1. Introduction
For decades microbial oil has been regarded as an important substitute for oils derived from plants. Microbial oil was generated commercially for the first time in 1985 but was found to be too expensive compared to the plant source [1]. Later, microbial oil was regarded as an important generator for the production of novel fatty acids that were difficult to be generated via agricultural means [1]. Many oleaginous microorganisms have been reported to produce polyunsaturated fatty acids (PUFAs) that cannot be synthesized by the human body [2,3,4]. Thus, lipogenic fungi are considered vital sources of ω- 6 and ω- 3 PUFAs, which have substantial economic benefit [5,6]. The medicinal significance of PUFAs has been investigated intensively in the past decades [7]. Together, these factors have contributed to the emergence of microbial oil being valuable in terms of industry and economics. Moreover, microbial lipid production is becoming attractive because of its extensive utilization as a biofuel and nutraceutical source [8,9,10,11].
In microbial oil-producing organisms like fungi, microalgae, yeast, and bacteria, lipid accumulation takes place under nitrogen depletion and carbon excess conditions [12]. Mucor circinelloides is deemed an exemplary oleaginous filamentous fungus for the accumulation of lipids because of the existence of genomic tools, known genomic data, and its capability to generate PUFAs rich in γ-linolenic acid (GLA, 18:3; n-6). A study about recent molecular tools to genetically manipulate various fungi provides important insights about genetic engineering of M. circinelloides [13].
In M. circinelloides, the mechanism of the accumulation of lipids at biochemical and molecular stages has been unfolded by the application of metabolic flux analysis [14], proteomics [15], genomics [16], the activity analysis of key enzymes [17,18], and the influence of metal, phosphate, and calcium ions [19,20]. The pentose phosphate pathway and malic enzyme (ME; EC 1.1.1.40) have been suggested to be chief NADPH suppliers in M. circinelloides [17,18]. Employing stoichiometric evaluation, it has been proposed that ME has a vital role in various organisms, even though all the NADPH needed for lipid biosynthesis cannot be provided by it [21]. In the elucidation of the biosynthesis of lipids in lipogenic fungi, the determination of this NADPH source is among one of the major problems of metabolism. Key functions of the NADP+-dependent cytosolic malic enzyme have been reported and discussed in various reviews and research articles [1,12,22,23].
Investigation conducted by Zhang et al. [24] showed a 2.5-fold elevation in the accumulation of lipids when the ME gene malEMc was overexpressed. On the other hand, when a specific ME inhibitor called sesamol was supplemented in the culture media of M. circinelloides CBS 108.16 [25], a reduction from 24% to 2% was noted. Furthermore, when the activity of ME diminishes, lipid accumulation is observed to be throttled in M. circinelloides and related fungus-like Mortierella alpina [26]. Rodríguez-Frómeta et al. [17] also overexpressed the ME gene malA in the CBS 277.49 strain of M. circinelloides, but the mutant strain failed to show any elevation in the accumulation of lipids, regardless of the elevation in ME activity and mRNA levels, signifying the presence of other lipid accumulation holdups. Interestingly, in the M. circinelloides genome (
In the current study, we overexpressed two cytosolic ME genes under the control of a strong promoter in the M. circinelloides lipid-overproducing strain WJ11. We named these genes cmalA and cmalB. Though the cmalA gene has been studied in other strains of M. circinelloides, we investigated it for the first time in the WJ11 strain. On the other hand, the cmalB gene has not been investigated before in any strain of M. circinelloides, and to the best of our information, we are investigating it for the first time. We found that overexpression of both genes produced a considerable elevation in ME activity, mRNA levels, and lipid accumulation, although lipid levels increased more significantly in the case of the cmalA gene.
2. Materials and Methods
2.1. Strains, Growth, and Transformation Conditions
cmalA and cmalB genes were amplified from the M. circinelloides WJ11 strain (CCTCC No. M 2014424). In all the transformation experiments, as a recipient strain, M. circinelloides M65, the uracil auxotroph of M. circinelloides WJ11, was used. Electroporation was employed to carry out the transformation [28]. YPG or MMC media was used to grow the cultures at 26 °C [29,30]. When needed, 200 mg/mL of uridine was added to the media. For mycelia and colonial growth, pH was regulated to 4.5 and 3, respectively.
150 mL of Kendrick and Ratledge (K & R) medium [31] was poured into 500 mL baffled flasks, and in it Mc-cmalA-4 (cmalA overexpression) and Mc-cmalB-5 (cmalB overexpression) strains and wild-type control strain (Mc-cmal-2075) were cultivated independently, for 24 h, with 150 rpm shaking at 28 °C. Cultures from these baffled flasks were then utilized as seed culture to be poured in a 2 L fermenter (BioFlo/CelliGen 115, New Brunswick Scientific, Edison, NJ, USA). The 2 L fermenter contained 1.5 L of altered K & R medium with 80 g glucose/L. Conditions maintained in the fermenter were as follows: shaking = 700 rpm, temperature = 28 °C, pH = 6, and aeration = 0.5 air volume per fermenter volume per minute (v/v min−1). 2 M HCL and 2 M NaOH were utilized for automatic pH maintenance. For the propagation and maintenance of the overexpression plasmids, Escherichia coli DH5α cells were used. LB media was used for their growth at 37 °C and 220 rpm shaking [32].
2.2. Plasmid Construction
For constructing cmalA and cmalB overexpression plasmids, two independent pMAT2075 plasmids where used. To enable their chromosomal integration via homologous recombination, they included one kb downstream and upstream CarRP sequences flanking pyrF genes [33]. From the M. circinelloides WJ11 genome, PCR amplicfication of cmalA and cmalB genes was carried out using primes cmalA-1F-XhoI/cmalA-1R-XhoI and cmalB-1F-XhoI/cmalB-1R-XhoI, respectively (Supplementary Table S1). Within these primers, the incorporation of 30 bp homologous sequences of XhoI restriction sites was carried out. To generate plasmids pMAT2075-cmalA and pMAT2075-cmalB, respective PCR fragments were digested with XhoI restriction endonucleases and ligated into plasmid pMAT2075 (One-step cloning kit, Takara, Shiga, Japan). These plasmids were then extracted from E. coli and PCR analysis was conducted using Ch-1F/Ch-1R and Ch-2F/Ch-2R primers, respectively, for cmalA and cmalB genes. Moreover, to confirm the sequence of the genes, DNA sequencing was performed. To release overexpression cmalA and cmalB constructs from their respective plasmids, SmaI was used as the digestive enzyme. The released constructs were then transformed into M65 protoplasts, wherein white colonies were produced because of carRP locus integration, and where integration of the carRP locus did not take place, yellow colonies were generated. The Rodríguez-Frómeta et al. [17] method was used for albino colony selection.
2.3. Preparation of Genomic DNA
For the extraction of DNA, K & R medium was used to grow M. circinelloides at conditions: shaking = 150 rpm, temperature = 28 °C, and time = 3 days. Under reduced pressure, a Büchner funnel was employed to harvest the mycelium. Distilled water was then used thrice to wash this mycelium. A DNA Quick Plant System kit was employed for the extraction of genomic DNA (Tiangen Biotech Co., Ltd., Beijing, China).
2.4. Reverse Transcription-Quantitative PCR (RT-qPCR) to Analyse the Expression of Genes
In a 2 L fermenter Mc-cmalA-4 and Mc-cmalB-5 and control strain Mc-cmal-2075 were cultivated, and subsequently, mycelium was harvested at 6 h, 24 h, 48 h, 72 h, and 96 h to do RT-qPCR analysis. From the collected mycelia, total RNA was extracted by the application of Trizol after grinding mycelia using liquid nitrogen and then transformed into cDNA by using PrimeScriptTM RT reagent kit (Takara) according to instructions of the manufacturer. The quantitative RT-PCR was executed with the help of Light Cycler 96 Instrument (Roche Diagnostics GmbH, Basel, Switzerland) and Maxima SYBR Green qPCR Master Mix (Thermo scientific, Waltham, MA, USA) according to instructions of the manufacturer. Using the method of 2−ΔΔCt and actin as housekeeping gene, data quantification was performed as described previously [34].
2.5. Measurement of the Concentration of Nitrogen and Glucose in Culture Medium
The indophenol method was employed to estimate the concentration of ammonium in the culture medium [35]. The glucose oxidase Perid-test kit (Shanghai Rongsheng Biotech Co., Ltd., Shanghai, China) was employed to determine the concentration of glucose as per the guidelines of the manufacturer.
2.6. Analysis of CDW and Lipid Accumulation
A Büchner funnel was used to harvest the mycelia that had been cultured for 4 d. The harvested mycelia were subsequently washed with distilled water three times. It was then frozen at −80 °C overnight and a freeze dryer was employed to dry up the mycelia. Then, the cell dry weight (CDW) was calculated gravimetrically [29] and a slightly modified Folch method [36] was employed to perform lipid extraction, which was followed by fatty acid methyl esters (FAME) analysis using gas chromatography (GC). FAME was prepared in tubes using 1 mL of 10% methanol in HCl and kept for 3 h at 60 °C in a water bath. 1 mL of NaCl (saturated) and 2 mL of hexane were utilized to extract methyl esters. After this, a vertical 360 tube rotator was utilized to rotate the tubes for 1 h. This was followed by vortexing and centrifugation at 3000 rpm for 5 min. GC analysis was performed on the top organic hexane layer containing methyl esters. Detectors for flame ionization were incorporated into the gas chromatography machine. A fatty acid standard (Supelco® 37 Component FAME Mix) was used to identify individual chromatographic peaks, and respective chromatographic peaks corresponded to respective fatty acids [37].
2.7. Determination of ME Activity
The Hsu and Lardy method [38] was used to determine ME activity with minor modifications. Extraction buffer maintained at pH 7.5, comprising of 50 mM Tris–HCL and 20% glycerol (w/w) was used during the process.
2.8. Statistical Analysis
Three independent experiments were carried out to obtain mean values. Multiple comparison tests using GraphPad Prism (version 7, San Diego, CA, USA) preceded by one-way/two-way ANOVA (wherever applicable) was employed to perform statistical analysis. p < 0.05 was considered significantly different.
3. Results
3.1. Genetic Engineering for Generation of cmalA and cmalB Overexpressing M. circinelloides Strains
The cmalA (scaffold00036.12) and cmalB (scaffold00049.37) genes were retrieved from M. circinelloides WJ11 genomic data. Using plasmid pMAT2075 (carrying the strong promoter zrt1 and pyrF gene as a selectable marker) [28], the overexpression recombinant strains of cmalA (pMAT2075-cmalA) and cmalB (pMAT2075-cmalB) genes were generated to assess their role in lipid production. The M65 strain (M. circinelloides WJ11 uracil auxotroph strain) was used as the recipient strain to transform pMAT2075-cmalA and pMAT2075-cmalB. Uridine auxotrophy in the M65 strain was complemented by the pyrF gene present in the plasmids. Mc-cmalA-4 and Mc-cmalB-5 were chosen as independent transformants, respectively, for the cmalA and cmalB genes. The Mc-cmal-2075 strain, having wild-type features, was used as the control strain.
Verification of the integration of transforming fragments into the M. circinelloides genome was conducted with the help of PCR. The amplification of the 2033 bp band with the primer pair 2F and 2R indicated the integration of the cmalA gene in the carRP locus of M65 (Figure 1b and Figure 2a). Similarly, the amplification of the 2032 bp band with the primer pair 3F and 3R indicated the integration of the cmalB gene in the carRP locus of M65 (Figure 1c and Figure 2b). Moreover, the integration of the pyrF gene into the M. circinelloides genome in Mc-cmalA-4 and Mc-cmalB-5 strains was confirmed by PCR amplification with the primer pairs 1F and 1R producing a band size of 1531 bp (Figure 1b,c and Figure 2c). Altogether, these outcomes authenticated the incorporation of the cmalA and cmalB genes into the carRP locus of the M. circinelloides genome under the regulation of the strong promoter pzrt1 in the Mc-cmalA-4 and Mc-cmalB-5 recombinant strains.
3.2. The Expression Levels of cmalA and cmalB Genes in Overexpressing Strains
To evaluate and compare the mRNA levels of the cmalA and cmalB genes in the respective Mc-cmalA-4 and Mc-cmalB-5 strains and the control strain Mc-cmal-2075, reverse transcription quantitative PCR (RT-qPCR) was performed. It was carried out at 6, 24, 48, 72, and 96 h of growth using primer pairs cmalA-3-F/cmalA-3-R and cmalB-3-F/cmalB-3-R for the cmalA and cmalB genes, respectively (Supplementary File Table S1). In comparison to the control strain, mRNA levels of the cmalA and cmalB genes in Mc-cmalA-4 and Mc-cmalB-5 overexpression strains were found to be considerably elevated. The level of expression at 24 h was observed to be 6.58- and 4.78-fold for Mc-cmalA-4 and Mc-cmalB-5 strains. A decrease in the trend was noted after 24 h, but elevation in comparison to the control strain was maintained throughout the process of fermentation, validating the overexpression of respective genes in the recombinant strains (Figure 3). During expression analysis across various specified hours, MC-cmal-2075 was used as reference and its relative expression value was taken as 1.
3.3. ME Activity Analysis of cmalA and cmalB Overexpressing Strains
The trend of specific ME activity in overexpressing strains (Mc-cmalA-4 and Mc-cmalB-5) compared to the control strain (Mc-cmal-1557) at 24, 48, 72, and 96 h is shown in Figure 4. Both overexpressing strains showed higher specific ME activity than the control strain throughout the whole culture. In addition, a consistent increase in specific ME activity for both overexpressing strains was observed up to 72 h of culture, whereas the control strain showed a reduction in this activity as the culture progressed. At 72 h of culture, the Mc-cmalA-4 and Mc-cmalB-5 strains showed a 9.8- and 6.4-fold elevation in specific ME activity, respectively, compared to the control strain. Despite the fact that a dip in ME activity in overexpressing strains was observed at 96 h, it was noted to be greater than in the control strain.
3.4. Cell Growth and Lipid Accumulation in cmalA and cmalB Overexpressing Strains
The effect of cmalA and cmalB overexpression on cell dry weight (CDW), lipid accumulation, and ammonium and glucose concentrations were analysed (Figure 5) in 96-h cultures. CDW increased with the incubation, and it was found to be maximum at 96 h for the overexpressing and control strains (Figure 5c). The concentration of glucose was found to be sufficient throughout the fermentation process, while the rate of consumption of glucose was found to be slightly faster in overexpression strains compared to the control strain (Figure 5a). However, nitrogen was depleted from the medium between 12 and 24 h, while lipid accumulation increased (Figure 5b). In the Mc-cmalA-4 and Mc-cmalB-5 strain, compared to the control strain Mc-cmal-2075, at 72 h, lipid accumulation was increased by 23.2% (from 34% in the control strain to 42% in the overexpressing strain) and 5.88% (from 34% in the control strain to 36% in the overexpressing strain), respectively (Figure 5d).
The fatty acid profiles of the recombinant and control strains are shown in Table 1. At 96 h, the GLA content in total fatty acids of Mc-cmal- 2075, Mc-cmalA-4, and Mc-cmalB-5 strains was 9.91%, 9.88%, and 10.35%, respectively. Since overexpression strains generated more lipid content than the control strain, the overall GLA content in the biomass of overexpression strains was noted to be higher than in the control strain. GLA content in biomass was noted to be elevated by 11.42% (from 3.5% to 3.9%) and 16.85% (from 3.5% to 4.09%) in Mc-cmalA-4 and Mc-cmalB-5 strains, respectively.
4. Discussion
The conditions of the culture and levels of nutrients are deemed to be important influencers for lipid accumulation. Commercially proficient lipid production necessitates the development of recombinant strains that can accumulate elevated amounts of lipids, independent of the environmental fluctuations. So far, several genetic engineering techniques have been employed to boost lipid production via overexpression or the deletion of important genes and transporters such as malate and citrate transporters [39,40,41,42,43], AMP-activated protein kinase [44,45], the pentose phosphate pathway [18,46,47], Acyl CoA: Diacylglycerol Acyltransferase [48], lipases [49,50,51], malic enzymes [17,24], and the utilization of lignocellulosic biomasses [52].
When ME genes were overexpressed in M. circinelloides, lipid accumulation was found to be enhanced 2.5-fold [24], while Rodríguez-Frómeta et al. [17] found no increase in the accumulation of lipids. The difference between these two studies was that the former utilized a self-replicative plasmid resulting in the generation of an unstable and industrially infeasible strain, whereas the latter generated M. circinelloides strains that were genetically stable because of the incorporation of the ME gene in the genome’s precise locus. Rodríguez-Frómeta et al. [17] developed an important scheme for the replacement of the gene, preventing the biosynthesis of carotenoids so that their strain could be utilized for the production of a biomass suitable for biodiesel transformation. This replacement strategy generated transformants of two types; yellow transformants in which the gene did not integrate at the desired location and white transformants lacking carotene with gene integration at the desired location. This approach, therefore, added an important tool for M. circinelloides research and for probable applications in biotechnology. In our study, we have also utilized the method of Rodríguez-Frómeta et al. [17]. To give an insight into different results produced during the works of Zhang et al. [24] and Rodríguez-Frómeta et al. [17], Hao et al. [53] overexpressed the ME gene malE1 in a related oleaginous fungus Mortierella alpina, and found that the content of fatty acid was elevated by 30%. Their results suggested that in oleaginous fungi, ME has an important role in the biosynthesis of fatty acids but is not the only rate-limiting enzyme. The ME gene (GenBank: DQ975377.1) from M. circinelloides has also been incorporated and overexpressed in Rhodotorula glutinis, an oleaginous yeast, and it has resulted in the elevation of lipid accumulation by 18.74% [54]. Other studies have shown that ME is not the major enzyme responsible for NADPH generation, but it does have a role in the synthesis of fatty acids [14,55]. In the current study, we have demonstrated that lipid accumulation increased when cmalA and cmalB genes were put under the regulation of a strong promoter zrt1. In the case of the cmalA gene, lipid accumulation was elevated by 23.2%, while for cmalB there was an elevation of 5.8%. By observing the trend of ME activity at 96 h, it becomes clear that some factors limit its persistent activity. It is hypothesized that the ME-cleaving enzyme may be responsible for the loss of activity [24].
The overexpression of cmalA and cmalB genes elevated the lipid content of the cell dry weight by 23.2% (from 34% to 42%) and 5.88% (from 34% to 36%), respectively, even though the activity of ME was elevated by 9.8- and 6.4-fold in respective genes. These outcomes indicate that the enhanced activity of ME did not necessarily result in an adequate elevation in NADPH levels required for the synthesis of fatty acids. Alternately, other pathways of metabolism may provide NADPH for lipid production. Thus, it is probable that the role of ME in the synthesis of fatty acids, perhaps, would not be as vital as suggested earlier [1,24]. Several researchers have exhibited that, among some yeasts or molds, ME overexpression considerably boosts the accumulation of lipids [24,53,54]. The ME recombinant strain of bacteria Rhodococcus josti, has been found to enhance lipid accumulation [56]. In microalgae, the mitochondrial malic enzyme of P. tricornutum, was found to significantly influence the accumulation of lipids in P. tricornutum and C. pyrenoidosa [57,58,59]. C. protothecoides and Nannochloropsis salina has also shown elevation in lipid synthesis when their malic enzymes were overexpressed [60].
Among plant cells, there is a lot of ambiguity about the effect of ME activity on accumulation of lipids. In the case of plants, ME is considered an ubiquitous enzyme having a role in various metabolic pathways such as photosynthesis, stress responses, and the development and growth of plants [61,62,63]; although, various roles of ME in plants still remain unidentified [64]. Hence, it may be difficult to find the gene or genes encoding for ME, having a role in NADPH generation aimed at the biosynthesis of fatty acids. However, a study on the development of embryos in rapeseed has shown that ME may have a role in the synthesis of fatty acids [65].
5. Conclusions
This is the first study to investigate the role played in the accumulation of lipids by cytosolic ME genes cmalA and cmalB in the M. circinelloides WJ11 strain, though previously the cmalA gene has been evaluated for its role in the accumulation of lipids in the CBS 277.49 strain. The study of cytosolic ME genes in the WJ11 strain revealed that the cmalA gene has a more significant role in the accumulation of lipids compared to the cmalB gene. In the case of both the genes, the activity of ME is found to be significantly reduced in the end. In conclusion, it can be said that, like some highly competent oleaginous yeasts which retain the activity of ME for a long time, M. circinelloides, if engineered to repel ME degradation, can also accumulate a high amount of lipid (~70%). If our study is analyzed within the prism of previous studies, our results further emphasize that ME is not the only rate-limiting enzyme, though it has a significant role in the synthesis of fatty acids.
Conceptualization, data generation, analysis, interpretation, and writing initial draft, A.B.A.F.; data generation, analysis, interpretation, and writing, A.M.S. and T.N.; data generation and analysis, S.N. and W.Y.; data curation and editing, V.G. and Y.M.; writing and funding acquisition, M.K.A.-S.; conceptualization, resources, funding acquisition, writing, and editing, Y.S. All authors have read and agreed to the published version of the manuscript.
This work was supported by the National Natural Science Foundation of China (grant 31972851617 and 31670064), TaiShan Industrial Experts Programme (tscy 20160101), and Researchers Supporting Project Number (RSP2022R410), King Saud University, Riyadh, Saudi Arabia.
Not applicable.
Not applicable.
All relevant data generated or analyzed during this study are included in this article.
The laboratory support provided by Khalid M Fazili, Department of Biotechnology for carrying out some of the experiments is greatly appreciated. We acknowledge the support in terms of infra structural facilities made available by the Department of Biotechnology, University of Kashmir, Srinagar. The authors would also like to extend their sincere appreciation to the Researchers Supporting Project Number (RSP2022R410), King Saud University, Riyadh, Saudi Arabia. We thank Syed Nahida, Junhuan Yang, Samee Ullah and Yusuf Nazir for supporting us during this work.
The authors declare no conflict of interest.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Enlarge this image.Figure 1. Overexpression of cmalA and cmalB genes. (a) Plasmid structure of plasmid pMAT2075. (b) Plasmid structure of pMAT2075-cmalA in Mc-cmalA-4 strain. Positions for primers 1F, 1R, 2F, and 2R are indicated. (c) Plasmid structure of pMAT2075-cmalB in Mc-cmalB-5 strain. Positions for primers 1F, 1R, 3F, and 3R are indicated.
Enlarge this image.Figure 2. PCR amplification in Mc-cmal-2075, Mc-cmalA-4, and Mc-cmalB-5 strains. (a) PCR amplification of cmalA gene and 3′ carRP region using primer pair 2F/2R in overexpressing transformant Mc-cmalA-4 strain. Lane M: Marker. Lane 1 represents the control, Lane 2 shows the result of the PCR amplification that confirmed the presence of cmalA gene in the overexpressing transformant. (b) PCR amplification of cmalB gene and 3′ carRP region using primer pair 3F/3R in overexpressing transformant Mc-cmalB-5 strain. Lane M: Marker, Lane 1 represents the control, Lane 2 shows the result of the PCR amplification that confirmed the presence of cmalB gene in the overexpressing transformant. (c) PCR amplification of pyrF genes and 5′ carRP regions in Mc-cmal-2075, Mc-cmalA-4, and Mc-cmalB-5 strains using primer pairs 1F/1R. Lane M: Marker. Lane 1 represents the control. Lanes 2 and 3 show presence of pyrF gene in Mc-cmalA-4 and Mc-cmalB-5 strains, respectively.
Enlarge this image.Figure 3. Levels of expression of cmalA and cmalB genes in control strain Mc-cmal-2075 and overexpressing strains Mc-cmalA-4 and Mc-cmalB-5. Values are the mean of three biological replicates. Error bars represent the standard error of the mean. Asterisks indicate significant differences: * p < 0.05 and ** p < 0.01.
Enlarge this image.Figure 4. Specific ME activity of the control strain Mc-cmal-2075 and overexpressing strains Mc-cmalA-4 and Mc-cmalB-5. Values are the mean of three biological replicates. Error bars represent the standard error of the mean. Asterisks indicate significant differences: * p < 0.05, ** p < 0.01, *** p < 0.001.
Enlarge this image.Figure 5. Determination of cell growth of lipid content analysis of recombinant strains (Mc-cmalA-4 and Mc-cmalB-5) and control strain (Mc-cmal-2075) at specific time points. (a) Concentration of glucose, (b) concentration of ammonium, (c) cell dry weight (CDW), (d) lipid content. Values are the mean of three biological replicates. Error bars represent the standard error of the mean. Asterisks indicate significant differences: * p < 0.05 and ** p < 0.01.
Composition of fatty acids (%, w/w of total fatty acids) * in Mc-cmal- 2075, Mc-cmalA-4, and Mc-cmalB-5 strains.
| Strains | Time (h) | 16:0 | 16:1 | 18:0 | 18:1 | 18:2 (LA) | 18:3 (GLA) |
|---|---|---|---|---|---|---|---|
| Mc-cmal-2075 | 24 | 17.44 ± 0.3 | 2.61 ± 0.2 | 4.90 ± 0.1 | 39.10 ± 0.9 | 11.01 ± 0.8 | 13.59 ± 0.4 |
| 48 | 20.18 ± 0.5 | 2.92 ± 0.2 | 5.18 ± 0.3 | 41.12 ± 1.0 | 10.74 ± 0.6 | 10.80 ± 0.2 | |
| 72 | 22.55 ± 0.7 | 3.41 ± 0.1 | 4.62 ± 0.3 | 41.88 ± 2.1 | 11.09 ± 0.6 | 9.76 ± 0.3 | |
| 96 | 22.59 ± 0.5 | 4.00 ± 0.3 | 3.93 ± 0.1 | 42.94 ± 1.5 | 12.41 ± 0.3 | 9.91 ± 05 | |
| Mc-cmalA-4 | 24 | 17.96 ± 0.4 * | 1.92 ± 0.2 * | 4.89 ± 0.9 | 35.37 ± 0.8 * | 11.48 ± 0.5 ** | 12.72 ± 0.7 * |
| 48 | 21.72 ± 0.4 ** | 2.57 ± 0.1 * | 5.01 ± 0.6 * | 40.38 ± 1.5 ** | 10.45 ± 0.3 * | 10.68 ± 0.2 ** | |
| 72 | 21.00 ± 1.1 * | 2.79 ± 0.2 ** | 4.46 ± 0.3 | 41.00 ± 1.5 ** | 10.56 ± 0.2 | 9.05 ± 0.6 * | |
| 96 | 22.95 ± 0.7 ** | 3.59 ± 0.1 * | 4.18 ± 0.2 * | 41.97 ± 1.3 * | 11.36 ± 0.3 | 9.88 ± 0.6 * | |
| Mc-cmalB-5 | 24 | 18.95 ± 0.7 * | 2.14 ± 0.3 * | 4.88 ± 0.4 | 39.20 ± 0.7 * | 12.92 ± 0.2 * | 13.49 ± 0.4 * |
| 48 | 21.82 ± 0.6 ** | 2.99 ± 0.2 * | 6.95 ± 0.4 * | 43.74 ± 1.3 * | 11.89 ± 0.3 ** | 10.89 ± 0.3 ** | |
| 72 | 22.75 ± 0.9 ** | 3.23 ± 0.4 | 5.31 ± 0.3 | 45.06 ± 1.5 ** | 11.45 ± 0.4 ** | 10.30 ± 0.2 * | |
| 96 | 23.66 ± 0.2 *** | 4.89 ± 0.3 * | 4.94 ± 0.2 * | 45.10 ± 1.8 * | 12.26 ± 0.1 | 10.35 ± 0.4 * |
* The fatty acid composition displayed at different time point. The values are means ± standard deviations of three independent experiments. Asterisks indicate significant differences: * p < 0.05, ** p < 0.01; *** p < 0.001.
Supplementary Materials
The following supporting information can be downloaded at:
References
1. Ratledge, C. Fatty acid biosynthesis in microorganisms being used for Single Cell Oil production. Biochimie; 2004; 86, pp. 807-815. [DOI: https://dx.doi.org/10.1016/j.biochi.2004.09.017]
2. Khan, A.K.; Yang, J.; Hussain, S.A.; Zhang, H.; Garre, V.; Song, Y. Genetic Modification of Mucor circinelloides to Construct Stearidonic Acid Producing Cell Factory. Int. J. Mol. Sci.; 2019; 20, 1683. [DOI: https://dx.doi.org/10.3390/ijms20071683] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30987311]
3. Yazawa, H.; Iwahashi, H.; Kamisaka, Y.; Kimura, K.; Aki, T.; Ono, K.; Uemura, H. Heterologous Production of Dihomo-γ-Linolenic Acid in Saccharomyces cerevisiae. Appl. Environ. Microbiol.; 2007; 73, pp. 6965-6971. [DOI: https://dx.doi.org/10.1128/AEM.01008-07] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17873077]
4. AbdElrazak, A.; Ward, A.; Glassey, J. Response surface methodology for optimising the culture conditions for eicosapentaenoic acid production by marine bacteria. J. Ind. Microbiol. Biotechnol.; 2013; 40, pp. 477-487. [DOI: https://dx.doi.org/10.1007/s10295-013-1238-x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23455698]
5. Christophe, G.; Fontanille, P.; Larroche, C. Research and Production of Microbial Polyunsaturated Fatty Acids. Bioprocess Biomol. Prod.; 2019; 1, pp. 167-194. [DOI: https://dx.doi.org/10.1002/9781119434436.ch8]
6. Wang, W.-J.; Huang, H.; Ji, X.-J. Systems and Synthetic Biotechnology for the Production of Polyunsaturated Fatty Acids. Systems and Synthetic Biotechnology for Production of Nutraceuticals; Springer: Singapore, 2019; pp. 189-202. [DOI: https://dx.doi.org/10.1007/978-981-15-0446-4_8]
7. Nazir, Y.; Halim, H.; Prabhakaran, P.; Ren, X.; Naz, T.; Mohamed, H.; Nosheen, S.; Mustafa, K.; Yang, W.; Hamid, A.A. et al. Different Classes of Phytohormones Act Synergistically to Enhance the Growth, Lipid and DHA Biosynthetic Capacity of Aurantiochytrium sp. SW1. Biomolecules; 2020; 10, 755. [DOI: https://dx.doi.org/10.3390/biom10050755]
8. Liang, M.H.; Jiang, J.-G. Advancing oleaginous microorganisms to produce lipid via metabolic engineering technology. Prog. Lipid Res.; 2013; 52, pp. 395-408. [DOI: https://dx.doi.org/10.1016/j.plipres.2013.05.002]
9. Li-Beisson, Y.; Peltier, G. Third-generation biofuels: Current and future research on microalgal lipid biotechnology. OCL; 2013; 20, D606. [DOI: https://dx.doi.org/10.1051/ocl/2013031]
10. Runguphan, W.; Keasling, J.D. Metabolic engineering of Saccharomyces cerevisiae for production of fatty acid-derived biofuels and chemicals. Metab. Eng.; 2014; 21, pp. 103-113. [DOI: https://dx.doi.org/10.1016/j.ymben.2013.07.003]
11. Ratledge, C. Microbial production of polyunsaturated fatty acids as nutraceuticals. Microbial Production of Food Ingredients, Enzymes and Nutraceuticals; Elsevier: Amsterdam, The Netherlands, 2013; pp. 531-558. [DOI: https://dx.doi.org/10.1533/9780857093547.2.531]
12. Ratledge, C. Microbial lipids. Biotechnology; 2nd ed. Wiley: New York, NY, USA, 1997; Volume 7, pp. 133-197.
13. Mohamed, H.; Naz, T.; Yang, J.; Shah, A.M.; Nazir, Y.; Song, Y. Recent Molecular Tools for the Genetic Manipulation of Highly Industrially Important Mucoromycota Fungi. J. Fungi; 2021; 7, 1061. [DOI: https://dx.doi.org/10.3390/jof7121061]
14. Zhao, L.; Zhang, H.; Wang, L.; Chen, H.; Chen, Y.Q.; Chen, W.; Song, Y. 13 C-metabolic flux analysis of lipid accumulation in the oleaginous fungus Mucor circinelloides. Bioresour. Technol.; 2015; 197, pp. 23-29. [DOI: https://dx.doi.org/10.1016/j.biortech.2015.08.035] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26318243]
15. Tang, X.; Zan, X.; Zhao, L.; Chen, H.; Chen, Y.Q.; Chen, W.; Song, Y.; Ratledge, C. Proteomics analysis of high lipid-producing strain Mucor circinelloides WJ11: An explanation for the mechanism of lipid accumulation at the proteomic level. Microb. Cell Fact.; 2016; 15, 35. [DOI: https://dx.doi.org/10.1186/s12934-016-0428-4] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26867592]
16. Tang, X.; Zhao, L.; Chen, H.; Chen, Y.Q.; Chen, W.; Song, Y.; Ratledge, C. Complete Genome Sequence of a High Lipid-Producing Strain of Mucor circinelloides WJ11 and Comparative Genome Analysis with a Low Lipid-Producing Strain CBS 277.49. PLoS ONE; 2015; 10, e0137543. [DOI: https://dx.doi.org/10.1371/journal.pone.0137543] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26352831]
17. Rodríguez-Frómeta, R.A.; Gutiérrez, A.; Torres-Martínez, S.; Garre, V. Malic enzyme activity is not the only bottleneck for lipid accumulation in the oleaginous fungus Mucor circinelloides. Appl. Microbiol. Biotechnol.; 2013; 97, pp. 3063-3072. [DOI: https://dx.doi.org/10.1007/s00253-012-4432-2] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23053085]
18. Zhao, L.; Tang, X.; Luan, X.; Chen, H.; Chen, Y.Q.; Chen, W.; Song, Y.; Ratledge, C. Role of pentose phosphate pathway in lipid accumulation of oleaginous fungus Mucor circinelloides. RSC Adv.; 2015; 5, pp. 97658-97664. [DOI: https://dx.doi.org/10.1039/C5RA20364C]
19. Dzurendova, S.; Zimmermann, B.; Kohler, A.; Reitzel, K.; Nielsen, U.; Dupuy-Galet, B.; Leivers, S.; Horn, S.; Shapaval, V. Calcium Affects Polyphosphate and Lipid Accumulation in Mucoromycota Fungi. J. Fungi; 2021; 7, 300. [DOI: https://dx.doi.org/10.3390/jof7040300]
20. Dzurendova, S.; Zimmermann, B.; Tafintseva, V.; Kohler, A.; Horn, S.J.; Shapaval, V. Metal and Phosphate Ions Show Remarkable Influence on the Biomass Production and Lipid Accumulation in Oleaginous Mucor circinelloides. J. Fungi; 2020; 6, 260. [DOI: https://dx.doi.org/10.3390/jof6040260]
21. Ratledge, C. The role of malic enzyme as the provider of NADPH in oleaginous microorganisms: A reappraisal and unsolved problems. Biotechnol. Lett.; 2014; 36, pp. 1557-1568. [DOI: https://dx.doi.org/10.1007/s10529-014-1532-3]
22. Botham, P.A.; Ratledge, C. A Biochemical Explanation for Lipid Accumulation in Candida 107 and Other Oleaginous Micro-organisms. J. Gen. Microbiol.; 1979; 114, pp. 361-375. [DOI: https://dx.doi.org/10.1099/00221287-114-2-361]
23. Ratledge, C.; Wynn, J.P. The Biochemistry and Molecular Biology of Lipid Accumulation in Oleaginous Microorganisms. Adv. Appl. Microbiol.; 2002; 51, pp. 1-52. [DOI: https://dx.doi.org/10.1016/s0065-2164(02)51000-5]
24. Zhang, Y.; Adams, I.P.; Ratledge, C. Malic enzyme: The controlling activity for lipid production? Overexpression of malic enzyme in Mucor circinelloides leads to a 2.5-fold increase in lipid accumulation. Microbiology; 2007; 153, pp. 2013-2025. [DOI: https://dx.doi.org/10.1099/mic.0.2006/002683-0]
25. Wynn, J.P.; Kendrick, A.; Ratledge, C. Sesamol as an inhibitor of growth and lipid metabolism in Mucor circinelloides via its action on malic enzyme. Lipids; 1997; 32, pp. 605-610. [DOI: https://dx.doi.org/10.1007/s11745-997-0077-1]
26. Wynn, J.P.; Hamid, A.B.A.; Ratledge, C. The role of malic enzyme in the regulation of lipid accumulation in filamentous fungi. Microbiology; 1999; 145, pp. 1911-1917. [DOI: https://dx.doi.org/10.1099/13500872-145-8-1911]
27. Vongsangnak, W.; Zhang, Y.; Chen, W.; Ratledge, C.; Song, Y. Annotation and analysis of malic enzyme genes encoding for multiple isoforms in the fungus Mucor circinelloides CBS 277.49. Biotechnol. Lett.; 2012; 34, pp. 941-947. [DOI: https://dx.doi.org/10.1007/s10529-012-0859-x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22367279]
28. Yang, J.; Khan, A.K.; Nosheen, S.; Nazir, M.Y.B.M.; Zhang, H.; Garre, V.; Song, Y. Improved SDA Production in High Lipid Accumulating Strain of Mucor circinelloides WJ11 by Genetic Modification. Am. J. Biochem. Biotechnol.; 2020; 16, pp. 138-147. [DOI: https://dx.doi.org/10.3844/ajbbsp.2020.138.147]
29. Hussain, S.A.; Hameed, A.; Khan, A.K.; Zhang, Y.; Zhang, H.; Garre, V.; Song, Y. Engineering of Fatty Acid Synthases (FASs) to Boost the Production of Medium-Chain Fatty Acids (MCFAs) in Mucor circinelloides. Int. J. Mol. Sci.; 2019; 20, 786. [DOI: https://dx.doi.org/10.3390/ijms20030786]
30. Nicolas, F.E.; De Haro, J.P.; Torres-Martínez, S.; Ruiz-Vázquez, R.M. Mutants defective in a Mucor circinelloides dicer-like gene are not compromised in siRNA silencing but display developmental defects. Fungal Genet. Biol.; 2007; 44, pp. 504-516. [DOI: https://dx.doi.org/10.1016/j.fgb.2006.09.003]
31. Kendrick, A.; Ratledge, C. Desaturation of polyunsaturated fatty acids in Mucor circinelloides and the involvement of a novel membrane-bound malic enzyme. Eur. J. Biochem.; 1992; 209, pp. 667-673. [DOI: https://dx.doi.org/10.1111/j.1432-1033.1992.tb17334.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/1425673]
32. Hanahan, D. Studies on transformation of Escherichia coli with plasmids. J. Mol. Biol.; 1983; 166, pp. 557-580. [DOI: https://dx.doi.org/10.1016/S0022-2836(83)80284-8]
33. Zhao, L.; Cánovas-Márquez, J.T.; Tang, X.; Chen, H.; Chen, Y.Q.; Chen, W.; Garre, V.; Song, Y.; Ratledge, C. Role of malate transporter in lipid accumulation of oleaginous fungus Mucor circinelloides. Appl. Microbiol. Biotechnol.; 2016; 100, pp. 1297-1305. [DOI: https://dx.doi.org/10.1007/s00253-015-7079-y]
34. Naz, T.; Yang, J.; Nosheen, S.; Sun, C.; Nazir, Y.; Mohamed, H.; Fazili, A.B.A.; Ullah, S.; Li, S.; Yang, W. et al. Genetic Modification of Mucor circinelloides for Canthaxanthin Production by Heterologous Expression of β-carotene Ketolase Gene. Front. Nutr.; 2021; 8, 756218. [DOI: https://dx.doi.org/10.3389/fnut.2021.756218] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34722614]
35. Chaney, A.L.; Marbach, E.P. Modified Reagents for Determination of Urea and Ammonia. Clin. Chem.; 1962; 8, pp. 130-132. [DOI: https://dx.doi.org/10.1093/clinchem/8.2.130] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/13878063]
36. Folch, J.; Lees, M.; Stanley, G.H.S. A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem.; 1957; 226, pp. 497-509. [DOI: https://dx.doi.org/10.1016/S0021-9258(18)64849-5]
37. Shah, A.M.; Mohamed, H.; Zhang, Z.; Song, Y. Isolation, characterization and fatty acid analysis of Gilbertella persicaria DSR1: A potential new source of high value single-cell oil. Biomass-Bioenergy; 2021; 151, 106156. [DOI: https://dx.doi.org/10.1016/j.biombioe.2021.106156]
38. Hsu, R.Y.; Lardy, H.A. Pigeon liver malic enzyme: II. Isolation, crystallization, and some properties. J. Biol. Chem.; 1967; 242, pp. 520-526. [DOI: https://dx.doi.org/10.1016/S0021-9258(18)96304-0]
39. Athenaki, M.; Gardeli, C.; Diamantopoulou, P.; Tchakouteu, S.S.; Sarris, D.; Philippoussis, A.; Papanikolaou, S. Lipids from yeasts and fungi: Physiology, production and analytical considerations. J. Appl. Microbiol.; 2018; 124, pp. 336-367. [DOI: https://dx.doi.org/10.1111/jam.13633]
40. Yang, J.; Cánovas-Márquez, J.T.; Li, P.; Li, S.; Niu, J.; Wang, X.; Nazir, Y.; López-García, S.; Garre, V.; Song, Y. Deletion of Plasma Membrane Malate Transporters Increased Lipid Accumulation in the Oleaginous Fungus Mucor circinelloides WJ11. J. Agric. Food Chem.; 2021; 69, pp. 9632-9641. [DOI: https://dx.doi.org/10.1021/acs.jafc.1c03307]
41. Wang, L.; Zhang, H.; Zhang, Y.; Song, Y. 13C metabolic flux analysis on roles of malate transporter in lipid accumulation of Mucor circinelloides. Microb. Cell Fact.; 2019; 18, 154. [DOI: https://dx.doi.org/10.1186/s12934-019-1207-9]
42. Yang, J.; Li, S.; Khan, A.K.; Garre, V.; Vongsangnak, W.; Song, Y. Increased Lipid Accumulation in Mucor circinelloides by Overexpression of Mitochondrial Citrate Transporter Genes. Ind. Eng. Chem. Res.; 2019; 58, pp. 2125-2134. [DOI: https://dx.doi.org/10.1021/acs.iecr.8b05564]
43. Yang, W.; Dong, S.; Yang, J.; Mohamed, H.; Shah, A.M.; Nazir, Y.; Gao, X.; Fan, H.; Song, Y. Molecular Mechanism of Citrate Efflux by the Mitochondrial Citrate Transporter CT in Filamentous Fungus Mucor circinelloides WJ11. Front. Microbiol.; 2021; 12, 673881. [DOI: https://dx.doi.org/10.3389/fmicb.2021.673881]
44. Nosheen, S.; Yang, J.; Naz, T.; Nazir, Y.; Ahmad, M.I.; Fazili, A.B.A.; Li, S.; Mustafa, K.; Song, Y. Annotation of AMP-activated protein kinase genes and its comparative transcriptional analysis between high and low lipid producing strains of Mucor circinelloides. Biotechnol. Lett.; 2020; 43, pp. 193-202. [DOI: https://dx.doi.org/10.1007/s10529-020-02990-2] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32809159]
45. Nosheen, S.; Naz, T.; Yang, J.; Hussain, S.A.; Fazili, A.B.A.; Nazir, Y.; Li, S.; Mohamed, H.; Yang, W.; Mustafa, K. et al. Role of Snf-β in lipid accumulation in the high lipid-producing fungus Mucor circinelloides WJ11. Microb. Cell Fact.; 2021; 20, 52. [DOI: https://dx.doi.org/10.1186/s12934-021-01545-y] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33639948]
46. Tang, X.; Chen, H.; Gu, Z.; Zhang, H.; Chen, Y.Q.; Song, Y.; Chen, W. Role of g6pdh and leuB on Lipid Accumulation in Mucor circinelloides. J. Agric. Food Chem.; 2020; 68, pp. 4245-4251. [DOI: https://dx.doi.org/10.1021/acs.jafc.9b08155] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32181644]
47. Chu, L.; Zan, X.; Tang, X.; Zhao, L.; Chen, H.; Chen, Y.Q.; Chen, W.; Song, Y. The role of a xylose isomerase pathway in the conversion of xylose to lipid in Mucor circinelloides. RSC Adv.; 2016; 6, pp. 77944-77952. [DOI: https://dx.doi.org/10.1039/C6RA12379A]
48. Zhang, L.; Zhang, H.; Song, Y. Identification and Characterization of Diacylglycerol Acyltransferase from Oleaginous Fungus Mucor circinelloides. J. Agric. Food Chem.; 2018; 66, pp. 674-681. [DOI: https://dx.doi.org/10.1021/acs.jafc.7b04295]
49. Zan, X.; Tang, X.; Chu, L.; Zhao, L.; Chen, H.; Chen, Y.Q.; Chen, W.; Song, Y. Lipase genes in Mucor circinelloides: Identification, sub-cellular location, phylogenetic analysis and expression profiling during growth and lipid accumulation. J. Ind. Microbiol. Biotechnol.; 2016; 43, pp. 1467-1480. [DOI: https://dx.doi.org/10.1007/s10295-016-1820-0]
50. Zan, X.; Tang, X.; Chu, L.; Song, Y. Dual Functions of Lip6 and Its Regulation of Lipid Metabolism in the Oleaginous Fungus Mucor circinelloides. J. Agric. Food Chem.; 2018; 66, pp. 2796-2804. [DOI: https://dx.doi.org/10.1021/acs.jafc.7b06024]
51. Zan, X.; Cui, F.; Sun, J.; Zhou, S.; Song, Y. Novel Dual-Functional Enzyme Lip10 Catalyzes Lipase and Acyltransferase Activities in the Oleaginous Fungus Mucor circinelloides. J. Agric. Food Chem.; 2019; 67, pp. 13176-13184. [DOI: https://dx.doi.org/10.1021/acs.jafc.9b05617]
52. Grubišić, M.; Mihajlovski, K.; Gruičić, A.M.; Beluhan, S.; Šantek, B.; Ivančić Šantek, M. Strategies for Improvement of Lipid Production by Yeast Trichosporon oleaginosus from Lignocellulosic Biomass. J. Fungi; 2021; 7, 934. [DOI: https://dx.doi.org/10.3390/jof7110934]
53. Hao, G.; Chen, H.; Wang, L.; Gu, Z.; Song, Y.; Zhang, H.; Chen, W.; Chen, Y.Q. Role of Malic Enzyme during Fatty Acid Synthesis in the Oleaginous Fungus Mortierella alpina. Appl. Environ. Microbiol.; 2014; 80, pp. 2672-2678. [DOI: https://dx.doi.org/10.1128/AEM.00140-14]
54. Li, Z.; Sun, H.; Mo, X.; Li, X.; Xu, B.; Tian, P. Overexpression of malic enzyme (ME) of Mucor circinelloides improved lipid accumulation in engineered Rhodotorula glutinis. Appl. Microbiol. Biotechnol.; 2013; 97, pp. 4927-4936. [DOI: https://dx.doi.org/10.1007/s00253-012-4571-5]
55. Tang, X.; Chen, H.; Chen, Y.Q.; Chen, W.; Garre, V.; Song, Y.; Ratledge, C. Comparison of Biochemical Activities between High and Low Lipid-Producing Strains of Mucor circinelloides: An Explanation for the High Oleaginicity of Strain WJ11. PLoS ONE; 2015; 10, e0128396. [DOI: https://dx.doi.org/10.1371/journal.pone.0128396] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26046932]
56. Hernández, M.A.; Alvarez, H.M. Increasing lipid production using an NADP+-dependent malic enzyme from Rhodococcus jostii. Microbiology; 2019; 165, pp. 4-14. [DOI: https://dx.doi.org/10.1099/mic.0.000736]
57. Zhu, B.-H.; Zhang, R.-H.; Lv, N.-N.; Yang, G.-P.; Wang, Y.-S.; Pan, K.-H. The Role of Malic Enzyme on Promoting Total Lipid and Fatty Acid Production in Phaeodactylum tricornutum. Front. Plant Sci.; 2018; 9, 826. [DOI: https://dx.doi.org/10.3389/fpls.2018.00826] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29971080]
58. Xue, J.; Wang, L.; Zhang, L.; Balamurugan, S.; Li, D.-W.; Zeng, H.; Yang, W.-D.; Liu, J.-S.; Li, H.-Y. The pivotal role of malic enzyme in enhancing oil accumulation in green microalga Chlorella pyrenoidosa. Microb. Cell Fact.; 2016; 15, 120. [DOI: https://dx.doi.org/10.1186/s12934-016-0519-2] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27387324]
59. Xue, J.; Niu, Y.-F.; Huang, T.; Yang, W.-D.; Liu, J.-S.; Li, H.-Y. Genetic improvement of the microalga Phaeodactylum tricornutum for boosting neutral lipid accumulation. Metab. Eng.; 2015; 27, pp. 1-9. [DOI: https://dx.doi.org/10.1016/j.ymben.2014.10.002] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25447640]
60. Yan, J.; Kuang, Y.; Gui, X.; Han, X.; Yan, Y. Engineering a malic enzyme to enhance lipid accumulation in Chlorella protothecoides and direct production of biodiesel from the microalgal biomass. Biomass-Bioenergy; 2019; 122, pp. 298-304. [DOI: https://dx.doi.org/10.1016/j.biombioe.2019.01.046]
61. Sun, X.; Han, G.; Meng, Z.; Lin, L.; Sui, N. Roles of malic enzymes in plant development and stress responses. Plant Signal. Behav.; 2019; 14, e1644596. [DOI: https://dx.doi.org/10.1080/15592324.2019.1644596]
62. Brutnell, T.P.; Wang, L.; Swartwood, K.; Goldshmidt, A.; Jackson, D.; Zhu, X.-G.; Kellogg, E.; Van Eck, J. Setaria viridis: A Model for C4 Photosynthesis. Plant Cell; 2010; 22, pp. 2537-2544. [DOI: https://dx.doi.org/10.1105/tpc.110.075309]
63. Zhou, H.; Liu, S.K.; Yang, C.P. Over-Expression of a NAD-Malic Enzyme Gene from Rice in Arabidopsis thaliana Confers Tolerances to Several Abiotic Stresses. Adv. Mater. Res.; 2011; 393–395, pp. 863-866. [DOI: https://dx.doi.org/10.4028/www.scientific.net/AMR.393-395.863]
64. Drincovich, M.F.; Casati, P.; Andreo, C.S. NADP-malic enzyme from plants: A ubiquitous enzyme involved in different metabolic pathways. FEBS Lett.; 2001; 490, pp. 1-6. [DOI: https://dx.doi.org/10.1016/S0014-5793(00)02331-0]
65. Kang, F.; Rawsthorne, S. Starch and fatty acid synthesis in plastids from developing embryos of oilseed rape (Brassica napus L.). Plant J.; 1994; 6, pp. 795-805. [DOI: https://dx.doi.org/10.1046/j.1365-313X.1994.6060795.x]
© 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/). Notwithstanding the ProQuest Terms and Conditions, you may use this content in accordance with the terms of the License.
Details
; Aabid Manzoor Shah 1 ; Naz, Tahira 1
; Nosheen, Shaista 1
; Wu, Yang 1 ; Garre, Victoriano 2
; Younis Majeed 3 ; Mohammed Khalid Al-Sadoon 4 ; Song, Yuanda 1
1 Colin Ratledge Center for Microbial Lipids, School of Agricultural Engineering and Food Science, Shandong University of Technology, Zibo 255000, China;
2 Departamento de Genética y Microbiología, Facultad de Biología, Universidad de Murcia, 30100 Murcia, Spain;
3 Department of Biotechnology, University of Kashmir, Srinagar 190006, Jammu and Kashmir, India;
4 Department of Zoology, College of Science, King Saud University, P.O. Box 2455, Riyadh 11451, Saudi Arabia;