Skip to main content
Download PDF

Lipid production from lignocellulosic biomass using an engineered Yarrowia lipolytica strain

Microbial Cell Factories volume 21, Article number: 226 (2022) Cite this article

Abstract

Background

The utilization of industrial wastes as feedstock in microbial-based processes is a one of the high-potential approach for the development of sustainable, environmentally beneficial and valuable bioproduction, inter alia, lipids. Rye straw hydrolysate, a possible renewable carbon source for bioconversion, contains a large amount of xylose, inaccessible to the wild-type Yarrowia lipolytica strains. Although these oleaginous yeasts possesses all crucial genes for xylose utilization, it is necessary to induce their metabolic pathway for efficient growth on xylose and mixed sugars from agricultural wastes. Either way, biotechnological production of single cell oils (SCO) from lignocellulosic hydrolysate requires yeast genome modification or adaptation to a suboptimal environment.

Results

The presented Y. lipolytica strain was developed using minimal genome modification—overexpression of endogenous xylitol dehydrogenase (XDH) and xylulose kinase (XK) genes was sufficient to allow yeast to grow on xylose as a sole carbon source. Diacylglycerol acyltransferase (DGA1) expression remained stable and provided lipid overproduction. Obtained an engineered Y. lipolytica strain produced 5.51 g/L biomass and 2.19 g/L lipids from nitrogen-supplemented rye straw hydrolysate, which represents an increase of 64% and an almost 10 times higher level, respectively, compared to the wild type (WT) strain. Glucose and xylose were depleted after 120 h of fermentation. No increase in byproducts such as xylitol was observed.

Conclusions

Xylose-rich rye straw hydrolysate was exploited efficiently for the benefit of production of lipids. This study indicates that it is possible to fine-tune a newly strain with as minimally genetic changes as possible by adjusting to an unfavorable environment, thus limiting multi-level genome modification. It is documented here the use of Y. lipolytica as a microbial cell factory for lipid synthesis from rye straw hydrolysate as a low-cost feedstock.

Background

The development of environmentally friendly branches of the biofuel industry is indispensable to establishing the definite suitability of the biorefinery concept. Microbial oils, in contrast to the traditionally used plant oils, have the potential to partially reduce the use of food raw materials for energy purposes and reduce the potential risk of deforestation to acquire new arable land. Additionally, in the process of developing the single cell oils (SCO) production, it is possible to plan targeted changes of the fatty acid profile [1,2,3]. Therefore, many attempts have been made to create a biological factory based on the microbial cells. The lipid production should take a twin-track approach. The first issue is the selection of inexpensive carbon sources as alternatives to the commonly used glucose; thus, attention should be paid to the opportunity of reuse of waste materials obtained from various branches of industry. Some of the previous research has shown that used cooking oil [4, 5], hydrolyzed castor oil [6], olive-mill wastewater [7, 8], crude glycerol [1, 9,10,11,12], molasses [13, 14] or food waste hydrolysates [15, 16] could be useful as feedstock for yeast bioprocesses. Lignocellulosic material, which can be hydrolyzed and converted to monosaccharides, appears to be another good candidate for low-cost feedstock [17,18,19,20]. Three varied fractions, cellulose, hemicellulose and lignin, have been identified from lignocellulosic biomass. Due to the chemical composition of plant cell walls, lignocellulosic hydrolysate is mainly a mixture of glucose and xylose. The high content of xylose, which is poorly metabolized by the most microorganism, due to the carbon catabolite repression suppressing the assimilation of C5 sugars, is one of the obstacles limiting the applications of lignocellulose [21]. In the present study, rye straw, which is a commonly used non-wood waste agricultural raw material in European countries, was used as a potential carbon source. The compositional analysis of the rye straw hydrolysate revealed a diverse sugar content—over 60% xylose, 27% glucose and 9% arabinose, in total 28 g/L monosaccharaides [22]. As a consequence, the second issue is choice of the most appropriate microbial factory cell. Rye straw hydrolysate has already been used for ethanol fermentation by Saccharomyces cerevisiae. However, xylose was detected in hydrolysate after the process [23].

In the context of biofuel production, Y. lipolytica appears justified as a microbe of interest. Y. lipolytica is a non-conventional, but well-studied, oleaginous yeast with huge industrial potential. In recent years, Y. lipolytica has been widely used for biosynthesis of metabolites such as citric acid [10, 24], polyols [25], enzymes [26], terpenoids [27,28,29] and polyketides [30, 31]. The high capacity of Y. lipolytica to convert carbon source into lipid provides an attractive platform for single cell oil (SCO) production. Lipid metabolism in this yeast has been expanded considerably. The overexpression of endogenous genes related to the lipid synthesis pathway caused a well-documented increased amount of SCO [1]. Tai and Stephanopoulos proposed compact strategy co-expression of ACC1 and DGA1 [32]. Furthermore, lipid production can be enhanced by deletion of genes related to lipid degradation such as MFE2 or genes associated with peroxisome biogenesis [33,34,35]. Additionally, evolution-based strategies were used as a tool for improving lipid accumulation [36]. The combination of increased lipid accumulation with efficient growth on a lignocellulosic substrate could lead to the creation of a biotechnological chassis for viable lipid production. One obstacle is that commonly used laboratory Y. lipolytica strains for biotechnology applications are unable to utilize xylose as its sole carbon source. The xylose utilization pathway, apart from the necessary transporters, requires activity of xylose reductase (XR), which mediates conversion of xylose to xylitol, xylulose dehydrogenase (XDH), which converts xylitol to xylulose, and xylulose kinase (XK) for final conversion to xylulose-5-phospate. Xylose-degrading enzymes have been confirmed in the Y. lipolytica system [37], nevertheless, without adaptation or genetic modification, yeast growth is insufficient. Table 1 summarizes research to date on the use of xylose utilization or co-fermentation of mixed sugars. The fermentation based on pure xylose should be considered as preliminary research for further use of xylose-rich wastes or as basic and necessarily metabolic studies.

Table 1 Comparison of xylose and waste-based fermentation of previously reported Y. lipolytica
Full size table

The purpose of this study was to examine the feasibility of using rye straw hydrolysate as a carbon source for the engineered Y. lipolytica strain. The modified strain, with limited genome modification, using exclusively native genes, was able to convert the entire sugar pool from low-cost lignocellulosic hydrolysate and produce a significant amount of lipids.

Results

Overexpression of the endogenous xylose utilization pathway and improving growth kinetics

The wild-type Y. lipolytica A101 strain is incapable of growing on xylose as a sole carbon source, which was confirmed in the presented studies and which is quite common for biotechnological relevant strains [44, 47]. The aim of the study was to develop a new strain with as few genetic changes as possible. Endogenous xylose reductase, xylitol dehydrogenase and xylulose kinase were integrated in pairs into an AJD pAD-DGA1 strain [48]. The parental yeast strain with overexpression of DGA1 (YALI0E32769g) is characterized by increased lipid production and accumulation [1, 49]. Strains were prepared according to the scheme in Additional file 1: Fig. S1. To achieve high protein expression, all genes were overexpressed under the control of the UAS1B16-TEF hybrid promoter [50]. The expression level of the functional genes was evaluated by qRT-PCR. The results showed that the engineered strains exhibit high levels of gene expression (Fig. 1). Lack of statistically different magnitude of relative gene expression, provided by the high-strength promoter, allows the comparison of the obtained strains under identical growth-medium conditions. The expression level of the previously integrated DGA1 gene also remained at a similar level in all tested strains.

Fig. 1
figure 1

Relative quantification of RNA transcript in the engineered strains. The histograms show the relative expression of genes with respect to the control—actin was used as a reference gene and Y. lipolytica A101 was used as a control strain. Samples were analyzed in triplicate. One-way ANOVA at p ≤ 0.05 was calculated. The homogeneous groups between the same genes according to the Duncan test were estimated and they are represented by the same letter

Full size image

Next, the engineered strains were tested for growth ability under sole xylose conditions in a shake-flask experiment. After 120 h AJD-D/XDH/XK began the increase of biomass exclusively. In Fig. 2 the growth and xylose assimilation curves are presented. At the end of the 12-day period, the biomass of AJD-D/XDH/XK reached the level of 5.2 g/L (the biomass total productivity Qx = 18.06 ± 1.13 mg/L/h, the biomass total yield Yx = 0.26 g/g). The remaining strains did not demonstrate a sufficient capacity for growth.

Fig. 2
figure 2

Growth of different Y. lipolytica engineered strains on xylose—(I) biomass increase (solid line) of Y. lipolytica A101, AJD-D/XR/XK (introducing heterologous xylose reductase and xylulose kinase), AJD-D/XR/XDH (introducing heterologous xylose reductase and xylitol dehydrogenase) and AJD-D/XDH/XK (introducing heterologous xylitol dehydrogenase and xylulose kinase), (II) xylose concentration (dotted line)

Full size image

It was assumed that prolonged exposure to an unfavorable, but cell-growth-enabling medium composition, would lead to changes Y. lipolytica gene expression level and, consequently, a change in phenotype [51]. Despite obtaining a strain able to grow on pure xylose, the long duration of the lag phase was a predicament. Instead of introducing further genetic modifications, it was decided to use time-extended shake-flask culture to accelerate cell growth. The starting point was the YNB medium consisting of 20 g/L xylose inoculated with starved yeast cells. The starved strain was previously cultured on YNB medium supplemented with glucose and left under carbon deficient conditions for a 10 days, then double-washed with sterile water and transferred to appropriate xylose-based medium. The yeast was transferred to fresh medium at an interval of 7 days. Changes in xylose assimilation between successive passages are shown in Fig. 3. In the early passages the sugar concentration decreased slightly. With each subsequent passage, the yeast began to utilize xylose more and more efficiently. Finally obtained yeast cell population with the ability to assimilate 20 g/L of xylose within 7 days was considered homogeneous and was named AJD-D/XYL/ALE.

Fig. 3
figure 3

Changes in xylose assimilation from the medium during culture AJD-D/XDH/XK strain, performed by sequential serial passages in shake cultural tube. At intervals an aliquot of the culture was transferred to a new medium in the tube. Each medium contained 20 g/L xylose supplemented with 6.7 g/L YNB. The strain after the last passages was used in the next research and has been described as Y. lipolytica AJD-D/XYL/ALE

Full size image

Differences in growth kinetics between parental and newly developed strains were examined in a shake-flask experiment. The OD600 and xylose assimilation curves are presented in Fig. 4A. The strain AJD-D/XDH/XK started to grow in the logarithmic phase after 96 h. The overall xylose assimilation occurred after 312 h, and the final optical density reached a level less than 25. The strain AJD-D/XYL/ALE utilized the total amount of xylose during 144 h and attained the highest grown parameters, such as xylose consumption rate 138.89 ± 3.37 mg/L/h and optical density level—OD600 = 32.3. In comparison, the initial strain after 144 h OD600 stood at 5.6 and xylose consumption rate achieved 11.11 ± 0.41 mg/L/h. The duration of the lag phase was significantly reduced, which was observed through the microplate reader (Fig. 4B).

Fig. 4
figure 4

Variations in growth characteristics between wild type, AJD-D/XDH/XK and AJD-D/XYL/ALE. (A) OD600 (dashed line) and xylose concentration (solid line) curves. (B) OD600 during first 45 h of growth on media containing 20 g/L xylose supplemented with 6.7 g/L YNB

Full size image

Lipid production from waste-based medium

Rye straw is one of the many lignocellulosic agricultural wastes that can play a prominent role in replacing pure sugars, such as glucose, as a primary substrate used in the biotechnological industry. The main sugar found in straw hydrolysate is xylose, over 61%. The remainder of the sugar composition consists of glucose and arabinose [22]. To explore the feasibility of rye straw hydrolysate fermentation by the newly obtained strain of Y. lipolytica a shake-flask experiment with the new substrate was performed. Co-expression of the xylose metabolism pathway with diacylglycerol acyltransferase (DGA1) should ensure biovalorization of the waste substrate into SCO. After 120 h, the parental strain Y. lipolytica A101 reached only 3.35 g/L of biomass and 0.2 g/L of triacylglycerols (Fig. 5). The strain AJD-D/XYL/ALE greatly increased the biomass level by more than 60% and lipid level by almost tenfold at 5.51 g/L and 2.19 g/L, respectively. All sugars in the medium were depleted. Importantly, no xylitol or citric acid production was observed during the fermentation process. The total yield and productivity are shown in Table 2.

Fig. 5
figure 5

Parameters of biomass and lipid production by Y. lipolytica A101, AJD-D/XDH/XK and AJD-D/XYL/ALE grown on rye straw hydrolysates containing about 61% xylose, 27% glucose, 8% arabinose and insignificant amounts of mannose or galactose, supplemented with YNB. Data from triplicate fermentations are shown. One-way ANOVA at p ≤ 0.05 was calculated and homogeneous groups were estimated according to the Duncan test. Mean values over the bars that are not significantly different from each other (p > 0.05) are represented by the same letter

Full size image
Table 2 Lipids and biomass production by Y. lipolytica strains grown on rye straw hydrolysates supplemented with YNB at the end of the shake-flask experiment
Full size table

The fatty acid composition of the obtained biomasses was analyzed and is shown compared to rapeseed oil in Table 3. Overexpression of DGA1 not only increases the lipid accumulation but also affects the change of lipid profile. Particularly high levels of oleic acid (C18:1) were observed in the modified strains with a simultaneously reduced amount of linoleic acid (C18:2). Oleic acid accounted for more than 63% of all fatty acid in the lipid pool in the AJD-D/XYL/ALE biomass. Interestingly, long-chain docosanoic acid, C22:0, and tetracosanoic acid, C24:0, were identified in the yeast biomass, with the highest amount detected in the biomass of the strain AJD-D/XYL/ALE.

Table 3 Percent of total cellular lipids of Y. lipolytica biomass obtained on hydrolysates-based medium supplemented by YNB compared to rapeseed oil fatty acid profile
Full size table

Discussion

The effective production of valuable metabolites is closely related to the complete utilization of the carbon source in the medium. Lignocellulosic waste materials, such as straw, may provide varied sugar composition by introducing glucose, xylose, or arabinose, among others, into the medium. Previous studies have shown that acid-enzymatic hydrolysis of rye straw allows one to obtain a sugar blend available for yeast growth [22]. However, the amount of yeast biomass produced was at a moderate level. The preliminary view was taken that improving xylose assimilation appeared essential to increase the effectiveness of biomass and lipid production from lignocellulosic materials. Therefore, a number of researchers have attempted to modify Y. lipolytica to utilize xylose. Initial attempts to engineer xylose metabolism by the heterologous expression of Scheffersomyces stipitis genes, a microbial model for xylose metabolism, failed to achieve strain able to grown on xylose, until after overexpressed three genes involved in xylose utilization pathway and received strain efficiently grown on xylose but simultaneously produced xylitol [44]. Similar growth parameters were achieved by the overexpression of the native genes [46]. Different approach in which introduced mutated xylose isomerase gene and performed adaptive laboratory evolution also resulted in receiving new Y. lipolytica strain enable to utilize xylose [39]. The aim of the present study was to reduce the amount of genetic modification and as has already been pointed out, that the double overexpression of xylitol dehydrogenase (XDH) and xylulose kinase (XK) is sufficient to enable Y. lipolytica to grow on xylose as the sole carbon source without adaptation [37]. However, the long lag phase is a bottleneck during production processes. Appropriate tool for metabolic changes, avoiding additional genetic modification, is adaptive laboratory evolution—a process based on natural evolution and selection directed to obtaining a better fitness to adapt to specific environmental conditions. ALE has been employed for an improved various important parameters in different aspects of biotechnology [52]. ALE was also proved to be effective for a wild-type Y. lipolytica strain and after adaptation yeast grew on xylose, but the growth rate was negligible. It has also been shown that strong expression of xylitol dehydrogenase is considered a priority factor for xylose metabolism [42]. Although ALE was not applied in the full sense in the present study, the simpler technique of prolonged culture in suboptimal medium already achieved a reduction in the time to reach the logarithmic phase of growth, which was an essential point in the creation of the new strain. Potential changes in genotype and phenotype stability remained unstudied, which would require genome sequencing of the adopted strain.

Therefore, the UAS1B16-TEF promoter, which provides high levels of gene expression, was used to construct the strain. In the presented study, co-expression of all xylose metabolism pathway genes was abandoned. The lack of overexpression of xylose reductase, the enzyme directly responsible for the conversion of xylose to xylitol, reduces the level of the by-product. In the conducted cultures, xylitol was not detected based on the HPLC analysis. Strain AJD/XYL/ALE is characterized by a satisfactory growth rate, short lag phase and high biomass production, after fermentation on pure xylose based medium. The results obtained are comparable to other xylose utilization processes with a total use of 20 g/L xylose in 5 days or OD600 = 25.8 after 4 days of fermentation compared to OD600 = 26.8 obtained by Y. lipolytica Y14 strain with multiple genome modification [38]. An essential part of the study was the production of lipids on waste substrate as a carbon source, rye straw hydrolysate. The total biomass productivity of AJD/XYL/ALE on hydrolysate was almost 2.5 times higher than reached during growth on pure xylose (46 mg/L/h compared to 18.06 mg/L/h) and 1.6 times higher than wild Y. lipolytica A101 growth on lignocellulosic hydrolysate (28 mg/L/h). The obtained strain showed enhanced synthesis of SCO without any major delay in growth. Both strains AJD/XDH/XK and AJD/XYL/ALE showed the same level of lipid accumulation above 35%. The improved availability of xylose in production medium for AJD/XYL/ALE strain resulted in an increased biomass levels. The process based on hydrolysate from agave bagasse and ylXYL + Obese strain allowed to obtain over 10 g/L biomass with 24.5% lipid content [46]. The shake-flask experiment showed that rye straw hydrolysate is a proper low-cost medium for fatty acid bioproduction. A number of valuable studies on lipid production by Y. lipolytica have been carried out to date. Higher level of lipid content (grams of lipid per gram of cell dry weight) have been obtained in bioprocess using glucose as substrate—in the range of 77–80% [54,55,56]. However, glucose-based processes generate high production costs. To reduce the outlay on lipid production, alternative carbon sources, such as glycerol, have begun to be used. Glycerol as a carbon source made it possible to obtain a wide range of lipid content. On glycerol as a sole substrate 52% lipid content was obtained [35], a combination of glycerol and molasses allowed for synthesis of 31% lipids [57], and glycerol enriched with xylose enabled the production of 42% lipids [44]. However, not all lignocellulosic raw materials are mainly composed of xylose. In rice bran hydrolysate, glucose was the dominant sugar [58]. The lipid content of Y. lipolytica was over 48%. Due to complex and differentiated chemical composition of lignocellulosic materials, they give the possibility to select the most appropriate substrate according to the purpose of bioproduction. In addition to the amount of lipids accumulated in cells, the fatty acids profile is also influential. It is worth noting that overexpression of DGA1 resulted in a slightly different fatty acid pool in the biomass. An overall increase in saturated fatty acids (palmitic acid C16:0, stearic acid C18:0, behenic acid C22:0 and lignoceric acid C24:0) was observed. This is in agreement with the previous reports [1]. During the shake-flask experiment, the major fatty acid in the pool was oleic acid C18:1—partial Y. lipolytica A101 contained 50%, AJD/XYL/ALE increased this amount to over 63%. The domination of the acid C18:1 is typical for Y. lipolytica biolipids [59]. Interestingly, over half the linoleic acid (and isomers of C18:2 linolelaidic acid) content was reduced in xylose-targeted modified yeast strains. Prior researchers have demonstrated that various types of the provided substrate can affect the fatty acid profile [60]. The genetic modification of yeast also appears to have an effect on the lipid pool [1]. The present experiment verified that the medium based on a lignocellulosic waste such as rye straw is a relevant, low-cost medium for microbial lipid production by Y. lipolytica. The results provide a promising starting point for optimization aimed at enhancing the SCO titer.

Conclusions

This study showed that overexpression of the native xylitol dehydrogenase (XDH) and xylulose kinase (XK) genes in Y. lipolytica allowed utilization of xylose as a sole carbon source. Subsequently, the prolonged-term shake-flask culture was used to reduce the duration of the lag phase and increase biomass level. The study carried out on a yeast strain that was previously prepared to overproduce lipid enabled the efficient conversion of xylose into microbial lipids. Rye straw hydrolysate, mainly composed of xylose and glucose, was thereafter tested as a carbon source for single cell oil production. The most efficient growth was observed with the strain AJD/XYL/ALE, and the lipid titer was improved tenfold over the control. This study provides evidence of low-cost bioproduction of lipids from lignocellulosic material. It also provides prospects for process optimization studies. The presented results brought to development one of the elements of green-biotechnology processes for enabling industrial waste management.

Methods

Strains and plasmids

Primary strains used in this study were Y. lipolytica A101 [61] and AJD with overexpression of YALI0E32769g—diacylglycerol acyltransferase Dga1p [40]. Both strains belong to the Department of Biotechnology and Food Microbiology at Wroclaw University of Environmental and Life Sciences, Poland. Escherichia coli DH5α was primarily used for molecular cloning. Vector pAD [62], carrying the UAS1B16-TEF promoter, was the basis for developing new plasmids with the native XR, XDH or XK gene fragment. All plasmids and strains used in this study are listed in Additional file 1: Table S1 in the supplemental material. The all XYL genes fragments were amplified from the Y. lipolytica A101 genomic DNA. The list of primers used is shown in Additional file 1: Table S2. The PCR amplified genes were cloned into the pAD vector using SgsI and NheI/Pml1 sites, T4 DNA Ligase (Thermo Fisher Scientific) and used for transformation of E. coli. The obtained plasmids were isolated using the Plasmid Mini Kit (A&A Biotechnology, Poland), sequenced (Genomed, Poland) and digested with MssI. Linear expression cassettes were used to transform yeast according to the lithium acetate method [63]. The restriction enzymes were acquired from FastDigest Thermo Scientific.

Culture media

For molecular cloning in E. coli LB medium (A&A Biotechnology, Poland) was used supplemented with 100 µg/mL ampicillin for selection after transformation. For the yeast inoculum preparation 6.7 g/L YNB (Yeast Nitrogen Base, Merck, Germany) supplemented by 20 g/L of glucose was used.

  1. (i)

    Pure xylose-based medium Growth of Y. lipolytica was conducted at 28 °C, 250 rpm in a 300 mL flask. The flask contained 50 mL of medium consisting of 20 g/L xylose (Sigma, Germany) and 6.7 g/L YNB. The same medium was used during growth tests.

  2. (ii)

    Transcript quantification For RNA isolation the strains were grown for 24 h in medium consisting of 6.7 g/L YNB (Yeast Nitrogen Base, Merck, Germany) and 20 g/L glucose (Merck, Germany).

  3. (iii)

    Agricultural waste medium Y. lipolytica A101, AJD-D/XDH/XK and AJD-D/XYL/ALE were used for biomass and lipid production with waste-based medium. 50 mL of rye straw hydrolysates [22] was supplemented with 6.7 g/L YNB (Yeast Nitrogen Base, Merck, Germany), sterilized by membrane filtration (Stericup Filter Units, 0.22 µm Durapore, Merck Millipore) and used for growth test and shake-flask experiments.

Growth cultures

The AJD-D/XDH/XK strain was selected for cultivation aimed at reducing phase lag. The process lasted for 80 days. Every 7 days the current culture standardized to OD600 = 0.01 was used as an inoculum for the next fresh medium. The composition of the medium remained constant for each passages. Another 1 mL of current culture was immediately frozen with 0.7 mL of 30% glycerol in liquid nitrogen and stored at − 80 °C. Quantification of xylose was executed every day. The strain obtained after the last passage was named AJD-D/XYL/ALE. Then, the growth of the wild and all newly developed strains were tested in the Spark microplate reader (Tecan Group Ltd., Switzerland). The overnight inoculation cultures were centrifuged, washed with sterile water and standardized to OD600 = 0.15. The strains were grown in 100-well plates in 200 μL of xylose-based medium or waste-based medium. Quadruple experiments were performed under a constant agitation rate at 28 °C with measurement of optical density at 420–560 nm every 30 min for 72 h. Negative controls with no yeast were included.

RNA isolation and transcript quantification

For RNA isolation cultures after 24 h of growth were centrifuged for 5 min at 12,000×g. Biomass was treated using the Total RNA Mini Plus kit (A&A Biotechnology, Poland). The obtained RNA was standardized to an equal concentration and additionally purified by DNase I (Thermo Scientific, USA) according to the producer’s instructions. Quantification of RNA was executed using a Biochrom WPA Biowave II spectrophotometer (Biochrom Ltd., UK) equipped with a TrayCell (Hellma Analytics, Germany). Maxima First Strand cDNA Synthesis kits for RT-qPCR (Thermo Fisher Scientific) were used for cDNA synthesis. For qRT-PCR analyses the DyNAmo Flash SYBR Green qPCR Kit (Thermo Fisher Scientific) was applied with the Eco Real-Time PCR System (Illumina, USA). The list of primers used is shown in Additional file 1: Table S2. Each sample was evaluated in triplicate. Expression level of genes was normalized to the actin gene. Data were analyzed using the ddCT method [64].

Analytical method

The growth curves were determined by monitoring at 24-h intervals the optical density (OD600) with a UV spectrometer (SmartSpec Plus, Bio-Rad, USA) with a semi-micro cuvette (Sarstedt, Germany). Quantitative analysis of biomass was determined gravimetrically after centrifuging (5 min; 5500×g) 10 mL of samples from the shake-flask cultures, washed with distilled water, harvested by filtration on 0.22-μm membranes and drying at 105 °C.

  1. (i)

    HPLC The concentrations of xylose were determined by HPLC using a Carbohydrate Analysis Column Aminex HPX-87P (Bio-Rad, USA) coupled to a UV (Dionex, USA) and a refractive index (RI) detector Shodex (Showa Denko K.K, Japan). The column was eluted with sterile water at 65 °C and a flow rate of 0.6 mL/min.

  2. (ii)

    GC–MS Lipids from 15 to 20 mg of lyophilized biomass were converted into their methyl esters according the method described before [65]. Fatty acids were identified and quantified by comparison to reference material (37 FAME MIX, CRM47885, Sigma-Aldrich). GC–MS analysis was performed with a Shimadzu single quadrupole GCMS-QP2010SE instrument equipped with a Zebron ZB-FAME column. Hexane was used as a solvent, and helium was used as a carrier gas (linear velocity—35 cm/s).

Calculation of fermentation parameters

The formula Yc = X/S was used to calculated the biomass total yield. X mean the final amount of biomass after at the end of the experiment. S mean the total amount of sugar in the hydrolysate. The total lipid yield was calculated in in the same manner.

The formula Qx = X/120 was used to calculated the biomass total productivity. An analogical formula was use to calculated the total lipid productivity.

The formula Qs = ΔS/144 was used to calculated the xylose consumption rate.

Statistical analysis

Statistical analysis of gene expression data, one-way ANOVA, were conducted using Statistica 12.5 (StatSoft, Kraków, Poland). Significant differences (p ≤ 0.05) between mean were assessed by Duncan’s t-test.

Availability of data and materials

Not applicable.

References

  1. Dobrowolski A, Drzymala K, Mitula P, Mironczuk AM. Production of tailor-made fatty acids from crude glycerol at low pH by Yarrowia lipolytica. Bioresour Technol. 2020;314: 123746.

    Article  CAS  PubMed  Google Scholar 

  2. Bhutada G, Menard G, Bhunia RK, Hapeta PP, Ledesma-Amaro R, Eastmond PJ. Production of human milk fat substitute by engineered strains of Yarrowia lipolytica. Metab Eng Commun. 2022;14: e00192.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Konzock O, Matsushita Y, Zaghen S, Sako A, Norbeck J. Altering the fatty acid profile of Yarrowia lipolytica to mimic cocoa butter by genetic engineering of desaturases. Microb Cell Fact. 2022;21:25.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. Katre G, Ajmera N, Zinjarde S, RaviKumar A. Mutants of Yarrowia lipolytica NCIM 3589 grown on waste cooking oil as a biofactory for biodiesel production. Microb Cell Fact. 2017;16:176.

    Article  PubMed  PubMed Central  Google Scholar 

  5. Liu X, Lv J, Xu J, Zhang T, Deng Y, He J. Citric acid production in Yarrowia lipolytica SWJ-1b yeast when grown on waste cooking oil. Appl Biochem Biotechnol. 2015;175:2347–56.

    Article  CAS  PubMed  Google Scholar 

  6. Braga A, Belo I. Immobilization of Yarrowia lipolytica for aroma production from castor oil. Appl Biochem Biotechnol. 2013;169:2202–11.

    Article  CAS  PubMed  Google Scholar 

  7. Sarris D, Rapti A, Papafotis N, Koutinas AA, Papanikolaou S. Production of added-value chemical compounds through bioconversions of olive-mill wastewaters blended with crude glycerol by a Yarrowia lipolytica strain. Molecules. 2019;24:222.

    Article  PubMed Central  Google Scholar 

  8. Dourou MKA, Juszczyk P, Sarris D, Bellou S, Triantaphyllidou IE, Rywinska A, Papanikolaou S, Aggelis G. Bioconversion of olive mill wastewater into high-added value products. J Clean Prod. 2016;139:957–69.

    Article  CAS  Google Scholar 

  9. Dobrowolski A, Mitula P, Rymowicz W, Mironczuk AM. Efficient conversion of crude glycerol from various industrial wastes into single cell oil by yeast Yarrowia lipolytica. Bioresour Technol. 2016;207:237–43.

    Article  CAS  PubMed  Google Scholar 

  10. Rzechonek DA, Dobrowolski A, Rymowicz W, Mironczuk AM. Aseptic production of citric and isocitric acid from crude glycerol by genetically modified Yarrowia lipolytica. Bioresour Technol. 2019;271:340–4.

    Article  CAS  PubMed  Google Scholar 

  11. Luo Z, Miao J, Luo W, Li G, Du Y, Yu X. Crude glycerol from biodiesel as a carbon source for production of a recombinant highly thermostable beta-mannanase by Pichia pastoris. Biotechnol Lett. 2018;40:135–41.

    Article  CAS  PubMed  Google Scholar 

  12. Chmielarz M, Blomqvist J, Sampels S, Sandgren M, Passoth V. Microbial lipid production from crude glycerol and hemicellulosic hydrolysate with oleaginous yeasts. Biotechnol Biofuels. 2021;14:65.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Gajdos P, Nicaud JM, Rossignol T, Certik M. Single cell oil production on molasses by Yarrowia lipolytica strains overexpressing DGA2 in multicopy. Appl Microbiol Biotechnol. 2015;99:8065–74.

    Article  CAS  PubMed  Google Scholar 

  14. Mironczuk AM, Rakicka M, Biegalska A, Rymowicz W, Dobrowolski A. A two-stage fermentation process of erythritol production by yeast Y. lipolytica from molasses and glycerol. Bioresour Technol. 2015;198:445–55.

    Article  CAS  PubMed  Google Scholar 

  15. Li C, Gao S, Li X, Yang X, Lin CSK. Efficient metabolic evolution of engineered Yarrowia lipolytica for succinic acid production using a glucose-based medium in an in situ fibrous bioreactor under low-pH condition. Biotechnol Biofuels. 2018;11:236.

    Article  PubMed  PubMed Central  Google Scholar 

  16. Arous FFF, Triantaphyllidou IE, Aggelis G, Nasri M, Mechichi T. Potential utilization of agro-industrial wastewaters for lipid production by the oleaginous yeast Debaryomyces etchellsii. J Clean Prod. 2016;133:899–909.

    Article  CAS  Google Scholar 

  17. Brandenburg J, Poppele I, Blomqvist J, Puke M, Pickova J, Sandgren M, Rapoport A, Vedernikovs N, Passoth V. Bioethanol and lipid production from the enzymatic hydrolysate of wheat straw after furfural extraction. Appl Microbiol Biotechnol. 2018;102:6269–77.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  18. Fei Q, O’Brien M, Nelson R, Chen X, Lowell A, Dowe N. Enhanced lipid production by Rhodosporidium toruloides using different fed-batch feeding strategies with lignocellulosic hydrolysate as the sole carbon source. Biotechnol Biofuels. 2016;9:130.

    Article  PubMed  PubMed Central  Google Scholar 

  19. Mardetko NGM, Santek B, Trontel A, Grubišić M, Novak M. Bioethanol production from dilute-acid pre-treated wheat straw liquor hydrolysate by genetically engineered Saccharomyces cerevisiae. Chem Biochem Eng Q. 2019;32:483–99.

    Article  Google Scholar 

  20. Flores-Gomez CA, Escamilla Silva EM, Zhong C, Dale BE, da Costa SL, Balan V. Conversion of lignocellulosic agave residues into liquid biofuels using an AFEX-based biorefinery. Biotechnol Biofuels. 2018;11:7.

    Article  PubMed  PubMed Central  Google Scholar 

  21. Jin YS, Cate JH. Metabolic engineering of yeast for lignocellulosic biofuel production. Curr Opin Chem Biol. 2017;41:99–106.

    Article  CAS  PubMed  Google Scholar 

  22. Drzymala K, Mironczuk AM, Pietrzak W, Dobrowolski A. Rye and oat agricultural wastes as substrate candidates for biomass production of the non-conventional yeast Yarrowia lipolytica. Sustainability. 2020;12:7704.

    Article  Google Scholar 

  23. Robak K, Balcerek M, Dziekonska-Kubczak U, Dziugan P. Effect of dilute acid pretreatment on the saccharification and fermentation of rye straw. Biotechnol Prog. 2019;35: e2789.

    Article  PubMed  Google Scholar 

  24. Carsanba E, Papanikolaou S, Fickers P, Erten H. Screening various Yarrowia lipolytica strains for citric acid production. Yeast. 2019;36:319–27.

    Article  CAS  PubMed  Google Scholar 

  25. Rakicka M, Biegalska A, Rymowicz W, Dobrowolski A, Mirończuk AM. Polyol production from waste materials by genetically modified Yarrowia lipolytica. Bioresour Technol. 2017;243:393–9.

    Article  CAS  PubMed  Google Scholar 

  26. Yan J, Han B, Gui X, Wang G, Xu L, Yan Y, Madzak C, Pan D, Wang Y, Zha G, Jiao L. Engineering Yarrowia lipolytica to simultaneously produce lipase and single cell protein from agro-industrial wastes for feed. Sci Rep. 2018;8:758.

    Article  PubMed  PubMed Central  Google Scholar 

  27. Liu D, Liu H, Qi H, Guo XJ, Jia B, Zhang JL, Yuan YJ. Constructing yeast chimeric pathways to boost lipophilic terpene synthesis. ACS Synth Biol. 2019;8:724–33.

    Article  PubMed  Google Scholar 

  28. Tramontin LRR, Kildegaard KR, Sudarsan S, Borodina I. Enhancement of astaxanthin biosynthesis in oleaginous yeast Yarrowia lipolytica via microalgal pathway. Microorganisms. 2019;7:472.

    Article  CAS  PubMed Central  Google Scholar 

  29. Liu Y, Jiang X, Cui Z, Wang Z, Qi Q, Hou J. Engineering the oleaginous yeast Yarrowia lipolytica for production of alpha-farnesene. Biotechnol Biofuels. 2019;12:296.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Markham KA, Palmer CM, Chwatko M, Wagner JM, Murray C, Vazquez S, Swaminathan A, Chakravarty I, Lynd NA, Alper HS. Rewiring Yarrowia lipolytica toward triacetic acid lactone for materials generation. Proc Natl Acad Sci USA. 2018;115:2096–101.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. Palmer CM, Miller KK, Nguyen A, Alper HS. Engineering 4-coumaroyl-CoA derived polyketide production in Yarrowia lipolytica through a beta-oxidation mediated strategy. Metab Eng. 2020;57:174–81.

    Article  CAS  PubMed  Google Scholar 

  32. Tai M, Stephanopoulos G. Engineering the push and pull of lipid biosynthesis in oleaginous yeast Yarrowia lipolytica for biofuel production. Metab Eng. 2013;15:1–9.

    Article  CAS  PubMed  Google Scholar 

  33. Dulermo R, Dulermo T, Gamboa-Melendez H, Thevenieau F, Nicaud JM. Role of Pex11p in lipid homeostasis in Yarrowia lipolytica. Eukaryot Cell. 2015;14:511–25.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Blazeck J, Hill A, Liu L, Knight R, Miller J, Pan A, Otoupal P, Alper HS. Harnessing Yarrowia lipolytica lipogenesis to create a platform for lipid and biofuel production. Nat Commun. 2014;5:3131.

    Article  PubMed  Google Scholar 

  35. Bhutada G, Kavscek M, Ledesma-Amaro R, Thomas S, Rechberger GN, Nicaud JM, Natter K. Sugar versus fat: elimination of glycogen storage improves lipid accumulation in Yarrowia lipolytica. FEMS Yeast Res. 2017;17: fox020.

    Article  PubMed Central  Google Scholar 

  36. Daskalaki A, Perdikouli N, Aggeli D, Aggelis G. Laboratory evolution strategies for improving lipid accumulation in Yarrowia lipolytica. Appl Microbiol Biotechnol. 2019;103:8585–96.

    Article  CAS  PubMed  Google Scholar 

  37. Rodriguez GM, Hussain MS, Gambill L, Gao D, Yaguchi A, Blenner M. Engineering xylose utilization in Yarrowia lipolytica by understanding its cryptic xylose pathway. Biotechnol Biofuels. 2016;9:149.

    Article  PubMed  PubMed Central  Google Scholar 

  38. Wu Y, Xu S, Gao X, Li M, Li D, Lu W. Enhanced protopanaxadiol production from xylose by engineered Yarrowia lipolytica. Microb Cell Fact. 2019;18:83.

    Article  PubMed  PubMed Central  Google Scholar 

  39. Yook SDKJ, Gong G, Ko JK, Um Y, Han SO, Lee S. High-yield lipid production from lignocellulosic biomass using engineered xylose-utilizing Yarrowia lipolytica. GCB Bioenergy. 2020;00:1–10.

    Google Scholar 

  40. Prabhu AA, Thomas DJ, Ledesma-Amaro R, Leeke GA, Medina A, Verheecke-Vaessen C, Coulon F, Agrawal D, Kumar V. Biovalorisation of crude glycerol and xylose into xylitol by oleaginous yeast Yarrowia lipolytica. Microb Cell Fact. 2020;19:121.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. Li H, Alper HS. Producing Biochemicals in Yarrowia lipolytica from xylose through a strain mating approach. Biotechnol J. 2020;15: e1900304.

    Article  PubMed  Google Scholar 

  42. Ryu S, Hipp J, Trinh CT. Activating and elucidating metabolism of complex sugars in Yarrowia lipolytica. Appl Environ Microbiol. 2016;82:1334–45.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  43. Li H, Alper HS. Enabling xylose utilization in Yarrowia lipolytica for lipid production. Biotechnol J. 2016;11:1230–40.

    Article  CAS  PubMed  Google Scholar 

  44. Ledesma-Amaro R, Lazar Z, Rakicka M, Guo Z, Fouchard F, Coq AC, Nicaud JM. Metabolic engineering of Yarrowia lipolytica to produce chemicals and fuels from xylose. Metab Eng. 2016;38:115–24.

    Article  CAS  PubMed  Google Scholar 

  45. Prabhu AA, Ledesma-Amaro R, Lin CSK, Coulon F, Thakur VK, Kumar V. Bioproduction of succinic acid from xylose by engineered Yarrowia lipolytica without pH control. Biotechnol Biofuels. 2020;13:113.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  46. Niehus X, Crutz-Le Coq AM, Sandoval G, Nicaud JM, Ledesma-Amaro R. Engineering Yarrowia lipolytica to enhance lipid production from lignocellulosic materials. Biotechnol Biofuels. 2018;11:11.

    Article  PubMed  PubMed Central  Google Scholar 

  47. Zhao C, Gu D, Nambou K, Wei L, Chen J, Imanaka T, Hua Q. Metabolome analysis and pathway abundance profiling of Yarrowia lipolytica cultivated on different carbon sources. J Biotechnol. 2015;206:42–51.

    Article  CAS  PubMed  Google Scholar 

  48. Dobrowolski A, Drzymala K, Rzechonek DA, Mitula P, Mironczuk AM. Lipid production from waste materials in seawater-based medium by the yeast Yarrowia lipolytica. Front Microbiol. 2019;10:547.

    Article  PubMed  PubMed Central  Google Scholar 

  49. Athenstaedt K. YALI0E32769g (DGA1) and YALI0E16797g (LRO1) encode major triacylglycerol synthases of the oleaginous yeast Yarrowia lipolytica. Biochim Biophys Acta. 2011;1811:587–96.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  50. Blazeck J, Reed B, Garg R, Gerstner R, Pan A, Agarwala V, Alper HS. Generalizing a hybrid synthetic promoter approach in Yarrowia lipolytica. Appl Microbiol Biotechnol. 2013;97:3037–52.

    Article  CAS  PubMed  Google Scholar 

  51. Wang Z, Zhou L, Lu M, Zhang Y, Perveen S, Zhou H, Wen Z, Xu Z, Jin M. Adaptive laboratory evolution of Yarrowia lipolytica improves ferulic acid tolerance. Appl Microbiol Biotechnol. 2021;105:1745–58.

    Article  CAS  PubMed  Google Scholar 

  52. Mavrommati M, Daskalaki A, Papanikolaou S, Aggelis G. Adaptive laboratory evolution principles and applications in industrial biotechnology. Biotechnol Adv. 2022;54: 107795.

    Article  CAS  PubMed  Google Scholar 

  53. Encinar JM, Pardal A, Sánchez N, Nogales S. Biodiesel by transesterification of rapeseed oil using ultrasound: a kinetic study of base-catalysed reactions. Energies. 2018;11:2229.

    Article  Google Scholar 

  54. Xu P, Qiao K, Stephanopoulos G. Engineering oxidative stress defense pathways to build a robust lipid production platform in Yarrowia lipolytica. Biotechnol Bioeng. 2017;114:1521–30.

    Article  CAS  PubMed  Google Scholar 

  55. Liu L, Markham K, Blazeck J, Zhou N, Leon D, Otoupal P, Alper HS. Surveying the lipogenesis landscape in Yarrowia lipolytica through understanding the function of a Mga2p regulatory protein mutant. Metab Eng. 2015;31:102–11.

    Article  PubMed  Google Scholar 

  56. Yan FX, Dong GR, Qiang S, Niu YJ, Hu CY, Meng YH. Overexpression of Δ12, Δ15-desaturases for enhanced lipids synthesis in Yarrowia lipolytica. Front Microbiol. 2020;11:289.

    Article  PubMed  PubMed Central  Google Scholar 

  57. Rakicka M, Lazar Z, Dulermo T, Fickers P, Nicaud JM. Lipid production by the oleaginous yeast Yarrowia lipolytica using industrial by-products under different culture conditions. Biotechnol Biofuels. 2015;8:104.

    Article  PubMed  PubMed Central  Google Scholar 

  58. Tsigie YA, Wang CY, Kasim NS, Diem QD, Huynh LH, Ho QP, Truong CT, Ju YH. Oil production from Yarrowia lipolytica Po1g using rice bran hydrolysate. J Biomed Biotechnol. 2012;2012: 378384.

    Article  PubMed  PubMed Central  Google Scholar 

  59. Juszczyk PRW, Kita A, Rywińska A. Biomass production by Yarrowia lipolytica yeast using waste derived from the production of ethyl esters of polyunsaturated fatty acids of flaxseed oil. Ind Crops Prod. 2019;138: 111590.

    Article  CAS  Google Scholar 

  60. Papanikolaou S, Chevalot I, Komaitis M, Aggelis G, Marc I. Kinetic profile of the cellular lipid composition in an oleaginous Yarrowia lipolytica capable of producing a cocoa-butter substitute from industrial fats. Antonie Van Leeuwenhoek. 2001;80:215–24.

    Article  CAS  PubMed  Google Scholar 

  61. Wojtatowicz M, Rymowicz W, Kautola H. Comparison of different strains of the yeast Yarrowia lipolytica for citric acid production from glucose hydrol. Appl Biochem Biotechnol. 1991;31:165–74.

    Article  CAS  PubMed  Google Scholar 

  62. Mironczuk AM, Biegalska A, Dobrowolski A. Functional overexpression of genes involved in erythritol synthesis in the yeast Yarrowia lipolytica. Biotechnol Biofuels. 2017;10:77.

    Article  PubMed  PubMed Central  Google Scholar 

  63. Nicaud JM, Fabre E, Gaillardin C. Expression of invertase activity in Yarrowia lipolytica and its use as a selective marker. Curr Genet. 1989;16:253–60.

    Article  CAS  PubMed  Google Scholar 

  64. Schmittgen TD, Livak KJ. Analyzing real-time PCR data by the comparative C(T) method. Nat Protoc. 2008;3:1101–8.

    Article  CAS  PubMed  Google Scholar 

  65. Browse J, McCourt PJ, Somerville CR. Fatty acid composition of leaf lipids determined after combined digestion and fatty acid methyl ester formation from fresh tissue. Anal Biochem. 1986;152:141–5.

    Article  CAS  PubMed  Google Scholar 

Download references

Acknowledgements

Not applicable.

Funding

This work was financially supported by the National Science Center, Poland, project UMO-2017/26/E/NZ9/00975. The APC/BPC is financed by Wroclaw University of Environmental and Life Sciences.

Author information

Author notes

  1. Aleksandra M. Mirończuk & Adam Dobrowolski

    Present address: Laboratory for Biosustainability, Institute of Environmental Biology, Wrocław University of Environmental and Life Sciences, Wrocław, Poland

Authors and Affiliations

  1. Department of Biotechnology and Food Microbiology, Wrocław University of Environmental and Life Sciences, 37 Chełmońskiego Street, 51-630, Wrocław, Poland

    Katarzyna Drzymała-Kapinos, Aleksandra M. Mirończuk & Adam Dobrowolski

Authors
  1. Katarzyna Drzymała-Kapinos
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Aleksandra M. Mirończuk
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Adam Dobrowolski
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

KDK carried out the majority of the experimental work, analyzed the results and drafted the manuscript, AMM contributed to data analysis and interpretation, and revision of the manuscript, AD contributed to the conceptualization and design of the experiments, interpretation of the data, revision of the manuscript and funding acquisition. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Adam Dobrowolski.

Ethics declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

Additional file 1: Figure S1.

Schematic for xylose utilization pathway and new Y. lipolytica strains preparation. Table S1. Strains and plasmids used in presented study. Table S2. List of primers.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Drzymała-Kapinos, K., Mirończuk, A.M. & Dobrowolski, A. Lipid production from lignocellulosic biomass using an engineered Yarrowia lipolytica strain. Microb Cell Fact 21, 226 (2022). https://doi.org/10.1186/s12934-022-01951-w

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s12934-022-01951-w

Keywords

Microbial Cell Factories

ISSN: 1475-2859

Contact us