Institute of Technology and Evaluation (Tokyo, Japan) and was stored at −80 ◦C as 50% (*v*/*v*) glycerol stock culture.

#### *2.2. Media Compositions*

YEPS-medium was used for the preparation of agarplates and preculture (20 g·L−<sup>1</sup> sucrose, 10 g·L−<sup>1</sup> yeast extract, 20 g·L−<sup>1</sup> peptone, optional 20 g·L−<sup>1</sup> agar-agar).

If not mentioned otherwise, the production media was a Tabuchi-medium [18] containing 120 g·L−<sup>1</sup> glucose, 0.5 g·L−<sup>1</sup> KH2PO4, 1.6 g·L−<sup>1</sup> NH4Cl, 0.2 g·L−<sup>1</sup> MgSO4·7 H2O, 10 mg·L−<sup>1</sup> FeSO4·7 H2O, 1 g·L−<sup>1</sup> yeast extract, and 30 g·L−<sup>1</sup> CaCO3. All components were prepared separately in stock solutions; the pH-value was adjusted to pH 6.0 for all solutions with 0.5 m H2SO4 or 1 m NaOH and autoclaved. The pH-value of the iron-solution was not corrected, and the solution was sterile filtered. CaCO3 was weighed in the glassware and autoclaved.

All media components were p.a. quality and purchased from Merck (Darmstadt, Germany), Sigma-Aldrich (St. Louis, MO, USA) or Roth (Karlsruhe, Germany). In fed-batch cultivation the glucose concentration was monitored during the cultivation. If necessary, glucose was added in solid form without previous sterilization to prevent a glucose limitation.

#### *2.3. Cultivation*

Precultures were conducted in a 250 mL shake flask with three baffles, a filling volume of 50 mL, and inoculated with a single colony from a YEPS-agar plate (30 ◦C, 3 days). The preculture was cultivated at 30 ◦C and 120 rpm (50 mm shaking diameter) for 24 h until an optical density of 10 at 605 nm was achieved. All experiments were inoculated with 1% (*v*/*v*) of the preculture.

The main cultures in shake flasks were carried out at 30 ◦C and 120 rpm in 250 mL shake flasks with three baffles and a filling volume of 100 mL Tabuchi-medium.

To identify potential impurities and the utilization of monosaccharides based on lignocellulosic feedstock, test tubes with Kapsenberg caps were used and a working volume of 2 mL (ø 16 mm × 100 mm). The test tubes were incubated for 4 days at 30 ◦C and 120 rpm in an inclined test tube holder with an inclination angle of 30◦. Shake flasks and test tubes were continuously rotated by hand while sampling avoiding inhomogeneity.

The cultivation in bioreactors were conducted in four parallel 1 L-bioreactors, equipped with a Rushton impeller and an L-sparger (model SR0700ODLS, DASGIP GmbH, Jülich, Germany). DASGIP Control software (DASGIP GmbH, Jülich, Germany) was used for the regulation of gassing, temperature, pH-value, and stirring rate, as well as recording the data of dissolved oxygen (DO) and pH. The pH regulation to pH 6.0 was carried out with 4 m NaOH, if not otherwise mentioned. At the beginning of the cultivation, 0.5 mL antifoam solution (Ucolup N-115, Brenntag, Mühlheim/Ruhr, Germany) was added to the broth. The experiments were carried out at 30 ◦C, 500 rpm, a filling volume of 500 mL, and an aeration of 0.1 vvm, unless otherwise mentioned. All cultivations were carried out in minimum duplicates, whereby the deviation from the mean value was <5%. All results are presented as mean values without error bars on account of readability.

#### *2.4. Analytical Methods*

The samples were centrifuged at 21,000 *g* for 20 min at 20 ◦C and the supernatant was used for further analysis. A Shimadzu HPLC (Shimadzu Corp., Kyoto, Japan) with a HPX-87H column (BioRad, Munich, Germany) with a refractive index detector (RI) and UV detector at 210 nm was used to analyze the concentrations of sugars and organic acids. The column was tempered at 40 ◦C and as mobile phase a 5 <sup>m</sup> H2SO4 solution at a flow rate of 0.6 mL·min−<sup>1</sup> was used. The concentration of an unknown product was estimated by the peak area of the RI-signal compared to a calibration of succinic acid. For samples of bioreactor experiments, the pellet was washed twice with deionized water and dried to a constant weight at 105 ◦C for at least 48 h to determine the cell dry weight (CDW).

The composition of fatty acids was analyzed by transesterification of the fatty acids as described by Lewis et al. [31]. The biomass of reactor cultivation was separated from the broth by centrifugation (21,000 *g* for 20 min at 20 ◦C). The supernatant was discarded, and the pellet was washed twice with 0.9% (*v*/*v*) NaCl-solution and suspended in 0.9% (*v*/*v*) NaCl-solution. The cells were disrupted by an ultrasonic-homogenisator on ice (4 cycles: 15 s at 65%, break 30 s; Sonopuls HD2200 with sonotrodetype UW2200, Bandelin electronic, Berlin, Germany). The suspension was stored at −80 ◦C and freeze-dried (Alpha 1-2 LD, Christ, Osterode, Germany). The fatty acids were derivatized to fatty acid methyl esters (FAME) [31] and analyzed by GC-MS on a GC-17A (Shimadzu Corp., Kyoto, Japan) with a ZebronTM ZB-WAX plus column (60 m <sup>×</sup> 0.25 mm <sup>×</sup> 0.25 <sup>μ</sup>m), using 1.4 mL·min−<sup>1</sup> helium as carrier gas. The temperature gradient of 60 ◦C was increased to 150 ◦C at a rate of 30 ◦C·min−1, and afterwards increased up to 240 ◦C at a rate of 13 ◦C·min<sup>−</sup>1. The temperature of 240 ◦C was kept for 30 min and raised to 255 ◦C for 5 min. The FAMEs were identified with the software LabSolutions (Shimadzu Corp., Kyoto, Japan) and the mass spectral data were compared with the database of the National Institute of Standards and Technology (Gaithersburg, MD, USA).

#### *2.5. Microscopy*

The cells were examined using a phase-contrast microscope (Axioplan, Carl Zeiss AG, Jena, Germany) with the software analysis pro (Analysis 5.1, Olympus Soft Imaging Solutions GmbH, Münster, Germany). Intracellular lipids were visualized after coloring with nil-red by fluorescence microscopy [32].

#### **3. Results**

#### *3.1. Standard Cultivation in Shake Flasks*

A standard cultivation of *U. rabenhorstiana* with pure glucose as substrate was performed in shake flasks (Figure 1). After one day, the itaconic acid production started, and additionally, succinic acid and malic acid were produced. Furthermore, an unknown product accumulated after 48 h approximately in a concentration range of <sup>&</sup>lt;1 g·L<sup>−</sup>1. Malic acid was consumed in further course of cultivation, and after four days, α-ketoglutaric acid was formed and increased parallel with the itaconic acid concentration. Glucose was completely consumed after 9.7 days, resulting in 31.3 g·L−<sup>1</sup> itaconic acid, 13.6 g·L−<sup>1</sup> <sup>α</sup>-ketoglutaric acid, 2.3 g·L−<sup>1</sup> malic acid, and traces of an unknown metabolite, followed by a further production of <sup>α</sup>-ketoglutaric acid. The overall productivity was 0.13 g (L·h)−<sup>1</sup> with a yield of 0.26 (*w*/*w*) after 9.8 days. Despite the use of CaCO3 as buffer, the pH-value constantly decreased from pH 6.8 to 4.9 throughout the cultivation. The morphology of *U. rabenhorstiana* changed from yeast-like single cells (0–2 days; Figure 1A) via a development of pseudomycel (2–7 days, Figure 1B) to filamentous growth like long branched mycel (7–11 days, Figure 1C). Moreover, intracellular lipids deposits were visible under the microscope, which became smaller in size after the glucose limitation at day 9.7.

**Figure 1.** Cultivation of *U. rabenhorstiana* in 250 mL shake flasks in standard Tabuchi-medium (**D**) and its corresponding morphology (**A**): 0–2 days; (**B**): 2–7 days; (**C**): 7–11 days. Glucose (blue square), itaconic acid (green square), α-ketoglutaric acid (grey triangle), succinic acid (black triangle), malic acid (light grey triangle), pH (black circle), 120 rpm, 30 ◦C, 1% (*v*/*v*) inoculum.

#### *3.2. Influence of Media Components*

To determine the influence of media components, titer, yield, and productivity of a cultivation with 120 g·L−<sup>1</sup> initial glucose, performed in shake flasks after 7.8 days, are shown in Figure 2. In the case of the variating initial glucose concentrations (Figure 2A), the point in time of glucose limitation was analyzed. 50 g·L−<sup>1</sup> glucose were consumed in 3.7 days, 100 g·L−<sup>1</sup> in 6.7 days, 120 g·L−<sup>1</sup> in 8.7 days, and 150 g·L−<sup>1</sup> in 10.7 days. Further, 200 g·L−<sup>1</sup> glucose was not completely consumed by *U. rabenhorstiana* and 35 g·L−1, and remained while the concentration of itaconic acid was constant after 15 days. With increasing initial glucose concentration, the titer of itaconic acid increased, but the productivity and yield decreased slightly from 0.16 g (L·h)−<sup>1</sup> to 0.09 g (L·h)−<sup>1</sup> and from 0.27 (*w*/*w*) to 0.20 (*w*/*w*) for glucose concentrations larger than 100 g·L−1. The amount of <sup>α</sup>-ketoglutaric, succinic, and malic acid of the total organic acid concentration was raised from 15.5% (50 g·L−<sup>1</sup> glucose) over 33.7% (120 g·L−<sup>1</sup> glucose) to 50.1% (200 g·L−<sup>1</sup> glucose). In case of different ammonia chloride concentrations, the highest titer of 31.8 g·L−<sup>1</sup> itaconic acid, a productivity of 0.17 g (L·h)<sup>−</sup>1, and a yield of 0.26 (*w*/*w*) was achieved using 1.6 g·L−<sup>1</sup> NH4Cl, which corresponded to the used concentration in standard Tabuchi-medium (Figure 2B). With a lower concentration of 1 g·L−<sup>1</sup> NH4Cl and higher concentrations between 3 and 7 g·L−<sup>1</sup> NH4Cl the titer, productivity and yield decreased up to 35%. Also, the chosen concentration of 0.2 g·L−<sup>1</sup> MgSO4·7 H2O in Tabuchi-medium was optimal for itaconic acid production with *U*. *rabenhorstiana*, and a titer of 28.9 g·L−<sup>1</sup> with a productivity of 0.16 g (L·h)−<sup>1</sup> was reached (Figure 2C). Lower or higher levels of magnesium resulted in a decrease of all target values. In the concentration range of 0.1–1 g·L−<sup>1</sup> KH2PO4, there were no significant differences between the titer, yield, and productivity. All cultivations yielded in titers of 29.3 g·L−<sup>1</sup> <sup>±</sup> 1.2 g·L−<sup>1</sup> with a productivity between 0.15–0.16 g (L·h)−<sup>1</sup> and a yield of 0.24–0.25 (*w*/*w*) (Figure 2D). In the range of 0.5–25 mg·L−<sup>1</sup> FeSO4·7 H2O, the itaconic acid decreased from 32.4 g·L−<sup>1</sup> to 24.2 g·L−<sup>1</sup> (Figure 2E). The productivity of 0.17 g (L·h)−<sup>1</sup> was reduced by 17% and the yield of 0.3 (*w*/*w*) itaconic acid by 25%. With increasing yeast extract concentration (0.25–1.5 g·L<sup>−</sup>1) the titer increased to 27.2 g·L−<sup>1</sup> with a productivity of 0.15 g (L·h)−<sup>1</sup> at a concentration of 1.5 g·L−<sup>1</sup> yeast extract (Figure 2F). A further increase in the yeast extract concentration up to 2 g·L−<sup>1</sup> resulted in a decreased titer of 24 g·L−<sup>1</sup> and a lowered productivity of 0.13 g (L·h)−1. None of the media components had an influence on the filamentous growth.

#### *3.3. Monosaccharide Utilization*

It is intended to produce itaconic acid based on renewable feedstocks, e.g., lignocellulosic biomass, biomass with a high starch content, or molasses. The usability of the monosaccharides based on those feedstocks (arabinose, fructose, galactose, glucose, mannose, rhamnose, and xylose) were investigated with 100 g·L−<sup>1</sup> of each sugar in test tubes (Table 1). For the precultivation, sucrose was used. Filamentous growth occurred on glucose, fructose, mannose, and xylose. An accumulation of long hyphae, a buildup of pellets with a diameter of 50 μm grew with arabinose as substrate. The yield and productivity were very different depending on the substrate. For the reference cultivation with glucose, the productivity was 0.16 g (L·h)−<sup>1</sup> with a yield of 0.24 (*w*/*w*). The productivity of 0.09 g (L·h)−<sup>1</sup> of itaconic acid with mannose was 44% lower, while the yield was in the same range with 0.22 (*w*/*w*). Using fructose, the same productivity compared to mannose was achieved, but with a lower yield of 0.17 (*w*/*w*). *U. rabenhorstiana* was able to use both pentoses for itaconic acid production, whereby the productivity with arabinose with 0.04 g (L·h)−<sup>1</sup> was twice as high as with xylose. In the cultivation with glactose, only traces of itaconic acid were detected. The yeast was not able to produce itaconic acid or even grow with rhamnose as single substrate.


**Table 1.** Cultivation of *U. rabenhorstiana* in test tubes with different monosaccharides as substrate (four days, 30 ◦C, 120 rpm, inclination angle of 30◦, and 1% (*v*/*v*) inoculum).

**Figure 2.** Influence of Tabuchi-medium components in 250 mL shake flasks on the final titer (dashed bar), yield (black bar), and productivity (grey bar) of *U. rabenhorstiana* at 120 rpm and 30 ◦C after 7.8 days (**B**–**F**). Cultivation time for different initial glucose concentrations depended on the point of glucose limitation (**A**). Asterisks highlight the standard media composition.

#### *3.4. Influence of Sugar Degradation Products*

In case of lignocellulosic feedstocks, different sugar degradation products are formed due to the harsh conditions in the pretreatment. To test the inhibitory effect of sugar degradation products like weak acids or furan derivates, the components were added by the lowest expected concentration levels to the media. The effect of 0–2 g·L−<sup>1</sup> acetic acid, formic acid, furfural, or hydroxymethylfurfural (HMF) was carried out in test tubes (Figure 3). A productivity of 0.15 g (L·h)−<sup>1</sup> with standard Tabuchi-medium without the addition of inhibitory components was reached. Up to a concentration of 0.5 g·L−<sup>1</sup> formic

acid, the productivity did not differ. The addition of 1 g·L−<sup>1</sup> resulted in an increased productivity of approximately 1.4 times and decreased to 0.13 g (L·h)−<sup>1</sup> with 2 g·L−<sup>1</sup> formic acid. The result was very similar with the addition of acetic acid. The standard productivity increased up to 0.19 g (L·h)−<sup>1</sup> by adding 0.5 g·L−<sup>1</sup> acetic acid and was reduced to 0.13 g (L·h)−<sup>1</sup> by increasing the acetic acid concentration. Both furan derivates influenced the microorganism very strongly, amounts of 0.1 g·L−<sup>1</sup> of HMF or 0.5 g·L−<sup>1</sup> furfural already resulted in a growth inhibition. If the growth was not inhibited, the yeast grew filamentous and stored intracellular lipid droplets comparable with the cultivation without addition of inhibitors.

**Figure 3.** Inhibition effects of sugar degradation products on the itaconic acid productivity with *U. rabenhorstiana* in test tubes with Tabuchi-medium after four days, 30 ◦C, 120 rpm, inclination angle of 30◦, pH > 5.5, and 1% (*v*/*v*) inoculum. Acetic acid (blue circle), formic acid (green circle), HMF (red diamond), furfural (orange diamond).

#### *3.5. Influence of the pH-Value in 1 L-Bioreactor*

The pH-value dropped in shake flask cultivations from pH 6.7 to pH 4.9 with CaCO3 as buffer. To estimate the influence of the pH-value, the cultivation was transferred in 1 L-bioreactors with pH-control using 4 <sup>m</sup> NaOH (Table 2). Moreover, the modified Tabuchi-medium with 100 g·L−<sup>1</sup> glucose and 1.5 g·L−<sup>1</sup> yeast extract was used, based on the findings regarding the tested media components. The highest titer of 31.7 g·L−<sup>1</sup> itaconic acid with a productivity of 0.23 g (L·h)−<sup>1</sup> and a yield of 0.34 (*w*/*w*) was reached with a controlled pH of 6.0. Beside itaconic acid, 0.4 g·L−<sup>1</sup> <sup>α</sup>-ketoglutaric acid, 2 g·L−<sup>1</sup> malic acid, 2.9 g·L−<sup>1</sup> succinic acid, and the unknown product (<1 g·L<sup>−</sup>1) were produced. The rate of the byproducts did not differ among the tested pH-values; also, the pH-value did not have any influence on the filamentous growth of the yeast and formation of intracellular lipids.


**Table 2.** Cultivation results of *U. rabenhorstiana* with pH-control in modified Tabuchi-medium with 100 g·L−<sup>1</sup> glucose and 1.5 g·L−<sup>1</sup> yeast extract in 1 L-bioreactor (30 ◦C, 500 rpm, 0.1 vvm, 4 <sup>m</sup> NaOH).

#### *3.6. Influence of Aeration in 1 L-Bioreactor*

The effect of aeration was tested in 1 L-bioreactors by different aeration rates between 0.1–1 vvm, the stirring rate was kept constant, and the pH-value was regulated to pH 6.0 (Table 3). An increasing aeration rate from 0.1 to 1 vvm resulted in a decreased yield and titer of 20%, as well as in a 45% lower productivity. In contrast, the formed biomass increased from 15.7 g·L−<sup>1</sup> at 0.1 vvm to 21.3 g·L−<sup>1</sup> at 1 vvm. There were no significant differences between the by-product concentrations depending on the aeration rate, which corresponds to the concentrations described in Section 3.5.

**Table 3.** Influence of aeration on the cultivation of *U. rabenhorstiana* in modified Tabuchi-medium with 100 g·L−<sup>1</sup> glucose and 1.5 g·L−<sup>1</sup> yeast extract in 1 L-bioreactor (30 ◦C, 500 rpm, pH 6.0).


#### *3.7. Fed-Batch Mode in 1 L-Bioreactor with Glucose*

Glucose concentrations larger than 150 g·L−<sup>1</sup> resulted in a decreased yield and productivity in shake flasks (Figure 2A). For this reason, a fed batch with glucose was realized at a constant pH of pH 6.0 in a 1 L-bioreactor (Figure 4). An initial glucose concentration of 100 g·L−<sup>1</sup> was chosen. After five days, 73 g·L−<sup>1</sup> and after 10 days, 25 g·L−<sup>1</sup> glucose were added into the cultivation broth, in which the average glucose consumption rate was 0.73 g (L·h)−<sup>1</sup> for the first batch (0–5 days), 0.53 g (L·h)−<sup>1</sup> for the second batch (5–10 days), and 0.28 for the third batch (10–15 days). The yield amounted to 0.31 (*w*/*w*) in the first batch and was constant with 0.26 (*w*/*w*) in the second and third batches. The DO decreased to 2% within the first day and varied between 2%–20% during the further cultivation. After one day, the itaconic acid production started and rose to a final titer of 50.3 g·L−<sup>1</sup> within 15 days. Beside itaconic acid, 3.6 g·L−<sup>1</sup> malic acid, 13.6 g·L−<sup>1</sup> succinic acid, 2.5 g·L−<sup>1</sup> <sup>α</sup>-ketoglutaric acid, and the unknown product (<10 g·L<sup>−</sup>1) were formed by 17.2 g·L−<sup>1</sup> filamentous biomass (Appendix A, Figure A2). This cultivation resulted in a productivity of 0.14 g (L·h)−<sup>1</sup> with an overall yield of 0.27 (*w*/*w*) after 15 days. Further, 175 mL of a 4 m NaOH was used to keep the pH-value constant at pH 6.0. After 15 days, the cell dry weight was analyzed regarding the fatty acids (Appendix A, Table A1); C16:0, C18:0, and C18:2 were the main elements.

**Figure 4.** Fed batch with glucose in 1 L-bioreactor with modified Tabuchi-medium (initial glucose concentration 100 g·L−<sup>1</sup> and 1.5 g·L−<sup>1</sup> yeast extract) at 30 ◦C, 500 rpm, 0.1 vvm, pH 6.0 with 4 <sup>m</sup> NaOH as base and 1% (*v*/*v*) inoculum. Glucose (blue square), itaconic acid (green square), pO2 (grey line), pH (black line), cell dry weight (CDW) (orange asterisk), arrows symbolize the addition of glucose.

#### **4. Discussion**

In a standard cultivation in shake flasks, a final titer of 31.3 g·L−<sup>1</sup> itaconic acid was achieved after 9.7 days without media and process optimization of *U. rabenhorstiana*. Additionally, succinic acid, malic acid, α-ketoglutaric acid, an unknown product, and intracellular lipids were formed. Guevarra and Tabuchi reached a titer of about 16 g·L−<sup>1</sup> itaconic acid in the same media after seven days and verified 2-hydroxyparaconic acid, itatartaric acid, and erythritol as byproducts with a total concentration of 30 g·L−<sup>1</sup> [18]. Furthermore, they reduced the byproduct concentration to 19 g·L−<sup>1</sup> with a constant itaconic acid titer by using an unbuffered media, whereby the final pH-value was pH 2.8 [30]. Erythritol as the unknown byproduct could be excluded in this work. In comparison with the literature, the unknown product (<1 g·L−1) could either be 2-hydroxyparaconic acid or itatartaric acid (Appendix A, Figure A1) [18,23,30] and must be characterized precisely in a further study. Moreover, the formation of different organic acids and intracellular lipid bodies by *Ustilaginaceae* besides itaconic acid production are very well known [5,19], as well as the production of cellobiose lipids and mannosylerythritol lipids [20,29], which were not proved in this study.

A point-by-point analysis of each media component indicated that the chosen concentrations of each component in the Tabuchi-medium were nearly optimal for itaconic acid production with *U. rabenhorstiana*. Only the glucose and yeast extract concentrations were adjusted from 120 g·L−<sup>1</sup> to 100 g·L−<sup>1</sup> for glucose and from 1 g·L−<sup>1</sup> to 1.5 g·L−<sup>1</sup> for yeast extract in the modified Tabuchi-medium. Increased glucose concentration resulted in an increased final titer, but the yield and productivity decreased with glucose concentrations >100 g·L−1. The decreased yield and productivity can be explained by a higher osmotic stress and a higher number of byproducts. A very similar result was obtained by the production of itaconic acid with *U. maydis* [19]. This microorganism also showed lower yields at higher initial glucose concentrations, explained by the formation of other organic

acids, polyols, and glycolipids. The increase of yeast extract by 0.5 g·L−<sup>1</sup> raised the titer, productivity, and yield. A further increase of yeast extract did not result in an improved titer, suggesting that a limitation of vitamins, amino acids, salts, trace elements or nucleic acids [33] is prevented with a concentration >1.5 g·L−<sup>1</sup> yeast extract. The itaconic acid overproduction by *U. maydis* or *P. antarctica* are mainly triggered by an ammonium limitation, higher concentrations of NH4Cl <sup>&</sup>gt; 4 g·L−<sup>1</sup> resulted in lower itaconic acid yields and an increase in biomass for *U. maydis* [5,14,19]. What is more, in this study, the ammonium concentration had the strongest influence on the fermentation performance of *U. rabenhorstiana.* A concentration of 1.6 g·L−<sup>1</sup> NH4Cl was optimal for itaconic acid production, and higher concentrations resulted in reduced yields and productivities as described already for *U. maydis*. For this reason, it can be assumed that an ammonium limitation caused the itaconic acid overproduction of *U. rabenhorstiana*. Further, the nitrogen limitation caused the accumulation of intracellular lipid droplets, which are mostly triacylglycerols [21,34]. A secretion of cellobiose lipids and mannosylerythritol lipids in the form of needle-like crystals or oily droplets of *U. maydis* [20], could not be verified for *U. rabenhorstiana*.

To use alternative, low-cost, or lignocellulosic feedstocks for itaconic acid production, it is important to use a microorganism, which is able to consume different monosaccharides and is robust towards varying impurities. In this study, it was worked out that *U. rabenhorstiana* is able to grow and produce itaconic acid from different monosaccharides like glucose, fructose, mannose, xylose, arabinose, and galactose. The highest productivity was reached with glucose, followed by fructose and mannose. Because the precultivation was based on sucrose, composed of glucose and fructose, it can be assumed that the cultivation with fructose was therefore such successful. An adaption to the used monosaccharide of the microorganisms in the preculture or a mixture of several monosaccharides with glucose would probably lead to higher productivity and yield using that single monosaccharide in the main culture. Furthermore, the plant pathogen *U. rabenhorstiana* is supposed to degrade a range of biomass-based polymers [35–37]. For industrial itaconic acid production, the filamentous fungus *A. terreus* is used, which is very sensitive to weak acids, furan derivates, metal ions, and other impurities, which are contained in such substrates [11,12,38]. *U. maydis* is described as a very robust microorganism [5,22,39]. *U. rabenhorstiana* was also not influenced by the addition of weak acid concentrations up to 2 g·L−<sup>1</sup> in the main culture. A major advantage of the cultivation of *Ustilaginaceae* is the pH range of 5.0–6.5, whereby the dissociated weak acids cannot cross the plasma membrane into the cytosol and affect the intracellular pH-value [40,41]. A positive effect on the itaconic acid productivity was even achieved with the addition of 0.5 g·L−<sup>1</sup> acetic acid or 1 g·L−<sup>1</sup> formic acid. This positive influence of low weak acid concentrations in cultivation media is already known from itaconic acid production with *A. terreus*, ethanol production with *S. cerevisiae,* or enzyme production with *T. reesei* [12,42–44]. In contrast, low concentrations of 0.1 g·L−<sup>1</sup> HMF or 0.5 g·L−<sup>1</sup> furfural are growth limiting factors; both furan derivates reduce the activity of a number of important intracellular enzymes of the maintenance metabolism, e.g., pyruvate dehydrogenase [45]. In particular, the activity of the pyruvate dehydrogenase is essential for cells, because this enzyme links glycolysis and citric acid cycle, which supplies the cell with energy intermediates. When using lignocellulosic biomass as feedstock for *U. rabenhorstiana*, it should be taken into account, that some robustness in relation to weak acids exists, but furan derivates influence the microorganisms mostly up to growth inhibition.

For further characterization of *U. rabenhorstiana*, the fermentation was transferred in 1 L-bioreactors to investigate the influence of pH-value and aeration. A constant pH of 6.0 and modified Tabuchi-medium yielded in the highest itaconic acid titer of 31.7 g·L<sup>−</sup>1, which is comparable with the titer in standard shake flask cultivation with Tabuchi-medium. However, the overall productivity was 1.7 times higher at a constant pH-value than in shake flask experiments with CaCO3, whereby the pH-value continuously decreased to pH 4.9. Moreover, the total concentration of by-products decreased by 66%, which resulted in an increased itaconic acid yield. These suggested that the buffer capacity in shake flasks is insufficient. Buffer systems like CaCO3 or MES and its buffer capacity have a significant impact on the organic acid production of *Ustilaginaceae* not only on titer, but also on the ratio of products [16,24,30]. The higher the buffer capacity, the better is the itaconic acid titer in small-scale experiments, but *U. maydis* achieved the highest itaconic acid titer of 45.5–63.2 g·L−<sup>1</sup> in cultivations with a constant pH of 6.0–6.5 in bioreactors [19,23]. Consequently, *U. maydis* and *U. rabenhorstiana* have nearly the same requirements in pH and productivity. Titer and yield are positively influenced by a constant pH-value.

In the literature, no detailed studies regarding influence of oxygen levels on the organic acid production with *Ustilaginaceae* exist. Only cultivation parameters like high shaking frequencies or stirrer speeds led to the conclusion that a high input of oxygen is necessary [10]. Contrary results were achieved for itaconic acid production with *U. rabenhorstiana*; the lowest aeration rate of 0.1 vvm and a constant stirring rate of 500 rpm yielded the best result regarding titer, productivity, and yield. Presumably, the increase in these values is related to the formation of 36% more biomass at higher aeration rates, because of a better supply of oxygen. In batch experiments, a maximum itaconic acid concentration of 33.3 g·L−<sup>1</sup> with an initial glucose concentration of 200 g·L−<sup>1</sup> was achieved, proving that initial glucose concentration <sup>≥</sup>100 g·L−<sup>1</sup> and a constant pH-value of pH 6.0 have a significant impact on the itaconic acid production with *U. rabenhorstiana*. Therefore, a cultivation in fed-batch mode with glucose was realized at pH 6.0 and resulted in 50.3 g·L−<sup>1</sup> itaconic acid. Comparing the batch cultivation with 200 g·L−<sup>1</sup> glucose with the fed-batch cultivation in a bioreactor, the productivity and yield were 1.4 times and the final titer 1.5 times higher. Also, the formation of organic acid as byproducts was reduced. The number of byproducts of the overall organic acid concentration decreased from 50% in batch cultivation to 28% in fed-batch mode. Thereby, the amount of organic acids shifted from α-ketoglutaric acid as the main byproduct in batch mode to succinic acid in fed-batch mode, due to the pH-value of each cultivation [16]. The final titer and yield of this study are slightly increased compared to a wildtype strain of *U. maydis*, which reached 44.5 g·L−<sup>1</sup> [19]. A higher final titer up to 63.2 g·L−<sup>1</sup> or yield of 0.48 (*w*/*w*) was only obtained by a genetical modification of *U. maydis* [23]. All in all, the fermentation broth was diluted by addition of 175 mL NaOH as base. Considering the dilution, the wildtype of *U. rabenhorstiana* demonstrates the potential to produce up to 68 g·L−<sup>1</sup> itaconic acid. However, in all main cultures, the unicellular growth of *U. rabenhorstiana* shifted to filamentous cells with depots of intracellular lipids. Neither the variation of media components nor the investigation of process parameters influenced the filamentous growth. The typical unicellular yeast-like growth of *U. maydis* for itaconic acid could not be achieved for *U. rabenhorstiana* in this study. This morphology would be a great advantage compared to the filamentous *A. terreus* regarding oxygen supply or viscosity, especially in large-scale fermentations [5,22], but was not focused in this study. In general, it is possible to generate a stable unicellular growth by deleting several genes [24]. The filamentous growth involved the accumulation of intracellular lipid droplets, which is initiated by nitrogen limitation, which in turn is needed for itaconic acid overproduction [5,19,34]. Furthermore, 89% of all fatty acids in *U. rabenhorstiana* were long-chain fatty acids C16:0, C18:0, and C18:2 and suggested the accumulation of triacylglycerols in the cells. The lipid bodies in *U. maydis* mainly contain triacylglycerols consisting of palmitic, linoleic, and oleic acids [21].

#### **5. Conclusions**

This study describes a known, but so far unspecified, itaconic acid producer—*U. rabenhorstiana*. The cultivation in shake flasks with a maximal final titer of 33.3 g·L−<sup>1</sup> itaconic acid was transferred in a bioreactor. With a controlled pH-value, a low initial glucose concentration, and fed-batch mode, a final titer of 50.3 g·L−<sup>1</sup> was achieved, which is comparable with titer of other wildtype strains of *Ustilago* described in literature. However, the productivity and yield are rather low compared to *U. maydis*, which was studied very precisely in the last years regarding cultivation and process parameters as well as metabolic engineering strategies for further improvements in itaconic acid production. Transferring this knowledge from *U. maydis* to *U. rabenhorstiana* could result in a further increased final titer and improved yield and productivity. Particular attention should be paid to the morphology of the yeast and minimization of byproducts, mainly the formation of intracellular lipid droplets. Moreover,

the wildtype strain *U. rabenhorstiana* turned out to be a robust and promising alternative itaconic acid producer based on renewable resources. All in all, this study serves a basis for further promising research regarding lignocellulosic hydrolysates.

**Author Contributions:** S.K. and M.L. designed and performed the experiments. U.P. and A.K. supervised the experiments. S.K. wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was carried out in the framework of the European Research Area Network for Industrial Biotechnology (ERA-IB project "Production of Organic Acids for Polyester Synthesis (POAP)") and was funded by the German Federal Ministry of Food and Agriculture, following a decision of the German Bundestag, via the Agency of Renewable Resources (Grant No. 22029312).

**Conflicts of Interest:** The authors declare no conflict of interest. The funder had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

#### **Appendix A**

**Figure A1.** Representative HPLC analysis of a final sample (15 days) of a fed-batch cultivation in 1 L-bioreactor (Section 3.7). α-ketoglutaric acid (α-Ket), glucose (Glu), malic acid (Mal), unknown product, succinic acid (Suc), itaconic acid (Ita).

**Figure A2.** Corresponding morphology of *U. rabenhorstiana* in a fed-batch cultivation in 1 L-bioreactor (Section 3.7) after 1.1 days (**A**) and 6.6 days (**B**).


**Table A1.** Analysis of fatty acids in intracellular lipid droplets in *U. rabenhorstiana* of a fed-batch cultivation in 1 L-bioreactor (Section 3.7).

#### **References**


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