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Wood-feeding termite gut symbionts as an obscure yet promising source of novel manganese peroxidase-producing oleaginous yeasts intended for azo dye decolorization and biodiesel production
Biotechnology for Biofuels volume 14, Article number: 229 (2021)
The ability of oxidative enzyme-producing micro-organisms to efficiently valorize organic pollutants is critical in this context. Yeasts are promising enzyme producers with potential applications in waste management, while lipid accumulation offers significant bioenergy production opportunities. The aim of this study was to explore manganese peroxidase-producing oleaginous yeasts inhabiting the guts of wood-feeding termites for azo dye decolorization, tolerating lignocellulose degradation inhibitors, and biodiesel production.
Out of 38 yeast isolates screened from wood-feeding termite gut symbionts, nine isolates exhibited high levels of extracellular manganese peroxidase (MnP) activity ranged between 23 and 27 U/mL after 5 days of incubation in an optimal substrate. Of these MnP-producing yeasts, four strains had lipid accumulation greater than 20% (oleaginous nature), with Meyerozyma caribbica SSA1654 having the highest lipid content (47.25%, w/w). In terms of tolerance to lignocellulose degradation inhibitors, the four MnP-producing oleaginous yeast strains could grow in the presence of furfural, 5-hydroxymethyl furfural, acetic acid, vanillin, and formic acid in the tested range. M. caribbica SSA1654 showed the highest tolerance to furfural (1.0 g/L), 5-hydroxymethyl furfural (2.5 g/L) and vanillin (2.0 g/L). Furthermore, M. caribbica SSA1654 could grow in the presence of 2.5 g/L acetic acid but grew moderately. Furfural and formic acid had a significant inhibitory effect on lipid accumulation by M. caribbica SSA1654, compared to the other lignocellulose degradation inhibitors tested. On the other hand, a new MnP-producing oleaginous yeast consortium designated as NYC-1 was constructed. This consortium demonstrated effective decolorization of all individual azo dyes tested within 24 h, up to a dye concentration of 250 mg/L. The NYC-1 consortium's decolorization performance against Acid Orange 7 (AO7) was investigated under the influence of several parameters, such as temperature, pH, salt concentration, and co-substrates (e.g., carbon, nitrogen, or agricultural wastes). The main physicochemical properties of biodiesel produced by AO7-degraded NYC-1 consortium were estimated and the results were compared to those obtained from international standards.
The findings of this study open up a new avenue for using peroxidase-producing oleaginous yeasts inhabiting wood-feeding termite gut symbionts, which hold great promise for the remediation of recalcitrant azo dye wastewater and lignocellulosic biomass for biofuel production.
Lignocellulosic biomass is a critical raw resource for a number of existing industries, such as forestry, pulp and paper, and growing second-generation biofuel manufacturing [1,2,3,4,5,6,7]. Among lignocellulosic components, lignin is the major and most recalcitrant aromatic macromonomer compound responsible for its strength [8, 9]. So far, the only source of energy that has been known to be easily accessible for both biomass-based and green energy production as well as the bio-based manufacture of aromatics and new polymers is lignin [10, 11]. The amorphous, heteropolymeric lignin, which comprises phenylpropanoid units, may be deconstructed into low molecular weight and monomeric aromatic compounds that serve as the building block for high-value products [12, 13]. Recent economic analyses have revealed that lignin valorization might produce at least ten times more value than burning it for energy production [14,15,16,17]. However, the cost of delignification processing, required in many industrial systems, has proven difficult and expensive. To adequately address these difficulties, microbial degradation was introduced. Wood-rotting basidiomycetes have been widely investigated for their inherent and efficient delignification activity [18,19,20]. For large-scale lignin biodegradation, delignifying enzymes such as lignin peroxidase (LiP; EC 188.8.131.52), manganese peroxidase (MnP; EC 1:11:1:13), and laccase (Lac; EC 184.108.40.206) were introduced. MnP is the major enzyme class in lignin biodegradation by white-rot fungi [21, 22]. For the pulp and paper industries, as well as for the management of xenobiotic pollutants, such as textile dyes, using lignin-degrading organisms and their constituent enzymes has become appealing [23,24,25,26].
Large amounts of potentially hazardous pollutants have been released into the biosphere as a result of global industrialization [27, 28]. Cleaning up the environment by removing dangerous chemicals from textile effluents is a critical and difficult subject that necessitates a variety of ways to find long-term solutions . Among the numerous types of dyes used in the textile processing industry, azo-dyes are widely employed, and they account for over 70% of the dyestuff market . Physical and chemical methods for treating azo-dyes have been used, however, resulting in secondary contamination from the release of hazardous by-products [24, 31]. As a result of its safety, efficiency, and capacity to change dangerous chemicals into less toxic compounds, microbial treatment of azo-dyes has grown in popularity. The ability of dye-decolorizing/ligninolytic microbes to degrade resistant xenobiotics, such as lignin-mimicking synthetic dyes, may represent a significant opportunity for the biodegradation of such recalcitrant xenobiotics. Recent research has extensively looked at the bioremediation of azo dyes by bacteria and fungus [32, 33]. Still, the records on yeasts' success against difficult azo dyes are confined to only a few documents [34,35,36]. Yeasts possess higher resistance to harsh conditions, such as severe pH, osmolality, and temperature. At the same time, they have excellent biodegradation power by creating numerous enzymes that target certain harmful contaminants .
Biodiesel, also known as fatty acid methyl ester (FAME) is a viable alternative resource that can efficiently replace petroleum-based diesel in terms of new renewable energy sources [38, 39]. To this end, single cell oil (SCO), also known as a microbial lipid, has received significant attention due to its renewability, sustainability, and extensive biotechnological use. At the same time, one of the most critical criteria of SCO quality is the fatty acid composition and its similarity with vegetable oil . The major lipid type of SCO is triacylglycerides (TAG), which are composed of long-chain fatty acids. Several microbial species, including fungi, yeasts, bacteria, and microalgae, can produce and accumulate enormous amounts of lipids under particular conditions . Yeasts are classified as oleaginous when their ability to accumulate neutral lipids, particularly TAG, exceeds 20% (w/w) on a dry weight basis . Thus, lipid-accumulating yeast species from the yeast genera Yarrowia, Candida, Cryptococcus, Rhodotorula, and Lipomyces have received increased attention for biofuel production [43, 44]. Concurrently, the ability of oleaginous yeasts to metabolize aromatic substances may boost lipid yields, since the ortho-cleavage pathway of aromatic metabolism creates acetyl CoA and pyruvic acid, both of which are precursors for fatty acid biosynthesis . Furthermore, yeasts have been identified as prospective enzyme manufacturers, producing xylanases, cellulases, proteases, and lipases, all of which are widely used in bioenergy and waste management [35, 43, 46].
Several studies have recently focused on the gut digestome of xylophagous insects, such as wood-feeding termites (WFTs), to find and understand their specific symbiont activities and potential biotechnological applications. However, the highly specialized gut symbionts of WFTs, notably yeast symbionts, are still little understood. To our knowledge, the biotechnological potential of some promising yeast species derived from Reticulitermes chinensis and Coptotermes formosanus for degrading recalcitrant xenobiotics such as textile azo dyes has been emphasized for the first time [32, 47]. Furthermore, the potential of oleaginous yeasts inhabiting WFTs to degrade lignin and lignin-like dyes into metabolites usable for biodiesel production is almost unexplored. Under this scope, this study aims at exploring MnP-producing oleaginous yeasts capable of decolorizing textile azo dyes and tolerating lignocellulose degradation inhibitors. As a result, peroxidase-producing oleaginous yeasts may be a potential candidate for bioremediating textile wastewater, valorizing lignocellulose, and producing biodiesel.
Screening of peroxidase- and lipid-accumulating yeasts inhabiting WFT guts
In this study, C. formosanus and R. chinenesis were used to isolate and screen MnP- and lipid-accumulating yeasts intended for decolorizing textile-derived azo dyes, tolerating lignocellulose degradation inhibitors, and producing biodiesel (FAME). Based on clear zone development on yeast extract peptone dextrose (YEPD) supplemented with o-Dianisidine dihydrochloride as a precursor of numerous azo dyes and a peroxidase substrate, 38 isolated yeasts yielded positive findings for MnP production. Furthermore, the emergence of a reddish-brown color between 31 yeast colonies after plate treatment with H2O2 was confirmed, indicating the production of the MnP enzyme. Nine isolates designated as PPY-3, PPY-4, PPY-11, PPY-13, PPY-17, PPY-22, PPY-27, PPY-29, and PPY-35 exhibited high levels of extracellular MnP activity after 5 days of incubation in an optimal substrate. This activity ranged between 23 and 27 U/mL. On the other hand, 14 of 22 MnP-producing yeast isolates were qualitatively stained blue for TAG accumulation. Fluorometric analysis with Nile red staining, however, indicated seven yeast isolates capable of MnP production, namely, PPY-4, PPY-11, PPY-18, PPY-22, PPY-27, PPY-32, and PPY-35, with substantial yellow-gold colouring.
Table 1 depicts the molecular identification and relationship of these seven MnP-producing/TAG-accumulating yeasts to their closest phylogenetic relatives based on BLAST comparisons to the GeneBank database. The four yeast isolates PPY-11, PPY-22, PPY-4, and PPY-35 were identified as Meyerozyma caribbica strain SSA1654, Meyerozyma guilliermondii strain SSA1547, Candida stauntonica strain SSA1653, and Debaryomyces hansenii strain SSA1502, respectively, belonging to the Ascomycota phylum. The remaining three yeast isolates, PPY-32, PPY-27, and PPY-18, belonged to the Basidiomycota phylum and were identified as Fellozyma inositophila strain SSA1579, Sterigmatomyces halophilus strain SSA1655, and Vanrija humicola strain SSA1514, respectively.
The lipid profiles of the selected seven yeast strains were evaluated after 5 days of growth at 28 °C in the presence of glucose and under nitrogen limitation (Figs. 1 and 2). M. guilliermondii SSA1547 has the highest biomass value (15.78 ± 0.45 g/L). It revealed a non-significant difference (p 0.1065) with biomass produced by V. humicola SSA1514, but a significant difference (p 0.0158) with biomass produced by the closest yeast strain, M. caribbica SSA1654 (Fig. 1a). In terms of lipid yield (Fig. 1b), there was no significant difference between M. guilliermondii SSA1547 (7.10 ± 0.85 g/L), V. humicola SSA1514 (6.62 ± 1.13 g/L), and M. caribbica SSA1654 (6.95 ± 1.77 g/L). It differed significantly (p 0.0215) from the lipid produced by D. hansenii SSA1502 (4.51 ± 0.88 g/L) (Fig. 1b). The lipid content was subsequently quantified to verify the yeasts’ oleaginous nature (Fig. 1c). S. halophilus SSA1655, F. inositophila SSA1579, and C. stauntonica SSA1653 accumulated less than 20% lipids per dry biomass when lipid content was determined (Fig. 1c). In contrast, four yeast strains had lipid accumulation greater than 20%, with M. caribbica SSA1654 having the highest lipid content (47.25 ± 1.84%, w/w), followed by M. guilliermondii SSA1547 (44.74 ± 6.09%, w/w), V. humicola SSA1514 (43.46 ± 5.34%, w/w), and D. hansenii SSA1502 (33.96 ± 5.72%, w/w) (Fig. 1c). As a result, S. halophilus SSA1655, F. inositophila SSA1579, and C. stauntonica SSA1653 were found to be non-oleaginous yeasts, whereas M. caribbica SSA1654, M. guilliermondii SSA1547, V. humicola SSA1514, and D. hansenii SSA1502 were found to be oleaginous yeast strains. Although M. caribbica SSA1654 had no significant difference in lipid content from M. guilliermondii SSA1547 (p 0.5319) or V. humicola SSA1514 (p 0.3098), the latter three strains had significantly higher lipid content than S. halophilus SSA1655 (p < 0.0001) and D. hansenii (p 0.0186). YL/x, the lipid yield per gram of total biomass produced, is shown in Fig. 1d. When compared to oleaginous and non-oleaginous yeast strains examined, M. caribbica SSA1654 had the highest YL/x (0.340 ± 0.07 g/g). Furthermore, M. caribbica SSA1654 exhibited no significant difference (p 0.8206) from V. humicola SSA1514 in terms of YL/x. When compared to M. guilliermondii SSA1547 (p 0.6526) and D. hansenii SSA1502 (p 0.1361), the latter two strains exhibited no significant difference in YL/x. M. caribbica SSA1654, V. humicola SSA1514, and M. guilliermondii SSA1547, on the other hand, revealed a substantially greater YL/x than S. halophilus SSA1655 (p 0.0001) (Fig. 1d).
Yx/s, the yield of biomass produced per gram of glucose consumed, is shown in Fig. 2a. M. caribbica SSA1654 had the highest Yx/s (0.50 ± 0.03 g/g), with a considerably higher Yx/s (p 0.0038) than S. halophilus SSA1655. However, no significant difference in Yx/s was determined between M. caribbica SSA1654 and V. humicola SSA1514 (p 0.1695) or M. guilliermondii SSA1547 (p 0.3308). YL/s, the lipid yield produced per gram of glucose consumed, is shown in Fig. 2b. In terms of YL/s, V. humicola SSA1514 had the highest YL/s value of 0.23 ± 0.04 g/g, with a significantly higher YL/s than S. halophilus SSA1655 (0.09 ± 0.01 g/g; p 0.0042). There was no significant difference in volumetric productivity of biomass (Qx) between V. humicola SSA1514 (0.15 ± 0.009 g/L/h) and M. caribbica SSA1654 (p 0.1535) or M. guilliermondii SSA1547 (p > 0.99). V. humicola SSA1514, on the other hand, had a significantly higher Qx than S. halophilus SSA1655 (p < 0.0072) (Fig. 2c). M. caribbica SSA1654 had the highest volumetric production of lipids (QL; 0.08 ± 0.003 g/L/h), as shown in Fig. 2d, with a significant difference from M. guilliermondii SSA1547 (p 0.004). Table 2 compares lipid production of the oleaginous strains to that of other oleaginous yeasts reported in the literature.
Profiles of oleaginous and non-oleaginous yeast fermentation
Six yeast strains were compared in terms of growth, residual glucose, and nitrogen assimilation (Fig. 3). All oleaginous and non-oleaginous yeast strains were cultured for 5 days at 28 °C under nitrogen limitation in the presence of glucose. All yeast strains displayed typical growth curves, including exponential and stationary phases for the oleaginous yeast strains V. humicola SSA1514, M. caribbica SSA1654, M. guilliermondii SSA1547, and D. hansenii SSA1502 (Fig. 3a–d) and the non-oleaginous yeast strains S. halophilus SSA1655 and F. inositophila SSA1579 (Fig. 3e, f). Nitrogen limitation, as expected, did not greatly boost yeast growth, as evidenced by the evolution of OD600. As a result, cells entered the stationary phase after 24 h of growth. Furthermore, as a result of nitrogen limitation, lower glucose uptake rates were found, and hence the profiles of glucose intake followed the drop in nitrogen concentration. Glucose was consumed in that context for up to 120 h of cultivation. After 5 days of incubation at 28 °C, 10–12.7% of glucose remained in S. halophilus SSA1655 and F. inositophila SSA1579 cultures, but it reduced to 3.9–5.2% in V. humicola SSA1514, M. caribbica SSA1654, M. guilliermondii SSA1547, and D. hansenii SSA1502 cultures. After 24 h, however, nitrogen was significantly reduced (86–96.4%) and thereafter remained constant for all oleaginous and non-oleaginous yeasts (Fig. 3).
Construction of NYC-1 consortium
In this study, a new yeast consortium NYC-1 which stands for the molecularly identified species M. caribbica strain SSA1654, D. hansenii strain SSA1502, M. guilliermondii strain SSA1547, and V. humicola strain SSA1514 was successfully developed. The potential of MnP-producing oleaginous yeast consortium members to decolorize textile azo dyes, withstand lignocellulose degradation inhibitors, and produce biodiesel was investigated for multipurpose applications, including bioremediation, lignocellulose valorization, and biofuel production.
The NYC-1 consortium's M. caribbica SSA1654, D. hansenii SSA1502, M. guilliermondii SSA1547, and V. humicola SSA1514 strains were examined for their β-glucosidase, CMCase, xylanase, and lipase activities (Fig. 4). Among the lignocellulolytic enzymes tested, M. guilliermondii SSA1547 gave a maximum yield of β-glucosidase (0.64 ± 0.07 U/mg) on day 3, whereas D. hansenii SSA1502 produced the least β-glucosidase activity (0.061 ± 0.005 U/mg) (Fig. 4a). M. caribbica SSA1654 had the highest CMCase yield (0.17 ± 0.03 U/mg) on day 3, while V. humicola SSA1514 had the lowest CMCase activity (Fig. 4b). Furthermore, M. caribbica SSA1654 had the highest xylanase yield (5.8 ± 0.9 U/mg) on day 4 when compared to M. guilliermondii SSA1547, which had the lowest xylanase activity (2.62 ± 0.1 U/mg) on day 4 (Fig. 4c). The time-course analysis, on the other hand, revealed that the maximal lipase synthesis in all yeast strains occurred after 3 days of incubation (data not shown). At the time, the lipase production rates of the yeast strains tested ranged from 25.6 ± 3.2 U/mL for D. hansenii SSA1502 to 55.3 ± 5.4 U/mL for M. caribbica SSA1654 (Fig. 4d). M. guilliermondii SSA1547, M. caribbica SSA1654, D. hansenii SSA1502, and V. humicola SSA1514 produced β-glucosidase, CMCase, xylanase, and lipase activities simultaneously, implying that these MnP- and TAG-producing yeasts could be efficiently employed for the valorization of organic and fatty substrates.
Tolerance to inhibitors derived from lignocellulose degradation
To investigate the inhibitory effect on growth of the four MnP-producing oleaginous yeast strains comprising the NYC-1 consortium, five representative lignocellulose degradation inhibitors were chosen: furfural, 5-hydroxymethyl furfural (5-HMF), acetic acid, vanillin, and formic acid. As given in Table 3, all yeast strains M. caribbica SSA1654, M. guilliermondii SSA1547, D. hansenii SSA1502, and V. humicola SSA1514 could grow in the presence of all inhibitory substances in the tested range. Among the four strains, M. caribbica SSA1654 and M. guilliermondii SSA1547, showed the highest tolerance to furfural (1.0 g/L), while M. caribbica SSA1654 showed the highest tolerance to 5-HMF (2.5 g/L) and vanillin (2.0 g/L). Furthermore, M. caribbica SSA1654 could grow in the presence of 2.5 g/L acetic acid but grew moderately. Based on the findings, M. caribbica SSA1654 was chosen for future study, because it produced the highest lipid concentration, accumulated the most lipid, and was tolerant to lignocellulosic inhibitory compounds.
Effect of inhibitors derived from lignocellulose degradation on lipid accumulation
The effects of inhibitors derived from lignocellulose degradation, viz. furfural, formic acid, 5-HMF, acetic acid, and vanillin, on the biomass and lipid production of the selected MnP-producing oleaginous yeast strain, M. caribbica SSA1654, were investigated in this study (Table 4). The biomass and lipid concentrations in the absence of inhibitory compounds (the control) were 14.65 ± 0.18 and 6.88 ± 1.65 g/L, respectively, representing 47.3% of dry biomass. Clearly, furfural and formic acid had a significant inhibitory effect on biomass production and lipid accumulation by M. caribbica SSA1654, compared to the other lignocellulose degradation inhibitors tested, which had a minor effect. Our findings revealed that as formic acid concentrations increased (0.1 to 0.5 g/L), biomass and lipid concentrations decreased gradually, and growth completely stopped when formic acid concentration reached 0.5 g/L. Adding acetic acid at a concentration of 0.1 g/L, on the other hand, improved lipid accumulation (7.11 ± 1.84 g/L) when compared to the control (6.88 ± 1.65 g/L). When acetic acid concentrations were between 0.1 and 0.5 g/L, there was little difference in biomass production, but when acetic acid concentration was 1.0 g/L, there was a significant decrease in biomass production (5.12 ± 0.0 g/L). The lipid accumulation of M. caribbica SSA1654 was strongly inhibited by furfural. Its lipid production was clearly reduced even at the lowest tested concentration of furfural (0.05 g/L), with a 55% reduction in lipid production. However, even at concentrations of up to 5.0 g/L, 5-HMF had little effect on biomass production (27%) and lipid accumulation (31%). Similar to 5-HMF, vanillin showed less inhibitory effects on biomass production and lipid accumulation of M. caribbica SSA1654 at tested vanillin concentrations (0.1–1.0 g/L).
Azo dye decolorization performance
The decolorization of nine azo dyes was studied using the newly constructed MnP-producing oleaginous yeast consortium NYC-1. The constructed NYC-1 consortium, as shown in Fig. 5a, demonstrated effective decolorization of all individual azo dyes tested within 24 h, up to a dye concentration of 250 mg/L, indicating non-specific reduction of the azo bond. However, as the dye concentration increased, the percentage of decolorization decreased significantly, with the percentage of decolorization being 10 times lower at 500 mg AO7/L than at 50 mg/L. The maximum decolorization percentage and time required for dye decolorization clearly differed between the dyes tested (Fig. 5a), probably due to differences in molecular structure. More specifically, azo dyes (AO7; Acid Orange 7, MO; Methyl Orange, RB81; Reactive Blue 81, MR; Methyl Red, and SGR; Acid Brilliant Scarlet GR) with low molecular weight showed a higher decolorization percentage, as compared to azo dyes with complex structure and high molecular weight (RB5; Reactive Black 5, RR120; Reactive Red 120, RG19; Reactive Green 19, and RV5; Reactive Violet 5) (Fig. 5a).
The widespread use of azo dyes in the textile industry results in massive amounts of textile effluents, which are difficult to biodegrade. As a result, it is critical to screen and construct microbial consortia that are potentially suitable for the effective processing of azo dye-loaded industrial effluents. The NYC-1 consortium's efficiency in decolorizing azo dyes in simulated wastewater was evaluated in this study. Clearly, after decolorizing the simulated wastewater for 21 h, the derived peak between 330 and 580 nm completely disappeared (Additional file 1: Fig. S1). As a result, NYC-1 is a promising microbial consortium for effective decolorization of industrial wastewater containing various azo dyes. The efficacy of NYC-1 consortium on azo dye decolorization was compared to other individual and microbial consortia reported earlier (Table 5). The performance of the NYC-1 consortium on decolorization after repeated dye addition, on the other hand, was investigated. As shown in Fig. 5b, the time required to achieve maximum decolorization efficiency after the initial injection of AO7 (150 mg/L) was 9 and 6 h, respectively. At the same time, it had just been 3 h after repeated cycles of dye addition.
To further examine the efficacy of dye biodegradation, the effects of static and agitation conditions on the decolorization performance of AO7 by the MnP-producing oleaginous yeast consortium NYC-1 were investigated. Under static conditions, approximately complete AO7 decolorization was obtained by the NYC-1 consortium after cultivation for 6 h. In comparison, after 15 h of incubation, the decolorization efficiency of AO7 under agitation conditions (120 rpm) reached 49% (Fig. 6a). Furthermore, the effect of initial dye concentration on the AO7 decolorization efficacy was investigated by the NYC-1 consortium. Within 9 h, the medium supplemented with the model azo dye AO7 (initial 50 mg/L) was decolorized. However, further transfers demonstrated rapid decolorization, with more than 98% of AO7 decolorization acquired within 3 h from the original concentration of 50 mg/L. Even when the initial dye concentration was increased to 250 mg/L, the decolorization efficiency exceeded 92% after 18 h of incubation (Fig. 6b).
The NYC-1 consortium also investigated the influence of several parameters, such as temperature, pH, and salt concentration on the decolorization efficacy of AO7 (Fig. 7). Complete dye decolorization was accomplished after 18 h of incubation at 28 °C, but increasing temperature reduced decolorization efficiency, achieving over 85% decolorization after 24 h of incubation at 50 °C (Fig. 7a). On the contrary, at 5 °C, a significant decrease in decolorization efficiency was determined, with less than 10% decolorization . Thus, the NYC-1 consortium's decolorization effectiveness of AO7 varied significantly with temperature (p < 0.0001). Furthermore, as demonstrated in Fig. 7b, the effect of pH change varied. Within 18 h of incubation, the NYC-1 consortium showed complete decolorization of AO7 dye at pH 5. However, as the acidic or alkaline pH increased, the decolorization efficiency declined dramatically, achieving approximately 57 and 15% dye decolorization at pH 3 and 10, respectively. Therefore, the NYC-1 consortium's decolorization effectiveness of AO7 varied considerably with increasing acidity or alkalinity (p < 0.0001) (Fig. 7b).
In terms of salt concentration (NaCl up to 60 mg/L), the NYC-1 consortium investigated the decolorization efficiency of AO7 in the presence of high concentrations of NaCl (Fig. 7c). After 18 h of incubation, the dye decolorization efficiency exceeded 95% when the decolorization medium was supplemented with 10 g/L NaCl. However, increasing the salt concentration resulted in a considerable decline in the NYC-1 consortium's AO7 decolorization efficiency, which exceeded 65% when the concentration of NaCl was adjusted at 50 g/L. As a result, the NYC-1 consortium has been identified as a halotolerant consortium that could potentially be applied prior to the disposal of azo dye wastewater, which is also characterized by high salt concentrations (Fig. 7c).
The NYC-1 consortium's decolorization performance against AO7 in the presence of co-substrates (e.g., carbon, nitrogen, or agricultural wastes) was also investigated (Fig. 8). The addition of co-substrates is necessary to improve yeast growth and decolorization of the azo dye. Therefore, the dye decolorization performance of the NYC-1 consortium was investigated in the presence of various carbon (xylose, glucose, maltose, sucrose, starch) and nitrogen (peptone, urea, NaNO3, NH4Cl, yeast extract) sources at 0.5% (w/v) added to the Bushnell Haas synthetic medium. Furthermore, the effect of various agricultural wastes, such as rice straw and stalk, sorghum husk, wheat bran, and bagasse, on AO7 decolorization was investigated upon adding 0.5 mL extract of 0.5% boiled substrates. In the absence of any supplementation, NYC-1 showed the lowest decolorization activity (approximately 17%) in the synthetic medium. The presence of xylose resulted in the highest percentage of decolorization (98.25%) of the carbon sources tested. In comparison, no significant difference (p 0.417) in decolorization efficiency (94.18%) was observed in the presence of glucose. Within 24 h of incubation, the constructed consortium's decolorization efficiency for maltose, sucrose, and starch was 77.34, 69.71, and 53.41%, respectively (Fig. 8a).
In terms of nitrogen sources, yeast extract resulted in complete (100%) decolorization, whereas peptone resulted in 97.2% (Fig. 8b). Urea, NaNO3, and NH4Cl, on the other hand, inhibited AO7 decolorization by 44.32, 38.80, and 9.67%, respectively, within 24 h. Τhe decolorization efficiency of NYC-1 consortium against AO7 was also evaluated in the presence of agricultural wastes (Fig. 8c). Within 24 h of incubation, rice straw had the highest percentage of decolorization (92.33%), followed by sorghum husk (86.59%). Bagasse extract, on the other hand, showed the lowest decolorization efficiency (28.62%). Rice straw and sorghum husk had a significantly (p < 0.0001) greater effect than other agro-wastes used (wheat bran, rice stalk, and bagasse) (Fig. 8c). As a result, using lignocellulosic biomass to improve the decolorization process could be a promising approach while also contributing to the effective management of agro-wastes.
Influence of azo dye on the fatty acid composition and biodiesel properties
Textile wastewater bioremediation is a new dye treatment technology. Oleaginous yeasts are gaining popularity for their ability to degrade textile wastewater and accumulate fatty acids for biodiesel production. This could be the first study to explore the efficacy of MnP-producing oleaginous yeasts for coupling azo dye decolorization and biodiesel production. The lipids extracted from the AO7-degraded NYC-1 consortium were compared to those extracted from the control experiment (dye-free medium containing NYC-1 yeast consortium). Clearly, there was a decrease in the percentage of saturated fatty acids, particularly dodecanoic acid, as well as an increase in the amount of other alkenes and alkanes in the dye-treated oleaginous yeast consortium compared to the control one (data not shown). The main physicochemical properties of biodiesel produced by the azo dye AO7-degraded NYC-1 consortium were estimated and the results were compared to those obtained from international standards, vegetable oils, and oleaginous yeasts (Table 6). Determining the long-chain saturation factor (LCSF) is essential for estimating critical parameters, such as cetane number, oxidative stability, and kinematic viscosity. Under this scope, the values of LCSF, cetane number, oxidative stability, and kinematic viscosity of biodiesel produced by the AO7-degraded NYC-1 consortium were 4.21, 53, 7.85 h, and 4.38 mm2 s−1, respectively, while M. caribbica SSA1654 achieved biodiesel values of 3.34, 52.32, 8.3 h, and 3.98 mm2 s−1, respectively (Table 6). Furthermore, when compared to M. caribbica SSA1654 (95.85%), Rhodotorula glutinis R4 (95.2%), palm oil (64.2%), and olive oil (92.7%), the biodiesel produced by NYC-1 consortium had a degree of unsaturation of 96.12%.
The fatty acid profiles of four oleaginous yeast strains were determined: V. humicola SSA1514, M. caribbica SSA1654, M. guilliermondii SSA1547, and D. hansenii SSA1502, with only the profile of M. caribbica SSA1654 presented in this study (Additional file 1: Table S1). Meyerozyma caribbica SSA1654 had a high lipid content of 47.25 ± 1.84% (w/w) and a glucose and nitrogen assimilation rates of 40 g/L and 137.5 mg/L, respectively, resulting in an imbalanced metabolism and glucose conversion into neutral lipids. The profile of the extracted lipid of M. caribbica SSA1654 in terms of the main lipid classes, monoacylglycerol (MAG), diacylglycerol (DAG), TAG, and free fatty acids (FFA), was assessed qualitatively using thin layer chromatography (TLC). The results were compared with those obtained from the vegetable oil (palm oil) (Additional file 1: Table S1). TLC analysis revealed that the lipid bodies of M. caribbica SSA1654 are mostly TAG. Interestingly, the oleaginous yeast flow rate and composition were comparable to those of palm oil. The fatty acid profile of the oleaginous yeast M. caribbica SSA1654 was determined after it was grown at a C/N ratio of about 40. Long-chain fatty acids were abundant in the lipid generated under these conditions, particularly oleic acid (C18:1; 60.73% w/w) and palmitic acid (C16:0; 17.34% w/w). The prominent FAMEs were C18, which included stearic acid (C18:0), oleic acid (C18:1), linoleic acid (C18:2), and linolenic acid (C18:3), accounting for 76.22 percent of total FAMEs. On the other hand, total C16, including palmitic acid (C16:0) and palmitoleic acid (C16:1), accounted for 18.94% of total FAMEs (Additional file 1: Table S1). M. caribbica SSA1654 also synthesized more mono-unsaturated fatty acid (MUFA; MUFA; 62.33%) than saturated (SFA) and poly-unsaturated fatty acid (PUFA), 19.2% and 13.63%, respectively (Additional file 1: Table S1). Overall, the FAMEs profile, as well as the relative abundance of UFA and SFA, greatly influence biodiesel quality. As a result, M. caribbica SSA1654 is a good candidate for third-generation biodiesel production. The high MUFA is advantageous, because it improves the cold filter plugging point. High SFA, on the other hand, results in a high oxidative stability value. As a result, controlling the physicochemical properties of biodiesel with the optimal ratio of SFA to MUFA in FAMEs is critical. Furthermore, the produced biodiesel by M. caribbica SSA1654 and NYC-1 consortium had a C18:3 content (less than 12), which was in accordance with the international biodiesel standard EN 14214 (Table 6). Therefore, the MnP-producing oleaginous yeast consortium NYC-1 and the individual Mnp-producing oleaginous yeast strain M. caribbica SSA1654 could be novel biological candidates with promising biodiesel production potential.
The decolorization of textile dye wastewater and subsequent lipid production has been highly demanded by screening MnP-producing yeasts from WFTs. Within 120 h, nine of 38 yeast isolates demonstrated the highest peroxidase potential, up to 27 U/mL. Recently, MnP was patented for enzymatic hydrolysis of lignocellulosic substances , with ligninolytic enzymes, such as MnP, LiP, Lac, and dye-decolorizing peroxidase, which are commonly derived from various white-rot basidiomycetes and effectively used in the degradation of lignin and other xenobiotics . MnP-producing yeasts have been shown to be highly effective at converting textile azo dye wastewater into innocuous end products . Ligninolytic yeasts produce a variety of peroxidases, which are used in lignin degradation, dye decolorization, and phenolics removal . Biomass digestion by termites, on the other hand, is far more efficient, typically achieving over 95% within a day [37, 52]. Sterigmatomyces halophilus strain SSA-1575, S. halophilus strain SSA1511 , Barnettozyma californica strain SSA1518 , Yarrowia sp. strain SSA1642 , and Meyerozyma guilliermondii strain SSA1522  have all been tested for azo dye decolorization. However, in the current study, the efficacy of MnP-producing oleaginous yeasts isolated from WFTs for coupling azo dye decolorization and biodiesel production was first explored.
Nile red, a common hydrophobic red fluorescent dye with metachromatic properties, was applied to detect intracellular neutral lipids. In this context, seven MnP-producing yeast strains: C. stauntonica SSA1653, M. caribbica SSA1654, D. hansenii SSA1502, V. humicola SSA1514, M. guilliermondii SSA1547, S. halophilus SSA1655, and F. inositophila SSA1579, showed high yellow-gold coloration using Nile red staining. Between them, M. guilliermondii strain B1281A and M. caribbica strain DMKURK258 can be considered potential biodiesel production sources . Similarly, the oleaginous yeast D. hansenii isolated from kefir showed a high ability to accumulate neutral lipids , while C. humicola strain UCDFST 10–1004 has been previously reported to convert lignocellulose into neutral lipids, easily trans-esterified to biodiesel .
Out of the 1600 yeast species known, approximately 100 are oleaginous, with lipid content greater than 20% (w/w) in the form of neutral lipids, primarily TAG . Oleaginous yeasts are promising micro-organisms that can use a variety of carbon sources to synthesize lipids, which can then be used to produce biodiesel. In this study, the accumulation of TAG in the oleaginous yeast strains varied between 33.9 and 47.2%, since D. hansenii SSA1502 (33.96 ± 5.72%, w/w), V. humicola SSA1514 (43.46 ± 5.34%, w/w), M. guilliermondii SSA1547 (44.74 ± 6.09%, w/w), and M. caribbica SSA1654 (47.25 ± 1.84%, w/w) showed high lipid-producing ability. Compared to the oleaginous yeasts, the highest concentration of biomass (9.27 ± 0.25 to 11.23 ± 0.71 g/L), lipid production (0.96 ± 0.09 to 1.82 ± 0.61 g/L), lipid content (10.35 ± 1.07 to 10.41 ± 1.14%, w/w), YL/x (0.104 ± 0.009 to 0.173 ± 0.02 g/g), Yx/s (0.28 ± 0.05 to 0.39 ± 0.01 g/g), YL/s (0.05 ± 0.01 to 0.09 ± 0.01 g/g), Qx (0.08 ± 0.001 to 0.12 ± 0.005 g/L/h) and QL (0.005 ± 0.001 to 0.02 ± 0.006 g/L/h) were achieved by the non-oleaginous yeast strains F. inositophila SSA1579, C. stauntonica SSA1653, and S. halophilus SSA1655, while the oleaginous yeasts Cutaneotrichosporon curvatum and Cyberlindnera saturnus showed the highest lipid content of 36.9 and 33.9% (w/w), respectively . The lipid production of MnP-producing oleaginous yeast strains was compared to that of other oleaginous yeasts reported in the literature [7, 58,59,60,61].
In terms of nitrogen consumption, almost complete removal was observed within 24 h of growth, with residual glucose of approximately 15% and assimilation reaching up to 96%. Nitrogen limitation via the de novo synthesis pathway is thought to be the most effective . AMP-deaminase converts adenine-monophosphate (AMP) to inosine-monophosphate (IMP) during nitrogen deficiency. As a result, the production of α-ketoglutarate from isocitrate is impossible, and the citrate that enters the cytoplasm produces acetyl-Co-A. Finally, the fatty acid chains C14–C16 are formed . In contrast to oleaginous yeasts, conventional or non-oleaginous yeasts convert excess carbon into mannan and glucan when nitrogen is limited .
The CMCase and xylanase activities produced by M. caribbica SSA1654 were maximum at 0.17 ± 0.03 and 5.8 ± 0.9 U/mg on days 3 and 4, respectively, while the β-glucosidase activity produced by M. guilliermondii SSA1547 was maximum (0.061 ± 0.005 U/mg) on day 3. The lipase activity of D. hansenii SSA1502 ranged from 25.6 ± 3.2 U/mL to 55.3 ± 5.4 U/mL for M. caribbica SSA1654. Other oleaginous yeasts with similar CMCase, β-glucosidase, xylanase, and lipase activities have previously been reported, including Cryptococcus curvatus , Trichosporon mycotoxinivorans , Sterigmatomyces halophilus , Barnettozyma californica , and Yarrowia sp. . As a result, the simultaneous production of CMCase, β-glucosidase, xylanase, and lipase by the MnP-oleaginous yeast strains tested in this study may serve as a biocatalyst for biocatalyzing the valorizing of agro-industrial biomass/fatty wastes and subsequent biofuel production.
Not only fermentable sugars, but also a variety of byproducts are produced and discharged into the hydrolysates during the pretreatment and hydrolysis of lignocellulosic biomass. These compounds may have a deleterious impact on micro-organism growth, metabolism, and product production . The existence of inhibitory chemicals in lignocellulosic hydrolysates may be divided into three major categories depending on their origin: weak acids, phenolic compounds, and furan derivatives . However, the concentrations of these chemicals vary depending on the feedstock as well as the pretreatment and hydrolysis processes . As a result, five representative lignocellulose degradation compounds were chosen to investigate their inhibitory effects on growth of the four MnP-producing oleaginous yeast strains comprising the NYC-1 consortium (M. caribbica SSA1654, M. guilliermondii SSA1547, D. hansenii SSA1502, and V. humicola SSA1514): furfural from dehydration of pentose sugars; 5-HMF from dehydration of hexose sugars; vanillin from lignin breakdown during acid hydrolysis; acetic acid from de-acetylation of hemicellulose; and formic acid from furfural and 5-HMF breakdown . Among the four strains, M. caribbica SSA1654, showed the highest tolerance to furfural (1.0 g/L), 5-HMF (2.5 g/L), vanillin (2.0 g/L) and acetic acid (2.5 g/L). These findings demonstrated that tolerance to lignocellulose degradation inhibitors was affected by the nature and concentration of the compounds, as well as the micro-organisms .
Extensive research has been conducted on the effects of lignocellulose degradation inhibitors on ethanol fermentation, lipid accumulation, and oleaginous yeast growth, with the conclusion that lignocellulose degradation inhibitors had a greater impact on yeast growth than lipid accumulation . The uncoupling and intracellular anion accumulation hypotheses explain the inhibitory effects of weak acids. Strong acids are lipid-soluble and can diffuse across the plasma membrane, dissociating due to neutral cytosolic pH, lowering intracellular pH and reducing ATP levels (uncoupling theory) or inhibiting enzyme activity (intracellular anion accumulation theory) . Due to its higher acidity (pKa 3.75) and smaller molecular size (pKa 4.75), formic acid has a stronger inhibitory effect than acetic acid . Furfural and 5-HMF are major degradation products from xylose and glucose, respectively. They are major inhibitors of ethanol fermentation . These compounds inhibit the glycolytic enzymes hexokinase and glyceraldehyde-3-phosphate dehydrogenase . In this study, furfural had strong inhibitory effect on lipid accumulation of M. caribbica SSA1654. Enzyme matrices and selective barriers are affected by phenolic compounds, such as vanillin . According to Chandel et al. , higher molecular weight phenolic compounds are less harmful to micro-organisms. Our findings are in accordance with Zhao et al.  who found that lignocellulose degradation compounds are toxic in high concentrations.
In terms of dye decolorization, azo dyes with low molecular weight demonstrated a higher maximum decolorization percentage by the newly constructed MnP-producing oleaginous yeast consortium NYC-1 than azo dyes with complex structure and high molecular weight. The performance of NYC-1 consortium was compared to that of other consortia reported in the literature [8, 74,75,76]. It has been reported that simple structured dyes of low molecular weight show increased decolorization rates than complex structures and dyes of high molecular weight . Furthermore, the number of azo bonds, Van der Waals forces, hydrogen bonds, and hydrophobic/electrostatic interactions all have an effect on the rate of dye decolorization . However, after the fourth addition, the constructed consortium's AO7 decolorization performance was comparable. When nutrients are scarce, exponential yeast growth ceases, and microbial death occurs gradually. As a result, the yeast cell's enzyme system is gradually inhibited .
Under static conditions, AO7 decolorization by the NYC-1 consortium was significantly higher (p 0.0047) than under agitation conditions. The findings support the NYC-1 consortium's preference for static conditions for AO7 decolorization. Azoreductase, the enzyme that catalyzes azo dye decolorization, is inhibited by aeration due to competition between the azo group and oxygen (as an electron acceptor) in the oxidation of reduced electron carriers, such as NADH and quinones . The azo bond in azo dyes is known to be electron-withdrawing. As a result, it reduces the susceptibility of dye molecules to oxidation . In terms of increasing initial dye concentrations, similar results were obtained in the NYC-1 consortium's decolorization efficiency against AO7. At a dye concentration of 250 mg/L, the decolorization efficiency exceeded 92% in 18 h. Furthermore, there was no difference in the pH of the culture medium before and after AO7 decolorization, indicating that dye decolorization can be attributed to the biotic performance of the NYC-1 consortium rather than pH variation in the culture medium . Overall, the presence of azo dyes in aquatic bodies and effluents, ranging from 10 to 200 mg/L, is hazardous and aesthetically unappealing [81, 82]; thus, azo dye degradation is critical prior to discharge.
The effects of temperature, pH, and salt concentration on dye decolorization performance were also investigated by the NYC-1 consortium. The developed consortium demonstrated significantly higher decolorization efficiency at 28 °C than at 20 °C (p < 0.0001) and 30 °C (p 0.0065). In addition, at 5 °C, the decolorization efficiency dropped sharply to less than 10%, confirming previous findings of dye decolorization inhibition at low temperatures . Furthermore, the NYC-1 consortium retained its decolorizing efficiency at high temperatures (up to 50 °C) in the current study, demonstrating its valuable potential for azo dye bioremediation. The NYC-1 consortium also achieved complete AO7 decolorization at pH 5 within 18 h of incubation, whereas decolorization efficiency decreased significantly with increasing acidity (pH 3; 57% decolorization) or alkalinity (pH 10; 15% decolorization). The NYC-1 consortium demonstrated significantly higher decolorization efficiency at pH 5 than at pH 4 (p 0.0009) and pH 6 (p 0.0123). The low dye decolorization efficiency at higher acidic or alkaline pH values was most likely due to a change in the dye chemical structure caused by the formation of protonated azo dyes . Furthermore, after 18 h of incubation, the dye decolorization efficiency exceeded 95% in the presence of 10 g/L NaCl. However, increasing the salt concentration resulted in a significant decrease in decolorization efficiency (p < 0.0001), with 50% decolorization achieved at 60 g/L NaCl. As a result, the NYC-1 consortium represents a highly promising halo-tolerant yeast consortium with significant potential for bioremediation of textile wastewater containing azo dyes and high salt concentrations .
It has been reported that the addition of co-substrates (e.g., carbon, nitrogen, or agricultural wastes) is required for both yeast growth and improving azo dye decolorization performance . Decolorization performance took its highest value in the case of xylose (98.25%) with a non-significant difference with glucose (94.18%; p 0.417). However, both xylose and glucose revealed significant differences between maltose (p 0.0048) and sucrose (p 0.0015). In addition, xylose, glucose, maltose, and sucrose showed a significantly higher decolorization performance (p 0.0001) when compared with starch. Although the isolated bacteria's conversion of glucose to organic acids may inhibit textile dye decolorization , the presence of glucose or xylose can aid AO7 dye decolorization by the NYC-1 consortium. Due to the microbial preference for these added carbon sources, Saratale et al.  found that adding di- and polysaccharides is less effective in promoting decolorization efficiency than monosaccharides. On the other hand, organic and inorganic nitrogen sources are essential nutrients for yeast growth . The NYC-1 consortium determined that yeast extract (100%) and peptone (97.2%) are the best nitrogen sources for AO7 decolorization, because both yeast extract and peptone provide significantly higher decolorization efficiency (p < 0.0001) than other nitrogen sources tested (urea, NaNO3, and NH4Cl). It has been reported that peptone and yeast extract are responsible for activating NADH expression and thus effectively decolorizing the dye . The low decolorization efficiency in the presence of NH4Cl can be attributed to its inhibitory effect on the performance of the decolorization enzymes . Finally, the NYC-1 consortium investigated the decolorization efficiency of AO7 in the presence of agro-waste extracts. Rice straw demonstrated the highest decolorization performance (92.33%) compared to bagasse extract (28.62%), owing to the production of volatile organic acids or alcohols that could serve as electron donors for dye reduction. As a result, the addition of agro-wastes to improve AO7 decomposition is an environmentally friendly and low-cost process that addresses the issue of vast quantities of agro-wastes produced globally.
TLC is a tool for monitoring the conversion of extracted lipids to FAMEs during the transesterification reaction in the production of biodiesel . According to the TLC results obtained in this study, both M. caribbica SSA1654 and palm oil standard had a similar flow rate and composition. Furthermore, three types of neutral lipids, MAG, DAG, and TAG, have been identified in oleaginous yeasts, with TAG remaining the most abundant . Therefore, the TAG produced by M. caribbica SSA1654 could be valuable when aiming at biodiesel production. In contrast, Patel et al.  reported that lipid-producing yeasts in the presence of glucose and under nitrogen limitation mainly synthesize myristic acid (C14:0), palmitic acid (C16:0), stearic acid (C18:0), oleic acid (C18:1), linoleic acid (C18:2), and linolenic acid (C18:3). Furthermore, the total SFA, MUFA, and PUFA exceed 95% of the total FAMEs, similar to vegetable oils mainly comprising of C16 and C18 fatty acids (SFA or MUFA) [43, 93, 94], suggesting that M. caribbica SSA1654 could be a potential oleaginous yeast, isolated from WFTs, valued for biodiesel production.
The physicochemical properties of biodiesel derived from NYC-1 consortium and M. caribbica SSA1654 lipids were compared to those obtained from oleaginous yeasts  and vegetable oils . Increased LCSF has been shown to have a negative effect on the cold filter plugging point (cold flow behavior) of biodiesel . The crystallization of biodiesel in the engine pipeline under cold conditions is a significant challenge . The degree of unsaturation of fatty acids determines oxidative stability, which is another critical criterion used to determine the shelf-life of any fuel . The cetane number of the biodiesel produced by M. caribbica SSA1654 and the NYC-1 consortium met the EN 14,214 (minimum 51) and ASTM 6751–3 values (minimum 47). According to Hoekman et al. , there are two limit values for cetane numbers. The higher value reduces engine efficiency, while low cetane number values result in high hydrocarbon emissions.
Lignocellulose biomass and textile azo dyes are recalcitrant substrates that pose a challenge for microbial decomposition. Several microorganisms, on the other hand, exhibit remarkable enzymatic activity and have a high potential for wastewater treatment and simultaneous bioenergy production. V. humicola SSA1514, M. caribbica SSA1654, M. guilliermondii SSA1547, and D. hansenii SSA1502 were the most advantageous lipid-accumulating strains, exhibiting MnP, cellulase, xylanase, and lipase activities, as well as tolerance to common lignocellulose degradation inhibitors. Among the four strains, M. caribbica SSA1654, showed the highest tolerance to furfural, 5-HMF, vanillin and acetic acid. Furfural and formic acid had a significant inhibitory effect on biomass production and lipid accumulation by M. caribbica SSA1654, compared to the other lignocellulose degradation inhibitors tested, which had a minor effect. The decolorization of AO7 azo dye by the newly constructed MnP-producing oleaginous yeast consortium NYC-1 was investigated, and the results showed nearly complete decolorization, particularly in the presence of xylose and yeast extract, which were used as carbon and nitrogen sources, respectively. Simultaneously, rice straw supplementation resulted in 92.3% AO7 decolorization. The produced biodiesel by M. caribbica SSA1654 and NYC-1 consortium had a C18:3 content, which was in accordance with the international biodiesel standards. Overall, the enzymatic performance of yeasts and their high lipid content indicate that they have great potential for the effective valorization of lignocellulosic waste and industrial wastewater containing azo dyes, as well as the production of biodiesel. However, more research is required before a large-scale and environmentally friendly application can be established.
Dyestuff and agricultural wastes
Nine azo dyes (Sigma-Aldrich, St. Louis, USA) were used in this study. The names of these dyes are Reactive Green 19, Methyl Red, Methyl Orange, Reactive Blue 81, Reactive Red 120, Reactive Violet 5, Acid Orange 7, Acid Brilliant Scarlet GR, and Reactive Black 5. The chemical structure of dyes and their corresponding maximum wavelengths (λmax) are given in Fig. 9. In this work, several agricultural wastes (rice straw, bagasse, wheat bran, rice stalk, and sorghum husk) were obtained from local farmers and industries (Zhenjiang, China) to evaluate their impacts on the AO7 dye decolorization by the developed yeast consortium. The agricultural waste extracts were prepared in the Bushnell Haas medium (5.0 mL extract of 0.5% boiled agricultural residue).
Isolation and screening of oleaginous yeasts capable of azo dye decolorization
The experimental setup for the isolation and screening of MnP-producing oleaginous yeasts intended for azo dye decolorization and biodiesel production is depicted in Fig. 10. The WFTs, R. chinenesis and C. formosanus were collected from rotting wood trees at Wuhan and Nanjing, China as reported previously [32, 47]. Isolation of yeasts from the gut symbionts of R. chinenesis and C. formosanus was performed following Suh and Blackwell . The surface of insect samples was disinfected twice with 70% ethanol for 1 min prior to dissection. The insect guts were then eliminated aseptically and homogenized in a sterile saline solution (0.85% NaCl, w/v) . Isolation experiments were conducted in conical flasks (100 mL) with a working volume of 40 mL of the Bushnell Hass medium using aliquots of 500 µL crushed WFT gut solutions . This synthetic medium contained (g/L): 2.45 NaH2PO4, 6.8 KH2PO4, 1.72 MgSO4.7H2O, 0.067 MnSO4.7H2O, and 0.2 CaCl2.2H2O. The medium was also supplemented with 0.5 g/L o-Dianisidine dihydrochloride (as a precursor of many azo dyes as well as a peroxidase substrate) and glucose at a final concentration of 40 g/L. Yeast extract (3.0 g/L) was included as the sole nitrogen source and yielding a C/N ratio of around 40. As antibacterial and antifungal agents, 0.01% chloramphenicol and 0.02% sodium propionate were added to the medium. The flasks were then incubated for 3–10 days at 28 °C under shaking conditions of 150 rpm. Repeated sub-culturing on yeast extract–peptone–dextrose (YEPD) medium was used to isolate distinct yeast colonies with different morphotypes. The YEPD medium contained (g/L): 20 peptone, 10 dextrose, 10 yeast extract, and 20 agar. Inoculum preparation of purified yeast isolates was prepared as shown in Fig. 10.
The ability of yeast isolates to accumulate TAG was qualitatively assessed using Sudan Black B staining protocol  by incubating the isolates for 30 min, and observing blue color retention after rinsing with 70% ethanol. For TAG (lipid) accumulation among oleaginous and non-oleaginous yeasts, the selected isolates were examined under a fluorescence microscope (Olympus BX35) at 460–500 nm following the Nile red staining protocol . The most promising oleaginous yeasts found to be positive for Sudan Black B were tested for lipase activity on the agar plates of tributyrin medium (10 g/L tributyrin, 3 g/L yeast extract, 5 g/L, and 20 g/L agar) as per our earlier report . The lipid content of yeast isolates was then measured gravimetrically . More specifically, after cultivation in a lipid production medium at 28 °C for 5 days, the produced yeast biomass was harvested by centrifugation at 12,000 rpm for 10 min. The derived pellet was rinsed three times with distilled water, followed by drying at 105 °C, until constant weight. Quantification of lipids in cell biomass was performed following the method described by Vyas and Chhabra . The lipid content was calculated as a percentage of the lipid weight relative to the dry biomass weight (% w/w). Concurrently, the discharged supernatant was used for glucose and nitrogen determination throughout the cultivation period. The 3,5-Dinitrosalicylic acid method was used to determine residual glucose , while nitrogen assimilation was estimated to assess ammonia concentration in the cultivation medium .
The selected yeast isolates were tested for MnP production using phenol red as an indicator of ligninolytic activity, with the color of the phenol red changing from deep orange to light yellow . The isolates that showed positive phenol red results were later confirmed for peroxidase production by measuring the development of a reddish-brown color around yeast colonies grown on YEPD agar plates supplemented with the peroxidase substrate (o-Dianisidine dihydrochloride, 0.5 g/L) upon the addition of 0.4 mM H2O2. Subsequently, the selected isolates were incubated in Bushnell Hass medium containing 0.5 g/L o-Dianisidine dihydrochloride and 150 µM of MnSO4.4H2O as an inducer to stimulate MnP production. MnP activity in a liquid medium was also determined following Orth et al. . The MnP-producing oleaginous yeasts were then tested for dye decolorization using Acid Orange 7 (AO7) as a model azo dye.
Azo dye decolorization and enrichment
Decolorization experiments were carried out in Bushnell Hass broth medium amended with AO7 dye at an initial concentration of 50 mg/L and 150 µM of MnSO4.4H2O as an inducer to stimulate MnP production. Aliquots (10%, v/v) of MnP-producing oleaginous yeast cultures (OD600 of 0.2) were inoculated into flasks, which were incubated at 28 °C under static conditions. When the decolorization was realized, a 10 mL mixed culture sample was inoculated into a new dye-added medium for another round of enrichment. The procedures were repeated ten times until the decolorization efficiency was stable. An inoculated dye-free medium was used as an abiotic control. The dye decolorization ability of the enriched isolates was also evaluated on agar plates containing 50 mg dye/L. The fastest-growing MnP-producing oleaginous yeast colonies capable of azo dye decolorization and exhibiting a high ratio of a zone of decolorization to a colony diameter were selected for identification and further experiments. In addition, the performance of the constructed yeast consortium on the decolorization of various textile azo dyes was studied.
Dye decolorization was mointored quantitatively at its corresponding maximum wavelength (Fig. 9) using a UV–Vis spectrophotometer [81, 82]. The culture supernatant was centrifuged (10,000 xg, 10 min, 4 °C) and the percentage of decolorization (%D) was calculated following the formula given by Ali et al. . All assays were carried out in triplicate, with the average values used in calculations. %D = (A0 − At)/A0 ∗ 100, where A0 and At are the absorbances of dye solutions before and after decolorization, respectively.
The simulated wastewater was prepared according to the real textile dyeing effluent, including mixture of nine azo dyes (Fig. 9) at a concentration of 150 mg/L for each dye. The prepared medium was then supplemented with the synthetic YME substrate to obtain a final concentration of 250 mg/L. The performance of the constructed yeast consortium on the decolorization of the simulated wastewater evaluated using UV–Vis spectrophotometric analysis in the range of 400–800 nm.
Molecular identification and phylogenetic analysis
The genomic DNA of the selected MnP-producing oleaginous yeast isolates capable of azo dye decolorization was extracted using Dr. GenTLER High Recovery (TakaRa, Japan) according to the manufacturer's instructions. For the D1/D2 and ITS regions, the isolated DNA was amplified with NL1/NL4 and ITS1/ITS4 primers, respectively . The PCR amplification was performed as previously described  and the amplification products were sequenced at Sangon Biotech (Shanghai, China). The yeast strains were identified using the nucleotide BLAST (http://www.ncbi.nlm.nih.gov/BLAST/) database. Molecular evolutionary genetics analysis version 7.0 (MEGA 7.0) software was used for phylogenetic and evolutionary analyses .
Construction of MnP-producing oleaginous yeast consortium capable of azo dye decolorization
The NYC-1 consortium stands for molecularly identified MnP-producing oleaginous yeast species caribbica, hansenii, guilliermondii, and humicola were developed. A loopful of each yeast strain was cultured separately in screw cap tubes containing 5 mL YEPD broth and incubated at 28 °C for up to 48 h for inoculum preparation. The obtained culture biomass (1.0 g wet weight) was washed three times with phosphate buffer (pH 7.4) before being aseptically inoculated into 100 mL of the same buffer (50 mM) and gently mixed to prepare a homogeneous cell suspension. Finally, the NYC-1 consortium was developed by mixing the cell suspensions at a 1:1 rate.
Activities of MnP, lipase, β-glucosidase, CMCase, and xylanase enzymes produced by the individual strains (SSA1547, SSA1514, SSA1654, and SSA1502) consisting the NYC-1 yeast consortium were determined spectrophotometrically in cell-free extracts. The MnP activity of yeast strains was determined following the method described by Rekik et al. . After adding H2O2 to the reaction mixture, the increase in the absorbance at 510 nm was monitored for 2 min (30 s interval) at 40 °C. Lipase activity was determined using p-nitrophenyl palmitate as a substrate . On the basis of the 3,5-Dinitrosalicylic acid technique , the activities of endo-β-1,4-glucanase (CMCase) and xylanase were assessed using CMC and xylan as corresponding substrates, respectively . β-glucosidase enzymatic activity was measured using 5 mM p-nitrophenyl glucopyranoside (pH 5) as the corresponding substrate . One unit of enzyme activity was defined as the amount of enzyme required to release 1.0 μmol of the reaction product per minute under specified conditions.
Effect of lignocellulose degradation inhibitors on yeast growth and lipid accumulation
The effect of the predominant lignocellulose degradation inhibitors (furfural, 5-hydroxymethyl furfural, acetic acid, vanillin, and formic acid) as common inhibitors in lignocellulosic hydrolysates was studied on growth and lipid accumulation. The effect of these inhibitors on growth of the selected individual strains (SSA1547, SSA1514, SSA1654, and SSA1502) consisting the NYC-1 yeast consortium was investigated as described previously [23, 55]. The consortium members' ability to grow in the presence of furfural (0.5, 1.0 and 1.5 g/L), 5-hydroxymethyl furfural (0.5, 1.0, 2.0, and 2.5 g/L), acetic acid (0.5, 1.0, 1.5, and 2.5 g/L), vanillin (0.5, 1.0, 2.0, and 2.5 g/L), and formic acid (0.5, 1.0, 3.0, and 4.0 g/L), which are commonly found in lignocellulosic hydrolysate [55, 69, 105], was tested in Erlenmeyer flasks with yeast nitrogen base medium supplemented with 1% glucose  against the control (medium without addition of any inhibitor compound). An aliquot (20 µL) of each yeast cell suspension (108 cells/mL) was inoculated in the prepared sterilized media. Growth was estimated within 14 days of incubation in a rotary shaker at 28 °C with an agitation speed of 150 rpm. Growth was defined following Poontawee et al. .
The effect of lignocellulose degradation inhibitors on lipid accumulation by the selected individual strains consisting the NYC-1 consortium was also studied. The prepared inoculum of yeast cultures was inoculated in 50 mL of nitrogen base medium broth supplemented with 1% glucose and in the presence of various concentrations of lignocellulose degradation inhibitors, giving an initial cell concentration of 1.0 at OD600. The inoculated flasks were incubated in a rotary shaker at 28 °C with an agitation speed of 150 rpm for 7 days. Cells were subsequently harvested for determining biomass and lipid accumulation. In this experiment, various concentrations of lignocellulose degradation inhibitors were used, including furfural (0.05, 0.1, 0.2 and 0.5 g/L), 5-hydroxymethyl furfural (0.1, 0.5, 1.0, and 5.0 g/L), acetic acid (0.1, 0.5, and 1.0 g/L), vanillin (0.1, 0.5, and 1.0 g/L), and formic acid (0.1, 0.2, 0.3, and 0.5 g/L). The control was a medium without the addition of any inhibitor compound.
Lipid extraction and biodiesel properties
TLC with a K6 silica gel plate (Merck, India) and a solvent mixture of hexane: diethyl-ether: methanol: acetic acid (78:17:3:2, v/v) were employed. Lipid analysis was detected following exposure to iodine vapor . The microbial lipid generated after methanolysis was examined following Morrison and Smith . The FAME obtained by transesterification was further analyzed as per our earlier report . The main qualitative physicochemical characteristics of the biodiesel produced by the lipid-derived from Meyerozyma caribbica and the NYC-1 consortium were determined, including cetane number, kinematic viscosity, iodine, saponification value, oxidative stability, density, long-chain saturation factor, and unsaturation degree . Subsequently, a comparison with palm oil  and the oleaginous yeast Rhodotorula glutinis R4  was made, followed by an emphasis on the specifications described in the international biodiesel standards, EN 14214 (Europe) and ASTM D6751-3 (USA).
All experiments were carried out in triplicate, and the results were analyzed using NCSS 2020 (LIC, Utah, USA) and Minitab version 19.2020.1 (Minitab Inc., US). The values represent the mean of three independent replicates, with error bars indicating the standard deviation. In terms of statistical significance evaluation, one-way analysis of variance (ANOVA) with Tukey–Kramer multiple comparisons and t-Student's tests, applied at p value ≤ 0.05.
Availability of data and materials
The data sets used and/or analyzed during the current study are available from the corresponding author on reasonable request.
Ali SS, Haixin J, Mustafa AM, Koutra E, El-Sapagh S, Kornaros M, Elsamahy T, Khalil M, Bulgariu L, Sun J. Construction of a novel microbial consortium valued for the effective degradation and detoxification of creosote-treated sawdust along with enhanced methane production. J Hazard Mater. 2021;27:126091.
Ali SS, Mustafa AM, Kornaros M, Sun J, Khalil M, El-Shetehy M. Biodegradation of creosote-treated wood by two novel constructed microbial consortia for the enhancement of methane production. Bioresour Technol. 2021;323:124544.
Ali SS, Mustafa AM, Sun J. Wood-feeding termites as an obscure yet promising source of bacteria for biodegradation and detoxification of creosote-treated wood along with methane production enhancement. Bioresour Technol. 2021;338:125521.
Ali SS, Kornaros M, Manni A, Sun J, El-Shanshoury AE, Kenawy ER, Khalil MA. Enhanced anaerobic digestion performance by two artificially constructed microbial consortia capable of woody biomass degradation and chlorophenols detoxification. J Hazard Mater. 2020;389:122076.
Ali SS, Sun J. Effective thermal pretreatment of water hyacinth (Eichhornia crassipes) for the enhancement of biomethanation: VIT® gene probe technology for microbial community analysis with special reference to methanogenic Archaea. J Environ Chem Engin. 2019;7(1):102853.
Ali SS, Sun J. Physico-chemical pretreatment and fungal biotreatment for park wastes and cattle dung for biogas production. Springerplus. 2015;4(1):712.
Ali SS, Al-Tohamy R, Manni A, Luz FC, Elsamahy T, Sun J. Enhanced digestion of bio-pretreated sawdust using a novel bacterial consortium: microbial community structure and methane-producing pathways. Fuel. 2019;254:115604.
Ali SS, Nessem AA, Sun J, Li X. The effects of water hyacinth pretreated digestate on Lupinus termis L. seedlings under salinity stress: a complementary study. J Environ Chem Engin. 2019;7(3):103159.
Shen X, Geng C, Lv B, Xu W, Xu Y, Zhao H. Tire pyrolysis wastewater treatment by a combined process of coagulation detoxification and biodegradation. Environ Sci Ecotechnol. 2021;7:100129.
Wang Y, Sun J, Ali SS, Gao L, Ni X, Li X, Wu Y, Jiang J. Identification and expression analysis of Sorghum bicolor gibberellin oxidase genes with varied gibberellin levels involved in regulation of stem biomass. Ind Crops Prod. 2020;145:111951.
Gumisiriza R, Hawumba JF, Okure M, Hensel O. Biomass waste-to-energy valorisation technologies: a review case for banana processing in Uganda. Biotechnol Biofuels. 2017;10:11.
Morales GM, Ali SS, Si H, Zhang W, Zhang R, Hosseini K, Sun J, Zhu D. Acidic versus alkaline bacterial degradation of lignin through engineered strain E. coli BL21 (Lacc): exploring the differences in chemical structure, morphology, and degradation products. Front bioeng biotechnol. 2020;8:671.
Weng C, Peng X, Han Y. Depolymerization and conversion of lignin to value-added bioproducts by microbial and enzymatic catalysis. Biotechnol Biofuels. 2021;14:84.
Koutra E, Mastropetros SG, Ali SS, Tsigkou K, Kornaros M. Assessing the potential of Chlorella vulgaris for valorization of liquid digestates from agro-industrial and municipal organic wastes in a biorefinery approach. J Clean Prod. 2021;280:124352.
Ali SS, Elsamahy T, Koutra E, Kornaros M, El-Sheekh M, Abdelkarim E, Zhu D, Sun J. Degradation of conventional plastic wastes in the environment. A review on current status of knowledge and future perspectives of disposal. Sci Total Environ. 2021;771:144719.
Ali SS, Elsamahy T, Al-Tohamy R, Zhu D, Mahmoud Y, Koutra E, Metwally MA, Kornaros M, Sun J. Plastic wastes biodegradation: mechanisms, challenges and future prospects. Sci Total Environ. 2021;780:146590.
Ali SS, Al-Tohamy R, Koutra E, Moawad MS, Kornaros M, Mustafa AM, Mahmoud YA, Badr A, Osman ME, Elsamahy T, Jiao H. Nanobiotechnological advancements in agriculture and food industry: applications, nanotoxicity, and future perspectives. Sci Total Environ. 2021;792:148359.
Fernández-Fueyo E, Ruiz-Dueñas FJ, López-Lucendo MF, Pérez-Boada M, Rencoret J, Gutiérrez A, Pisabarro AG, Ramírez L, Martínez AT. A secretomic view of woody and nonwoody lignocellulose degradation by Pleurotus ostreatus. Biotechnol Biofuels. 2016;9:49.
Zhu N, Liu J, Yang J, Lin Y, Yang Y, Ji L, Li M, Yuan H. Comparative analysis of the secretomes of Schizophyllum commune and other wood-decay basidiomycetes during solid-state fermentation reveals its unique lignocellulose-degrading enzyme system. Biotechnol Biofuels. 2016;9:42.
Fernández-Fueyo E, Ruiz-Dueñas FJ, Martínez MJ, Romero A, Hammel KE, Medrano FJ, Martínez AT. Ligninolytic peroxidase genes in the oyster mushroom genome: heterologous expression, molecular structure, catalytic and stability properties, and lignin-degrading ability. Biotechnol Biofuels. 2014;7:2.
Qin X, Sun X, Huang H, Bai Y, Wang Y, Luo H, Yao B, Zhang X, Su X. Oxidation of a non-phenolic lignin model compound by two Irpex lacteus manganese peroxidases: evidence for implication of carboxylate and radicals. Biotechnol Biofuels. 2017;10:103.
Liu Y, Wu Y, Zhang Y, Yang X, Yang E, Xu H, Yang Q, Chagan I, Cui X, Chen W, Yan J. Lignin degradation potential and draft genome sequence of Trametes trogii S0301. Biotechnol Biofuels. 2019;12:256.
Ali SS, Al-Tohamy R, Koutra E, Kornaros M, Khalil M, Elsamahy T, El-Shetehy M, Sun J. Coupling azo dye degradation and biodiesel production by manganese-dependent peroxidase producing oleaginous yeasts isolated from wood-feeding termite gut symbionts. Biotechnol Biofuels. 2021;14(1):61.
Ali SS, Al-Tohamy R, Sun J. Performance of Meyerozyma caribbica as a novel manganese peroxidase-producing yeast inhabiting wood-feeding termite gut symbionts for azo dye decolorization and detoxification. Sci Total Environ. 2022;806:150665.
Ali SS, Abomohra AE, Sun J. Effective bio-pretreatment of sawdust waste with a novel microbial consortium for enhanced biomethanation. Bioresour Technol. 2017;238:425–32.
Ali SS, Mustafa AM, Kornaros M, Manni A, Sun J, Khalil MA. Construction of novel microbial consortia CS-5 and BC-4 valued for the degradation of catalpa sawdust and chlorophenols simultaneously with enhancing methane production. Bioresour Technol. 2020;301:122720.
Murillo G, He Y, Yan Y, Sun J, Bartocci P, Ali SS, Fantozzi F. Scaled-up biodiesel synthesis from Chinese Tallow Kernel oil catalyzed by Burkholderia cepacia lipase through ultrasonic assisted technology: a non-edible and alternative source of bio energy. Ultrason Sonochem. 2019;58:104658.
Murillo G, Ali SS, Sun J, Yan Y, Bartocci P, El-Zawawy N, Azab M, He Y, Fantozzi F. Ultrasonic emulsification assisted immobilized Burkholderia cepacia lipase catalyzed transesterification of soybean oil for biodiesel production in a novel reactor design. Renew Energy. 2019;135:1025–34.
Darwesh OM, Ali SS, Matter IA, Elsamahy T. Nanotextiles waste management: Controlling of release and remediation of wastes. In: Nanosensors and Nanodevices for Smart Multifunctional Textiles 2021, pp. 267–286. Elsevier. https://doi.org/10.1016/B978-0-12-820777-2.00016-9
Bhat AP, Gogate PR. Degradation of nitrogen-containing hazardous compounds using advanced oxidation processes: a review on aliphatic and aromatic amines, dyes, and pesticides. J Hazard Mater. 2021;403:123657.
Jin X, Wu C, Tian X, Wang P, Zhou Y, Zuo J. A magnetic-void-porous MnFe2O4/carbon microspheres nano-catalyst for catalytic ozonation: Preparation, performance and mechanism. Environ Sci Ecotechnol. 2021;7:100110.
Ali SS, Al-Tohamy R, Sun J, Wu J, Huizi L. Screening and construction of a novel microbial consortium SSA-6 enriched from the gut symbionts of wood-feeding termite, Coptotermes formosanus and its biomass-based biorefineries. Fuel. 2019;236:1128–45.
Li L, Wang T, Chen T, Huang W, Zhang Y, Jia R, He C. Revealing two important tryptophan residues with completely different roles in a dye-decolorizing peroxidase from Irpex lacteus F17. Biotechnol Biofuels. 2021;14:128.
Ali SS, Al-Tohamy R, Xie R, El-Sheekh MM, Sun J. Construction of a new lipase-and xylanase-producing oleaginous yeast consortium capable of reactive azo dye degradation and detoxification. Bioresour Technol. 2020;313:123631.
Ali SS, Al-Tohamy R, Koutra E, El-Naggar AH, Kornaros M, Sun J. Valorizing lignin-like dyes and textile dyeing wastewater by a newly constructed lipid-producing and lignin modifying oleaginous yeast consortium valued for biodiesel and bioremediation. J Hazard Mater. 2021;403:123575.
Ali SS, Sun J, Koutra E, El-Zawawy N, Elsamahy T, El-Shetehy M. Construction of a novel cold-adapted oleaginous yeast consortium valued for textile azo dye wastewater processing and biorefinery. Fuel. 2021;285:119050.
Ali SS, Al-Tohamy R, Sun J, Wu J, Huang M. The role of gut symbionts from termites: a unique hidden player from yeasts. Acta Microbiol Sin. 2018;58(6):1004–15.
Ferrero GO, Faba EM, Eimer GA. Biodiesel production from alternative raw materials using a heterogeneous low ordered biosilicified enzyme as biocatalyst. Biotechnol Biofuels. 2021;14:67.
Zhou ZW, Xing X, Li J, Hu ZE, Xie ZB, Wang N, Yu XQ. Magnetic COFs as satisfied support for lipase immobilization and recovery to effectively achieve the production of biodiesel by great maintenance of enzyme activity. Biotechnol Biofuels. 2021;14:156.
Karamerou EE, Parsons S, McManus MC, Chuck CJ. Using techno-economic modelling to determine the minimum cost possible for a microbial palm oil substitute. Biotechnol Biofuels. 2021;14:57.
Pagliano G, Ventorino V, Panico A, Pepe O. Integrated systems for biopolymers and bioenergy production from organic waste and by-products: a review of microbial processes. Biotechnol Biofuels. 2017;10:113.
Yaguchi A, Franaszek N, O’Neill K, Lee S, Sitepu I, Boundy-Mills K, Blenner M. Identification of oleaginous yeasts that metabolize aromatic compounds. J Ind Microbiol Biotechnol. 2020;47:801–13.
Ayadi I, Belghith H, Gargouri A, Guerfali M. Screening of new oleaginous yeasts for single cell oil production, hydrolytic potential exploitation and agro-industrial by-products valorization. Process Saf Environ Prot. 2018;119:104–14.
Slininger PJ, Dien BS, Kurtzman CP, Moser BR, Bakota EL, Thompson SR, O’Bryan PJ, Cotta MA, Balan V, Jin M, Sousa LD. Comparative lipid production by oleaginous yeasts in hydrolyzates of lignocellulosic biomass and process strategy for high titers. Biotechnol Bioeng. 2016;113:1676–90.
Xu Z, Lei P, Zhai R, Wen Z, Jin M. Recent advances in lignin valorization with bacterial cultures: microorganisms, metabolic pathways, and bio-products. Biotechnol Biofuels. 2019;12:32.
Vyas S, Chhabra M. Isolation, identification and characterization of Cystobasidium oligophagum JRC1: a cellulase and lipase producing oleaginous yeast. Bioresour Technol. 2017;223:250–8.
Ali SS, Wu J, Xie R, Zhou F, Sun J, Huang M. Screening and characterizing of xylanolytic and xylose-fermenting yeasts isolated from the wood-feeding termite, Reticulitermes chinensis. PLoS ONE. 2017;12(7):e0181141.
Al-Dhabi NA, Esmail GA, Arasu MV. Effective degradation of tetracycline by manganese peroxidase producing Bacillus velezensis strain Al-Dhabi 140 from Saudi Arabia using fibrous-bed reactor. Chemosphere. 2021;268:128726.
Martinez D, Larrondo LF, Putnam N, Gelpke MD, Huang K, Chapman J, Helfenbein KG, Ramaiya P, Detter JC, Larimer F, Coutinho PM. Genome sequence of the lignocellulose degrading fungus Phanerochaete chrysosporium strain RP78. Nat Biotechnol. 2004;22(6):695–700.
Baborová P, Möder M, Baldrian P, Cajthamlová K, Cajthaml T. Purification of a new manganese peroxidase of the white-rot fungus Irpex lacteus, and degradation of polycyclic aromatic hydrocarbons by the enzyme. Res Microbiol. 2006;157:248–53.
Debnath R, Saha T. An insight into the production strategies and applications of the ligninolytic enzyme laccase from bacteria and fungi. Biocatal Agric Biotechnol. 2020;12:101645.
Sun JZ, Ding SY, Peterson DJ. Biological conversion of biomass for fuels and chemicals: Explorations from natural utilization systems. Cambridge: Royal Society of Chemistry; 2014.
Ramírez-Castrillón M, Jaramillo-Garcia VP, Rosa PD, Landell MF, Vu D, Fabricio MF, Ayub MA, Robert V, Henriques JA, Valente P. The oleaginous yeast Meyerozyma guilliermondii BI281A as a new potential biodiesel feedstock: Selection and lipid production optimization. Front Microbiol. 2017;8:1776.
Gientka I, Kieliszek M, Jermacz K, Błażejak S. Identification and characterization of oleaginous yeast isolated from kefir and its ability to accumulate intracellular fats in deproteinated potato wastewater with different carbon sources. BioMed Res Int. 2017. https://doi.org/10.1155/2017/6061042.
Sitepu IR, Jin M, Fernandez JE, da Costa SL, Balan V, Boundy-Mills KL. Identification of oleaginous yeast strains able to accumulate high intracellular lipids when cultivated in alkaline pretreated corn stover. Appl Microbiol Biotechnol. 2014;98:7645–57.
Garay LA, Sitepu IR, Cajka T, Chandra I, Shi S, Lin T, German JB, Fiehn O, Boundy-Mills KL. Eighteen new oleaginous yeast species. J Ind Microbiol Biotechnol. 2016;43:887–900.
Llamas M, Dourou M, Gonzalez-Fernandez C, Aggelis G, Tomas-Pejo E. Screening of oleaginous yeasts for lipid production using volatile fatty acids as substrate. Biomass Bioenergy. 2020;138:105553.
Ayadi I, Kamoun O, Trigui-Lahiani H, Hdiji A, Gargouri A, Belghith H, Guerfali M. Single cell oil production from a newly isolated Candida viswanathii Y-E4 and agro-industrial by-products valorization. J Ind Microbiol Biotechnol. 2016;43:901–14.
Fontanille P, Kumar V, Christophe G, Nouaille R, Larroche C. Bioconversion of volatile fatty acids into lipids by the oleaginous yeast Yarrowia lipolytica. Bioresour Technol. 2012;114:443–9.
Deeba F, Pruthi V, Negi YS. Converting paper mill sludge into neutral lipids by oleaginous yeast Cryptococcus vishniaccii for biodiesel production. Bioresour Technol. 2016;213:96–102.
Patel A, Pravez M, Deeba F, Pruthi V, Singh RP, Pruthi PA. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour Technol. 2014;165:214–22.
Ördög V, Stirk WA, Bálint P, van Staden J, Lovász C. Changes in lipid, protein and pigment concentrations in nitrogen-stressed Chlorella minutissima cultures. J Appl Phycol. 2012;24:907–14.
Ratledge C, Wynn JP. The biochemistry and molecular biology of lipid accumulation in oleaginous microorganisms. Adv Appl Microbiol. 2002;51:1–52.
Papanikolaou S, Seraphim P, Papanikolaou S. Oleaginous yeasts: biochemical events related with lipid synthesis and potential biotechnological applications. Ferment Technol. 2012;1:1–3.
Chaturvedi S, Bhattacharya A, Nain L, Prasanna R, Khare SK. Valorization of agro-starchy wastes as substrates for oleaginous microbes. Biomass Bioenergy. 2019;127:105294.
Sagia S, Sharma A, Singh S, Chaturvedi S, Nain PK, Nain L. Single cell oil production by a novel yeast Trichosporon mycotoxinivorans for complete and ecofriendly valorization of paddy straw. Electron J Biotechnol. 2020;44:60–8.
Huang C, Wu H, Liu ZJ, Cai J, Lou WY, Zong MH. Effect of organic acids on the growth and lipid accumulation of oleaginous yeast Trichosporon fermentans. Biotechnol Biofuels. 2012;5:4.
Palmqvist E, Hahn-Hägerdal B. Fermentation of lignocellulosic hydrolysates. II: inhibitors and mechanisms of inhibition. Bioresour Technol. 2000;74:25–33.
Yu X, Zeng J, Zheng Y, Chen S. Effect of lignocellulose degradation products on microbial biomass and lipid production by the oleaginous yeast Cryptococcus curvatus. Process Biochem. 2014;49(3):457–65.
Almeida JR, Modig T, Petersson A, Hähn-Hägerdal B, Lidén G, Gorwa-Grauslund MF. Increased tolerance and conversion of inhibitors in lignocellulosic hydrolysates by Saccharomyces cerevisiae. J Chem Technol Biotechnol. 2007;82:340–9.
Chen X, Li Z, Zhang X, Hu F, Ryu DD, Bao J. Screening of oleaginous yeast strains tolerant to lignocellulose degradation compounds. Appl Biochem Biotechnol. 2009;159:591–604.
Chandel AK, Da Silva SS, Singh OV. Detoxification of lignocellulose hydrolysates: biochemical and metabolic engineering toward white biotechnology. Bioenergy Res. 2013;6:388–401.
Zhao X, Peng F, Du W, Liu C, Liu D. Effects of some inhibitors on the growth and lipid accumulation of oleaginous yeast Rhodosporidium toruloides and preparation of biodiesel by enzymatic transesterification of the lipid. Bioprocess Biosyst Eng. 2012;35:993–1004.
Olteanu Z, Rosu CM, Mihasan M, Surdu S, Lacramioara O. Preliminary consideration upon oxido-reductive system involved in aerobic biodegradation of some textile dyes. J Exp and Mol Biol. 2008;9(2):41–6.
Tony BD, Goyal D, Khanna S. Decolorization of textile azo dyes by aerobic bacterial consortium. Int Biodeterior Biodegrad. 2009;63:462–9.
Khehra MS, Saini HS, Sharma DK, Chadha BS, Chimni SS. Decolorization of various azo dyes by bacterial consortium. Dyes pigm. 2005;67:55–61.
Pearce CI, Lloyd JR, Guthrie JT. The removal of colour from textile wastewater using whole bacterial cells: a review. Dyes pigm. 2003;58(3):179–96.
Joshi T, Iyengar L, Singh K, Garg S. Isolation, identification and application of novel bacterial consortium TJ-1 for the decolourization of structurally different azo dyes. Bioresour Technol. 2008;99:7115–21.
Kalyani DC, Patil PS, Jadhav JP, Govindwar SP. Biodegradation of reactive textile dye Red BLI by an isolated bacterium Pseudomonas sp. SUK1. Bioresour Technol. 2008;99:4635–41.
Chang JS, Chou C, Lin YC, Lin PJ, Ho JY, Hu TL. Kinetic characteristics of bacterial azo-dye decolorization by Pseudomonas luteola. Water Res. 2001;35:2841–50.
Al-Tohamy R, Kenawy ER, Sun J, Ali SS. Performance of a newly isolated salt-tolerant yeast strain Sterigmatomyces halophilus SSA-1575 for azo dye decolorization and detoxification. Front Microbiol. 2020;11:1163.
Al-Tohamy R, Sun J, Fareed MF, Kenawy ER, Ali SS. Ecofriendly biodegradation of Reactive Black 5 by newly isolated Sterigmatomyces halophilus SSA1575, valued for textile azo dye wastewater processing and detoxification. Sci Rep. 2020;10(1):12370.
Tan L, He M, Song L, Fu X, Shi S. Aerobic decolorization, degradation and detoxification of azo dyes by a newly isolated salt-tolerant yeast Scheffersomyces spartinae TLHS-SF1. Bioresour Technol. 2016;203:287–94.
Hsueh CC, Chen BY. Comparative study on reaction selectivity of azo dye decolorization by Pseudomonas luteola. J Hazard Mater. 2007;141:842–9.
Chen Y, Feng L, Li H, Wang Y, Chen G, Zhang Q. Biodegradation and detoxification of Direct Black G textile dye by a newly isolated thermophilic microflora. Bioresour Technol. 2018;250:650–7.
Vantamuri AB, Shettar AK. Biodegradation of diazo reactive dye (Green HE4BD) by Marasmius sp. BBKAV79. Chem Data Collect. 2020;28:100422.
Chen KC, Wu JY, Liou DJ, Hwang SC. Decolorization of the textile dyes by newly isolated bacterial strains. J Biotechnol. 2003;101(1):57–68.
Chang JS, Kuo TS, Chao YP, Ho JY, Lin PJ. Azo dye decolorization with a mutant Escherichia coli strain. Biotechnol Lett. 2000;22:807–12.
Jadhav JP, Kalyani DC, Telke AA, Phugare SS, Govindwar SP. Evaluation of the efficacy of a bacterial consortium for the removal of color, reduction of heavy metals, and toxicity from textile dye effluent. Bioresour Technol. 2010;101:165–73.
Fedosov SN, Brask J, Xu X. Analysis of biodiesel conversion using thin layer chromatography and nonlinear calibration curves. J Chromatogr A. 2011;1218:2785–92.
Vinarta SC, Angelicola MV, Barros JM, Fernandez PM, Mac Cormak W, Aybar MJ, De Figueroa LI. Oleaginous yeasts from Antarctica: screening and preliminary approach on lipid accumulation. J Basic Microbiol. 2016;56:1360–8.
Patel A, Arora N, Sartaj K, Pruthi V, Pruthi PA. Sustainable biodiesel production from oleaginous yeasts utilizing hydrolysates of various non-edible lignocellulosic biomasses. Renew Sustain Energy Rev. 2016;62:836–55.
Maza DD, Viñarta SC, Su Y, Guillamón JM, Aybar MJ. Growth and lipid production of Rhodotorula glutinis R4, in comparison to other oleaginous yeasts. J Biotechnol. 2020;310:21–31.
Ramos MJ, Fernández CM, Casas A, Rodríguez L, Pérez Á. Influence of fatty acid composition of raw materials on biodiesel properties. Bioresource Technol. 2009;100:261–8.
Patel A, Matsakas L. A comparative study on de novo and ex novo lipid fermentation by oleaginous yeast using glucose and sonicated waste cooking oil. Ultrasonics Sonochem. 2019;52:364–74.
Hoekman SK, Broch A, Robbins C, Ceniceros E, Natarajan M. Review of biodiesel composition, properties, and specifications. Renew Sustain Energy Rev. 2012;16:143–69.
Suh SO, Blackwell M. Four new yeasts in the Candida mesenterica clade associated with basidiocarp-feeding beetles. Mycologia. 2005;97(1):167–77.
Thakur MS, Prapulla SG, Karanth NG. Microscopic observation of Sudan Black B staining to monitor lipid production by microbes. J Chem Technol Biotechnol. 1988;42(2):129–34.
Miller GL. Use of dinitrosalicyclic reagent for determination of reducing sugar. Anal Chem. 1959;31:426–8.
Gómez-Alonso S, Hermosín-Gutiérrez I, García-Romero E. Simultaneous HPLC analysis of biogenic amines, amino acids, and ammonium ion as aminoenone derivatives in wine and beer samples. J Agric Food Chem. 2007;55(3):608–13.
Orth AB, Royse DJ, Tien MI. Ubiquity of lignin-degrading peroxidases among various wood-degrading fungi. Appl Environ Microbiol. 1993;59(12):4017–23.
Kumar S, Stecher G, Tamura K. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol Biol Evol. 2016;33(7):1870–4.
Rekik H, Jaouadi NZ, Bouacem K, Zenati B, Kourdali S, Badis A, Annane R, Bouanane-Darenfed A, Bejar S, Jaouadi B. Physical and enzymatic properties of a new manganese peroxidase from the white-rot fungus Trametes pubescens strain i8 for lignin biodegradation and textile-dyes biodecolorization. Int J Biol Macromol. 2019;125:514–25.
Vorderwülbecke T, Kieslich K, Erdmann H. Comparison of lipases by different assays. Enzyme Microb Technol. 1992;14(8):631–9.
Hu C, Zhao X, Zhao J, Wu S, Zhao ZK. Effects of biomass hydrolysis by-products on oleaginous yeast Rhodosporidium toruloides. Bioresour Technol. 2009;100:4843–7.
Poontawee R, Yongmanitchai W, Limtong S. Efficient oleaginous yeasts for lipid production from lignocellulosic sugars and effects of lignocellulose degradation compounds on growth and lipid production. Process Biochem. 2017;53:44–60.
Alvarez AF, Alvarez HM, Kalscheuer R, Wältermann M, Steinbüchel A. Cloning and characterization of a gene involved in triacylglycerol biosynthesis and identification of additional homologous genes in the oleaginous bacterium Rhodococcus opacus PD630. Microbiol. 2008;154(8):2327–35.
Morrison WR, Smith LM. Preparation of fatty acid methyl esters and dimethylacetals from lipids with boron fluoride–methanol. J Lipid Res. 1964;5(4):600–8.
The authors are thankful to the Scientific Research Center and Measurement (SRCM) of Tanta University (Egypt), and King Abdulaziz University's Center of Excellence in Environmental Studies in Jeddah (Saudi Arabia) for providing technical assistance in chemical analyses performed in this study.
This work was supported by the National Key R&D Program of China (2018YFE0107100), the National Natural Science Foundation of China (31772529), and the project funded by the Priority of Academic Program Development of Jiangsu Higher Education Institutions (PAPD 4013000011). It was also supported by Taif University Researchers Supporting Project number (TURSP-2020/95), Taif University, Taif, Saudi Arabia.
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Fatty acid composition of M. caribbica SSA1654. Fig. S1. UV–Vis spectrophotometric analysis at 400–800 nm of the simulated wastewater containing 250 mg/L of each applied dye, within 21 h of incubation in the presence of the NYC-1 consortium.
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Al-Tohamy, R., Sun, J., Khalil, M.A. et al. Wood-feeding termite gut symbionts as an obscure yet promising source of novel manganese peroxidase-producing oleaginous yeasts intended for azo dye decolorization and biodiesel production. Biotechnol Biofuels 14, 229 (2021). https://doi.org/10.1186/s13068-021-02080-z
- Meyerozyma caribbica
- Oleaginous yeasts
- Azo dyes
- Manganese peroxidases
- Lignocellulose degradation inhibitors
- Wood-feeding termite gut symbionts