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Department of Biological Sciences, University of Hull, Hull HU6 7RX, UK
Correspondence
Colin Ratledge
c.ratledge{at}hull.ac.uk
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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-linolenic acid (18 : 3 n-6). At the end of lipid accumulation (96 h), ME activity in the recombinant strains had ceased, as it had done in the parent wild-type cells, indicating that additional, but unknown, controls over its activity must be in place to account for this loss of activity: this may be due to the presence of a specific ME-cleaving enzyme. The hypothesis that the rate-limiting step of fatty acid biosynthesis is therefore the supply of NADPH, as generated specifically and solely by ME, is therefore considerably strengthened by these results.
-linolenic acid; ME, malic enzyme
Present address: National Institutes of Health, NIDCR, 9000 Rockville Pike, Bethesda, MD 20892, USA.
Present address: Centre for Novel Agricultural Products (CNAP), Department of Biology, University of York, York YO10 5DD, UK.
The GenBank/EMBL/DDBJ accession numbers for the ME sequences of Mc. circinelloides and Mt. alpina are DQ975377 and DQ973624, respectively.
A table showing the strains and plasmids used in this study is available with the online version of this paper.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). The latter two products are currently used in infant formula preparations and the demand for them is now exceeding the present production capacity (Ratledge, 2005; Ratledge & Hopkins, 2006a, b). Although the process of fatty acid biosynthesis is now well known, the genetic factors that control the extent to which lipid accumulates in a particular organism are far from clear (almost invariably the lipid that is accumulated by animals, plants and micro-organisms is in the form of triacylglycerols, the chief components of which are the fatty acyl chains).
In both plants and micro-organisms, the extent of lipid accumulation is determined by the genetic constitution, as maximum attainable lipid contents can vary enormously between species and even between individual strains. In micro-organisms, the range of lipid contents amongst the so-called oleaginous species can vary from 20–25 % to >70 % (Ratledge, 1997). The oil content of commercial plant seeds can range from 
15 %, as with soybeans, to 50 % with groundnuts (peanuts) and other oil-seed crops. Various attempts have been made with plants to increase the flux of carbon into fatty acid biosynthesis and to identify possible bottlenecks, with subsequent genetic modification being carried out to remove postulated impediments to lipid accumulation. None of these attempts has been successful, with only relatively small increases (10–15 %) in lipid accumulation being recorded (see for example Broun et al., 1999; Thelen & Ohlrogge, 2002; Rangasamy & Ratledge, 2000; Ramli et al., 2005) following genetic manipulation.
For some time it has seemed to us that the overall regulation of fatty acid biosynthesis must lie outside the flux of carbon in view of this lack of success in overexpressing various supposed rate-limiting enzymes of fatty acid synthesis (Wynn & Ratledge, 1997; Wynn et al., 1999; Ratledge & Wynn, 2002; Ratledge, 2004). We have previously described the process of lipid accumulation in both oleaginous yeasts and filamentous moulds as requiring the key activity of ATP : citrate lyase (ACL) in order for cells to produce a sufficient supply of acetyl-CoA in the cytosol (Botham & Ratledge, 1979; Evans & Ratledge, 1985). Although this activity is essential for lipid accumulation to take place (i.e. lipid to exceed more than 20 % of the cells), it is insufficient to explain the range of lipid levels in the oleaginous cells, as there is no correlation between ACL activity and lipid accumulation (Ratledge & Wynn, 2002). This cytosolic enzyme is, however, absent in non-oleaginous yeasts such as Saccharomyces cerevisiae and Candida utilis (Botham & Ratledge, 1979). Overexpression of the gene encoding ACL in tobacco plants fails to increase lipid content by more than 15 % of that of the control plants (Rangasamy & Ratledge, 2000). (It should be noted that at that time it was not possible to carry out similar genetic modifications of oleaginous micro-organisms, as none of these species had been established as genetically manipulatable).
Of key significance, at least in our view, has been the role of malic enzyme (ME; EC 1 . 1 . 1 . 40) in lipid accumulation. This enzyme carries out the irreversible decarboxylation of malate to pyruvate with the formation of NADPH from NADP+. It is then this NADPH which is vital for fatty acid biosynthesis. No other NADPH-generating enzyme appears able to provide the requisite reducing power for fatty acid synthase to function (Wynn & Ratledge, 1997; Wynn et al., 1999). We have shown using Mucor circinelloides as a model organism that sesamol, as a specific inhibitor of ME activity, decreases lipid accumulation from 25 % of the cell biomass to 2 % without adversely affecting growth (Wynn et al., 1997). Moreover, there is a direct correlation between the decreasing activity of ME during the lipid-accumulation phase and the extent of lipid accumulation in both this fungus and the related Mortierella alpina (Wynn et al., 1999): when ME activity ceases, lipid accumulation stops. Thus, an explanation of why different organisms have different levels of their maximum lipid contents can be provided by the expression and possible control of ME.
Further, a mutant of the fungus Aspergillus nidulans that lacks ME activity has only 12 % lipid when grown under exactly the same conditions as the wild-type which has ME activity and accumulates 30 % of its biomass as lipid (Wynn & Ratledge, 1997). Although we have also shown that whilst there are several isoforms of ME present in Mc. circinelloides (Song et al., 2001), only one of these (termed isoform III) is involved in NADPH formation and is linked to the lipid-accumulation process. This isoform III is converted into isoform IV, presumably post-transciptionally, upon the commencement of lipid accumulation, which is triggered by depletion of nitrogen from the growth medium. We have isolated and sequenced another isoform (type II) of ME from Mc. circinelloides (Li et al., 2005), but this is the form of ME associated with anaerobic metabolism and not with fatty acid biosynthesis.
In this present paper, we have now identified the gene encoding ME isoforms III/IV and, having isolated and sequenced it, have reintroduced it into Mc. circinelloides under the control of a constitutive promoter. In this way we have been able to achieve expression of a higher activity of ME in the fungus, with the result that the content of lipid in the cells has increased from 12 % of the biomass to 30 %. However, ME activity still disappears by the end of the lipid-accumulation phase, with the result that lipid accumulation, though more than doubled, does not continue indefinitely. A possible reason for this is discussed later in the paper.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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), was used as the recipient in transformation experiments and as a control strain for gene expression with 20 µg leucine ml–1 in its medium. Escherichia coli strain InV
F' (Invitrogen) was used for DNA manipulation. E. coli strain LE392 (Promega) was used for genomic library construction. A list of the strains and plasmids used in this work is given in Supplementary Table S1.
Mt. alpina and Mc. circinelloides were maintained on solidified Kendrick medium (Kendrick & Ratledge, 1992) at 30 °C and stored at 4 °C. Mc. circinelloides R7B was grown at 30 °C on YPG complete medium (3 g yeast extract, 10 g peptone and 20–50 g glucose per litre distilled water) or, for plate cultures, in the minimal medium YNB (0.5 g Difco yeast nitrogen base without amino acids, but with 1.5 g ammonium sulphate l–1, 1.5 g glutamic acid l–1 and 10 g glucose l–1) with the glucose added post-sterilization. Fungi (about 0.3x105 spores of each strain) were initially cultivated in 1 l stirred bottles containing 800 ml Kendrick medium for 16 h at 30 °C, and then inoculated at 10 % (v/v) into a 5 l fermenter containing 4 l Kendrick medium modified to contain 2 g ammonium tartrate l–1 and 50 g glucose l–1. Leucine at 20 µg ml–1 was included in the medium used for Mc. circinelloides R7B as control. Fermenters were held at 30 °C, stirred at 500 r.p.m. with aeration at 0.5 volume of air per volume of fermenter per minute (v/v min–1), and pH was maintained at 5.5–6.5 by automatic addition of 2 M KOH or 2 M HCl.
Nucleic acid manipulation.
High-molecular-mass genomic DNA was extracted from mycelia of Mt. alpina CBS 696.70 and Mc. circinelloides CBS 108.16, and grown for 24 h in Kendrick medium which had been freeze-ground with liquid N2. A standard phenol/chloroform extraction procedure (Michaelson et al., 1998) was used prior to application of a DNA Clean-up kit (Promega), and the final DNA was dissolved in Tris/EDTA buffer (10 mM Tris/HCl, 1 mM EDTA, pH 7.0). Total RNA was isolated by using the RNeasy Plant Mini kit (Qiagen) with freeze-ground mycelium following the manufacturer's instructions. Southern and Northern blotting were performed using standard procedures for capillary transfer of nucleic acids to nylon membranes. DNA was labelled with [-32P]dCTP or a fluorescent labelling kit (Amersham). In all experiments, hybridization was carried out at 55 °C in a hybridization buffer, and the blots were subsequently washed at 55 °C successively with 2x saline sodium citrate (SSC), 1x SSC and 0.5x SSC containing 0.5 % SDS (SSC=0.15 M NaCl, 15 mM sodium citrate, pH 7.0). Signals were detected by autoradiography on X-ray film. Northern blot signals were corrected for fluctuations in RNA loading by comparison with band densities obtained from concurrent electrophoresis in an ethidium bromide-stained formaldehyde-containing agarose gel.
Amplification of fragments of the ME gene from Mt. alpina and Mc. circinelloides.
Degenerate primers were designed according to the homology with conserved amino acid sequences of ME and were obtained commercially (MWG-Biotech) as sense primers P1 [5'-GT(AGCT)GT(AGCT)AC(AGCT)GA(CT)GG(AGCT)CA(AG)-3'] and P2 [5'-GG(AGCT)AT(ACT)CC(AGCT)GTTGG(AGCT)AAA-3'], and antisense primers P3 [5'-(AG)TT(AGCT)GC(AG)AA(AG)TC(CT)TC(AG)AA(CT)TG-3'] and P4 [5'-(AGCT)CC(CT)TG(AGT)AT(AG)TC(AG)TC(AG)TT(AG)AA-3']. Two DNA fragments, 310 and 370 bp, were amplified using Mt. alpina and Mc. circinelloides genomic DNA, respectively, by primers P2 and P4. PCR conditions were 94 °C for 5 min, 30 cycles at 94 °C for 1 min, 44 °C for 1 min and 72 °C for 1 min, and a final extension at 72 °C for 10 min. PCR products were cloned into the pGEM-T Easy vector (Promega) and the resulting plasmids (named pFMt6 and pFMc4.1 for Mt. alpina and Mc. circinelloides, respectively) were purified using a Wizard column (Promega). A list of all plasmids used in this work is given in Supplementary Table S1.
Genomic library construction and screening.
Two genomic libraries were constructed using LambdaGME-11 Genomic Cloning Vector (BamHI arms) (Promega) with Sau3AI-digested genomic DNA prepared from Mc. circinelloides CBS 108.16 and Mt. alpina CBS 696.70, following the manufacturer's instructions. The libraries were screened with Amersham Redivue [-32P]dCTP-labelled PCR-amplified probe from EcoRI-digested pFMt6 and pFMc4.1. Approximately 2x105 p.f.u. was used in each primary screen. Eleven positive plaques were obtained in the secondary screen from the Mt. alpina genomic library and 13 from that of Mc. circinelloides.
Transformation conditions.
Transformation of E. coli strain InVF' (Invitrogen) was used in this study. Competent cells for transformation were obtained according to the manufacturer's instructions and transformed by heat shock at 42 °C for 90 s. The transformants were selected on LB plates with 50 µg ampicillin ml–1 and 160 µg X-Gal ml–1. Plasmids were isolated by growing the cells in liquid LB medium and using a Wizard column (Promega).
Transformation of Mc. circinelloides R7B was carried out as described elsewhere (van Heeswijck & Roncero, 1984) with the modifications of Wolff et al. (2002), as follows. Protoplasts were prepared by enzymic treatment of germlings with a mixture of 125 µg chitosanase-RD (US Biologicals) and 5 U chitinase (from Streptomyces griseus, Sigma) in a total volume of 2 ml. Cell wall digestion was carried out for 2–3 h at 28 °C. Typically, 2–5 µg DNA was used per transformation. Transformants were selected on YNB plates maintained at room temperature for 2 days.
DNA sequencing and bioinformatic analysis.
The DNA sequences from both strands were determined with the Sequencing kit (Beckman Coulter) and a capillary electrophoresis sequencer (CEQ 8000, Beckman Coulter) according to the manufacturer's instructions. The sequence data were interpreted and aligned by CodonCode Aligner. The position of introns was speculated by bioinformatic analysis using MacVector software (Oxford Molecular), and the sequences were confirmed by cDNA sequences.
Construction of expression vectors.
The episomal E. coli–Mc. circinelloides shuttle vector pEUKA11, containing the E. coli kanR gene under the transcriptional control of the Mc. circinelloides glyceraldehyde-3-phosphate dehydrogenase gene (gpd1) promoter and terminator, was a kind gift from Dr Jose Arnau at the Bioteknologisk Institut, Hoersholm, Denmark (see Appel et al., 2004; Wolff et al., 2002).
Each full-length malEMt and malEMc gene (from Mt. alpina and Mc. circinelloides) was cloned separately into the expression vector pEUKA11 to replace the kanR gene under the control of the gpd1 promoter and terminator. Overexpression of malEMt and malEMc was achieved by a high level of constitutive gene expression in Mc. circinelloides R7B under the control of the gpd1 promoter and terminator (see Fig. 1).
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The 3 kb Mt. alpina and 2.1 kb Mc. circinelloides ME gene fragments were created and inserted into the pGEM-T Easy vector (Promega). The resulting plasmids (pMtME and pMcME) were digested with XhoI and NotI, and the fragments containing the full-length malEMt and malEMc genes were then inserted into the linear pEUKA11 vector digested by XhoI and NotI and gel-purified to remove the kan gene. In the resulting plasmids (pEUKMt2.6 and pEUKMc4.8), the individual ME gene was placed under the control of the promoter and terminator regions of the gpd1 gene; the nucleotide sequences of the fusion plasmids were confirmed by sequencing.
Isolation of single Mc. circinelloides transformants with pEUKMt2.6 and pEUKMc4.8.
Transformed primary colonies were isolated on YNB plates (pH 3) to select for leucine-independent growth. Well-isolated colonies were transferred from the original transformation plates to fresh YNB plates and, after appropriate growth of the successive vegetative cycles, sporangiospores were harvested.
Determination of ME activity.
Biomass was harvested by filtration under reduced pressure through a Whatman no. 1 filter, washed with distilled water and then suspended in extraction buffer [100 mM KH2PO4/KOH, pH 7·5, containing 20 % (w/v) glycerol, 1 mM benzamidine and 1 mM DTT]. After being disrupted by a single pass through a One-Shot cell disrupter (Constant Systems) at 64 MPa, the material was centrifuged (10 000 g for 10 min at 4 °C) and the supernatant used immediately for the determination of ME activity (see Wynn et al., 1997). Protein was determined using the Bradford method with BSA as a standard.
Determination of cell dry weight, and glucose and ammonium ion concentrations.
Biomass was harvested by filtration through a pre-weighed glass-fibre filter (Whatman GF/A), washed twice with distilled water and dried at 110 °C to a constant weight. Glucose in the culture medium was determined using a glucose oxidase Perid test kit (Boehringer Mannheim) and ammonium using the indophenol method (see Song et al., 2001).
Analysis of cell lipid and fatty acids.
Lipid was extracted from freeze-dried biomass as previously described (Song et al., 2001) and its amount determined gravimetrically. Fatty acids, as their methyl esters, were prepared and analysed by GC, as described by Wynn & Ratledge (2000).
Chemicals.
Unless otherwise stated, all chemicals were purchased from Sigma.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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These two amplified DNA fragments were separately used as probes to isolate corresponding lambda clones from Mt. alpina and Mc. circinelloides CBS 108.16 genomic libraries. Screening of the genomic libraries resulted in the isolation of 13 positive clones for Mt. alpina from 6x104 plaques and 11 positive clones for Mc. circinelloides from 2x105 plaques. One of the clones from the Mt. alpina genomic library had a 22 kb DNA fragment containing the Mt. alpina ME gene and one from the Mc. circinelloides genomic library had a 23 kb DNA fragment containing the Mc. circinelloides ME gene. A 4.5 kb DNA fragment from the former and a 5.5 kb DNA fragment from the latter were obtained from each positive lambda clone after digestion by XhoI or PstI, respectively, and were inserted into XhoI- or PstI-digested plasmid pBluescript SKII (pBSK) (Stratagene). The resulting plasmids, pMtX1.4 and pMcP3.7, were purified, each inserted sequence was then determined, and genes were named malEMt and malEMc, respectively. The ATG codons in the Mt. alpina DNA fragment at nt 1006 and in the Mc. circinelloides DNA fragment at nt 2373 were assumed to be the translation initiation codons for each gene. This conclusion was based on their positions relative to the promoter elements, notably CCAAT-, TATA- and CT-rich regions, and the consensus sequence generated by using these frames. Therefore, the consensus sequences of the putative ME genes from Mt. alpina and Mc. circinelloides were considered to be, respectively, 2983 and 2104 bp.
Two introns (436 and 537 bp) in malEMt and four introns (69, 57, 56 and 59 bp) in malEMc were confirmed by cDNA sequencing using RT-PCR products as templates. The putative promoter regions (CCAAT-, TATA- and CT-rich regions) existed within 200 bp upstream of the translation initiation site (ATG) of both malEMt and malEMc.
Genomic Southern blot analysis
Southern blots of Mt. alpina CBS 696.70 genomic DNA, digested with XhoI, EcoRI and BamHI, and Mc. circinelloides CBS 108.16 genomic DNA, digested with PstI, EcoRI and BamHI, and hybridized with the respective full-length gene, were performed. Only one strong signal was obtained from each hybridization after film development, indicating that both fungal strains have a single gene copy (data not shown).
Analysis of malEMt and malEMc sequence
GenBank database searches with the deduced amino acid sequences of MEs from Mt. alpina and Mc. circinelloides revealed close matches to the complete amino acid sequences of MEs from a number of yeast and fungi. In an alignment, the putative amino acid sequences of the Mt. alpina ME showed, respectively, 49, 50, 47, 45, and 45 % identity to those from Neurospora crassa (Galagan et al., 2003), A. nidulans (Galagan et al., 2005), Gibberella zeae PH-1 (B. Birren and others, unpublished work; see http://www.broad.mit.edu/annotation/genome/fusarium_graminearum/Home.html), Magnaporthe grisea (Dean et al., 2005) and Dictyostelium discoideum (Eichinger et al., 2005). In a similar alignment, the putative amino acid sequence of the Mc. circinelloides MEs showed, respectively, 42, 44, 43, 42 and 45 % identity to the same five organisms.
The alignment showed that there was 55 % identity between the amino acid sequences of Mt. alpina and Mc. circinelloides, and that the identical residues were scattered throughout the sequence.
From these alignments, regions of functional significance could be found (Fig. 2). Consensus sequences in the primary structures of the two MEs revealed that two highly conserved dinucleotide binding sites, GXGXXG/A (highlighted in Fig. 2), are present regardless of the cofactor specificities of the enzyme (Wierenga et al., 1986; Rothermel & Nelson, 1989; Borsch & Westhoff, 1990). Regions Y(138) to C(146) in Mc. circinelloides and Y(194) to C(202) in Mt. alpina (highlighted in Fig. 2) matched perfectly with the binding site for malate [numbered Y(44)–C(52)] given by Kulkarni et al. (1993). The sequences from D(199) to D(231) in Mc. circinelloides and from D(256) to D(288) in Mt. alpina (highlighted in Fig. 2) could be assigned to the consensus sequence for the site at which the ADP ring of NADP+ is bound, which has a predicted ββ secondary structure [numbered D(193)–D(223) in Wierenga et al., 1986]. Furthermore, the crystal structures of the NAD(P)+-dependent ME from human mitochondria (Yang et al., 2000) and an NADP+-dependent ME from pigeon cytosol (Yang et al., 2002) (neither shown in Fig. 2) have been reported; various residues, F, E, D (numbered as 347, 348 and 349, respectively, in the Mt. alpina sequence) and F, N, D and D (369–372 in the same sequence and highlighted in Fig. 2) are also highly conserved and are probably involved in the binding of a divalent metal ion. The K(220) residue in Mc. circinelloides and the K(277) residue in Mt. alpina (numbered Lys183 in the pigeon ME by Yang et al., 2002) may be important residues for catalytic activity. In addition, the 2'-phosphate group of NADP+ would be predicted to interact with S(387) in Mc. circinelloides and S(440) in Mt. alpina (numbered Ser346 by Yang et al., 2002). Many of these residues are highly conserved in MEs from a wide range of species. For example, E(291), D(292) and D(315) in Mc. circinelloides correspond to K(456) in Mt. alpina (numbered Lys362 in the pigeon enzyme in Yang et al., 2002), and these residues are always highly conserved in NADP+-dependent MEs (Yang et al., 2002; Chang & Tong, 2003).
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Characterization of the transformed strains
(a) Identification of transformed Mucor strains by Southern blotting.
To investigate the role that ME plays in lipid accumulation and fatty acid desaturation in Mt. alpina and Mc. circinelloides, the respective ME gene overexpression vectors, pEUKMt2.6 and pEUKMc4.8, were transformed into Mc. circinelloides R7B, which is a leucine auxotroph (see Methods). Seventeen transformants with pEUKMt2.6, named Mucor-malEMt, and nine transformants with pEUKMc4.8, named Mucor-malEMc, were obtained and analysed for being stable Leu+ phenotypes by successive cycles of vegetative growth in the absence of selective pressure. One transformant of each type was finally selected for Southern analysis using a LeuA gene fragment (PstI-digested from plasmid pLEU4, kindly provided by Dr Santiago R. Torres Martines, University of Murcia) as a probe. The hybridization with genomic DNA from both mutant strains showed strong signals, while no signal was detected from genomic DNA of the wild-type strain (see Fig. 3). This result demonstrated that both overexpression vectors had been separately transformed into Mc. circinelloides R7B and that they carried both the leucine gene and the malEMt or malEMc gene.
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(b) Northern blot analysis.
To examine the transcript levels of malE genes, total RNA was isolated from all three strains grown in the 5 l fermenters. Northern blotting was performed on cells taken after 24, 48 and 72 h using the ME gene-specific probes from pMt6 or pMc4.1 (see Fig. 4). The transcript level of the wild-type strain decreased rapidly after nitrogen exhaustion from the medium (before 24 h; see Fig. 5), but in the two transformants it remained high throughout growth and the subsequent lipid-accumulation phase. Thus, in the wild-type strain, there is a rapid cessation of ME expression, thereby leading to loss of ME activity and consequent low lipid accumulation (see below). In the transformants, continued expression of ME was achieved by placing the gene under the control of the gpd1 promoter.
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. Activities of ME within cells at different stages of culture were consistently higher in the two transformants than in the original parent. Up to threefold higher activity was seen in the McmalEMt transformant and up to twofold in the McmalEMc transformant compared with the control (R7B) strain.
The overexpressed MEs were also detected by activity staining after non-denaturing PAGE (see Fig. 6). Unlike Mc. circinelloides CBS 108.16, which produces six ME isoforms (Song et al., 2001), only two isoforms were observed in the Mc. circinelloides R7B strain, and these corresponded to isoforms III and IV of the former strain. These are the two isoforms associated with lipid accumulation (see Introduction). The activity of the smaller isoform in each mutant strain (see Fig. 6) was stronger and remained longer than that in the control strain, and was thus instrumental in the increased accumulation of lipid in the transformants (see below).
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from the medium, was similar in the two transformants and in the control strain. 
was depleted in all three cases before 24 h, and lipid accumulation probably commenced 2–4 h prior to this. Glucose consumption rates of the two transformants were faster than that of control strain after 48 h, and concomitantly the cell dry weights also increased more rapidly in the control strain (see Fig. 5). Significantly, both transformed strains accumulated considerably more lipid than the original wild-type strain (see Fig. 5), and this explains the increased uptake of glucose and the higher biomass levels. The amounts of lipid accumulated in the transgenic strains malEMt and malEMc were, respectively, 2.5- and 2.4-fold more than in the control strain.
Not only was a much higher amount of lipid synthesized by the two transformants than by the original strain but also, again significantly, the higher activity of ME in the transformants maintained and even increased the amounts of unsaturated fatty acids produced in the lipid (see Table 1). Thus, increased ME activity led to increases of both lipid biosynthesis and unsaturated fatty acid production, including that of -linoleic acid (18 : 3 n-6), which is a feature of this mould.
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DISCUSSION |
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). To isolate the ME genes from Mt. alpina and Mc. circinelloides, degenerate primers were designed based on the amino acid sequence motifs conserved in multiple species, and the subequent PCR product was used as probe to isolate MEs from each of the respective genomic libraries. From both cloned ME genes, malEMt and malEMc, CCAAT-, TATA- and CT-rich boxes, which are generally present in the promoter regions of eukaryotes, were identified within 200 bp upstream of the transcriptional initiation site (ATG). Two introns of 436 and 537 bp are present in malEMt from Mt. alpina, for which relatively large introns (up to 1 kb) have been reported (Wongwathanarat et al., 1999; Mackenzie et al., 2000). Four introns of 69, 57, 56 and 59 bp are present in malEMc from Mc. circinelloides, and for this species the mean size of an intron is 69 bp (Velayos et al., 2000). The sequence of malEMt has a G+C content of 54 %, whereas the introns have a G+C content of 49 %. The sequence of malEMc has a G+C content of 56 %, with the introns having G+C contents of 36, 42, 41 and 39 %. These data fall within the general rule that a sequence in an intron contains more A+T than G+C.
The results of the Northern blot analysis suggest a similar manner of transcription of both ME genes. Therefore, the temporal difference in the persistence of ME activity, previously reported in the wild-type strains of Mt. alpina and Mc. circinelloides after nitrogen exhaustion (see Wynn et al., 1999), is probably due to the different cellular half-lives of the respective ME proteins.
The downstream flanking regions of malEMt and malEMc were examined. No mitochondrial leader sequence was found from the deduced amino acids of either gene. The results of a BLAST search with GenBank showed high homology to other cytosolic ME genes (data not shown, but see Results for references). Therefore, these two MEs are presumptive cytosolic enzymes.
A single gene copy number of malEMt and malEMc in Mt. alpina and Mc. circinelloides, respectively, suggests that at least some of the multiple ME isoforms noted in both these fungi (Song et al., 2001; Zhang, 2005) are encoded by different genes. Thus, the sequence of malEMc in this study is completely different from that of isoform II (mce1) from the same Mc. circinelloides strain that encodes a mitochondrial ME (Li et al., 2005; and see Fig. 2). With isoforms III and IV (see Introduction), it is considered that both arise from the gene that has been cloned in this work; isoform IV, which appears later than isoform III (see Fig. 6), is probably formed by post-translational modification.
The process of lipid accumulation in oleaginous micro-organisms, whereby the lipid content may reach 70 %, and possibly higher, is still imperfectly understood. Although it has long been established that the build-up of lipid storage reserves, mainly in the form of triacylglycerols, requires the presence of ACL to generate acetyl-CoA in the cytosol (Botham & Ratledge, 1979), the activity of this enzyme does not correlate with the amount of lipid that a particular micro-organism may accumulate (Boulton & Ratledge, 1981; Ratledge & Gilbert, 1985; Wynn et al., 1998). Indeed, a feature of lipid storage in micro-organisms, and also in plants, is that the maximum amount of lipid that can be stored in an individual species appears to be under genetic control in that there seems to be an absolute ceiling beyond which further increases in lipid content cannot proceed. The exceptions to this are some oleaginous yeasts, which accumulate lipid up to 70 % and beyond, and in which the maximum content is probably limited by the physical capacity of the cell to accommodate large oil droplets, bearing in mind that such cells still contain other organelles. In such organisms, all enzyme activities that participate in glycolysis and lipid biosynthesis, including ME, remain active throughout the lipid-accumulation phase (Evans & Ratledge, 1983).
Whilst various proposals have been made to determine the rate-limiting step in both micro-organisms and plants (Ratledge & Wynn, 2002; Ohlrogge & Jaworski, 1997; Ramli et al., 2005), none of the possible enzymes involved in either acetyl-CoA generation or fatty acid synthesis appears to have been vindicated by subsequent cloning experiments (Ohlrogge & Jaworski, 1997; Rangasamy & Ratledge, 2000). In our detailed biochemical analysis of lipid accumulation in Mc. circinelloides, which has been used commercially to produce an oil rich in -linolenic acid (GLA; 18 : 3 n-6), and also in Mt. alpina, currently used to produce arachidonic acid (20 : 4 n-6), we had identified ME activity as the most likely rate-limiting step in this process (Wynn et al., 1999, 2001; Song et al., 2001). ME, through its oxidative decarboxylation of malate to pyruvate, provides NADPH which is essential for biosynthetic reactions. Although other NADPH-generating enzymes exist [glucose-6-phosphate dehydrogenase, 6-phosphogluconate dehydrogenase (these two enzymes are involved in the initial reactions of the pentose phosphate cycle) and NADP+-dependent isocitrate dehydrogenase], their activities do not correlate with either the onset of lipid accumulation or its cessation. Of all the enzyme activities that have been monitored in both these fungi, only that of ME shows a close correlation with lipid accumulation. When ME activity ceases, so does lipid accumulation, whether it is at the limit of about 25 % cell dry weight for Mc. circinelloides or at 40 % for Mt. alpina (Wynn et al., 1999). Further, when ME activity was inhibited by sesamol, derived from sesame seed oil, lipid accumulation in Mc. circinelloides dropped from 25 to 2 % of the biomass, without any effect on cell growth (Wynn et al., 1997). Thus, although the general view would suggest that there is a common pool of NADPH within a cell, our previous results would refute this, at least in the fungi we have studied, and have strongly suggested a positive linkage of ME activity with fatty acid biosynthesis (Ratledge & Wynn, 2002; Ratledge, 2004).
In this present work, we have now shown that lipid production can be increased substantially by placing the gene encoding ME [either isoforms III/IV from Mc. circinelloides (Song et al., 2001) or its major isoform in Mt. alpina (Zhang, 2005)] under the control of a constitutive promoter. The transformed cells had a lipid content that was 2.5-fold that of the original parent, although it should be noted that the parent strain had a maximum lipid content of only 12 %; this was about half that of the strain we had originally used, but the latter had had no prior genetic work carried out with it and so was inappropriate for genetic manipulation. Simultaneous with the substantial increase in lipid accumulation was a slight but definite overall increase in the GLA content of the lipid, indicating that the increased activity of ME was probably also generating NAPDH for fatty acid desaturases (see Kendrick & Ratledge, 1992). Thus, increased ME activity led not only to an increased lipid content of the cells but also to an increased fatty acid desaturase activity.
It is, however, clear that there are still factors that are limiting the continued activity of ME even though it is now being expressed quasi-constitutively. Although we achieved higher expression of the enzyme, and consequently a higher activity than before, ME activity still declined after the exhaustion of nitrogen from the culture medium (see Fig. 5), in spite of the continued formation of the mRNA for ME (see Fig. 4). It is a prerequisite of lipid accumulation that nitrogen (or some nutrient other than carbon) be exhausted so that cell proliferation is stopped but carbon assimilation still continues. Lipid accumulation is then not so much an increased activity of appropriate lipid-synthesizing enzymes, but arises because other cell activities, including multiplication, have ceased or substantially declined. We have previously shown that during the lipid-accumulation phase in Mc. circinelloides, there is a change in ME from isoform III to isoform IV (Song et al., 2001), which is possibly due to the removal of a short peptide sequence from the protein (Z. Bing & C. Ratledge, unpublished work), but with the overall consequence that ME activity gradually declines and eventually the enzyme ceases to be functional. This was again noted with both the transformants: ME activity still substantially declined. One possible explanation for this is that there could be an enzyme that specifically degrades ME (an ME-converting enzyme?) once the cells are nitrogen limited and have commenced lipid storage.
Lipid accumulation thus can only continue for as long as ME continues to be active. The transformed cells were still limited in their capacity to accumulate lipid because of the continuing inactivation (conversion?) of ME. They were still some way off what would be regarded as a physical limit of lipid production. It may be conjectured that if ME could be engineered to resist degradation (or if the gene for the suggested ME-converting enzyme could be deleted) then fungi such as Mc. circinelloides and Mt. alpina should be able to accumulate as much lipid as the most efficient oleaginous yeasts, in which ME does indeed remain fully active (Evans & Ratledge, 1983) and lipid can accumulate to very high levels (>70 %).
Whether increasing ME activity in other systems, such as plant and animal cells, would also lead to increased lipid accumulation is less certain. In plants, there is a wide range of functions for ME, which is regarded as a ubiquitous enzyme that is involved in a variety of different metabolic pathways, ranging from a role in photosynthesis to other still unknown roles (Drincovich et al., 2001). Thus, identifying the correct ME gene involved in generating NADPH specifically for fatty acid biosynthesis may not be an easy task. In animal cells, although ME has been recognized for some time as being important for generating NADPH for fatty acid biosynthesis (e.g. see Castelein et al., 1994; Hillgartner & Charron, 1998; Sourdioux et al., 1999; Ceddia et al., 2000), it is not the only provider: both glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase also provide 50 % of the NADPH for fatty acid biosynthesis (Rognstad & Katz, 1979; Shimomura et al., 1998). These two enzymes are part of the pentose phosphate pathway, and like ME, remain active during lipid formation. They only appear to be repressed by starvation of the animal (Hillgartner & Charron, 1998). However, in oleaginous micro-organisms, the pentose phosphate pathway is repressed during lipid accumulation (Evans & Ratledge, 1983, 1984), as its operation is not required to provide intermediates for protein and nucleic acid biosynthesis, both of which are stopped by the exhaustion of nitrogen from the culture medium at the start of lipid accumulation. Such conditions, of course, could not apply to animals.
Thus, it is probably only in micro-organisms that ME assumes a unique role in fatty acid biosynthesis that cannot be replaced by another enzyme. This uniqueness would also seem to apply to the formation of unsaturated fatty acids (Kendrick & Ratledge, 1992), which also requires NADPH and, being a membrane-associated activity, clearly needs a mechanism by which the highly water-soluble cofactor can be supplied to the fatty acid desaturases contained within the membranes of the endoplasmic reticulum. ME evidently fulfils this second role in addition to its role in fatty acid biosynthesis, as we have shown here.
Codicil
This paper concludes the work of C. R. on the biochemistry of lipid accumulation in oleaginous micro-organisms. Although further papers may be published on various other aspects of lipid formation in micro-organisms, this is the final work in which the key role of ME and its attendant gene expression will be investigated. Other interested researchers are therefore free to develop whatever aspect of these studies may seem attractive to them.
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ACKNOWLEDGEMENTS |
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Edited by: N. L. Glass
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
Borsch, D. & Westhoff, P. (1990). Primary structure of NADP+-dependent malic enzyme in the dicotyledonous C4 plant Flaveria trinervia. FEBS Lett 273, 111–115.[CrossRef][Medline]
Botham, P. A. & Ratledge, C. (1979). A biochemical explanation for lipid accumulation in Candida 107 and other oleaginous microorganisms. J Gen Microbiol 114, 361–375.
Boulton, C. A. & Ratledge, C. (1981). Correlation of lipid accumulation in yeasts with possession of ATP : citrate lyase. J Gen Microbiol 127, 169–176.
Broun, P., Gettner, S. & Somerville, C. (1999). Genetic engineering of plant lipids. Annu Rev Nutr 19, 197–216.[CrossRef][Medline]
Castelein, H., Gulick, T., Declercq, P. E., Mannaerts, G. P., Moore, D. D. & Baes, M. I. (1994). The peroxisome proliferator activated receptor regulates malic enzyme gene expression. J Biol Chem 269, 26754–26758.
Ceddia, R. B., William, W. N., Lima, F. B., Flandin, P., Curi, R. & Giacobino, J. P. (2000). Leptin stimulates uncoupling protein-2 mRNA expression and Krebs cycle activity and inhibits lipid synthesis in isolated rat white adipocytes. Eur J Biochem 267, 5952–5958.[Medline]
Chang, G. G. & Tong, L. (2003). Structure and function of malic enzymes, a new class of oxidative decarboxylases. Biochemistry 42, 12721–12733.[CrossRef][Medline]
Chang, G. G., Wang, J. K., Huang, T. M., Lee, H. J., Chou, W. Y. & Meng, C. L. (1991). Purification and characterisation of the NADP+-dependent cytosolic malic enzyme from human cancer cell line. Eur J Biochem 202, 681–688.[Medline]
Cohen, Z. & Ratledge, C. (2005). Single Cell Oils. Champaign, IL: AOCS Press.
Dean, R. A., Talbot, N. J., Ebbole, D. J., Farman, M. L., Mitchell, T. K., Orbach, M. J., Thon, M., Kulkarni, R., Xu, J. R. & other authors (2005). The genome sequence of the rice blast fungus Magnaporthe grisea. Nature 434, 980–986.[CrossRef][Medline]
Drincovich, M. F., Casati, P. & Andreo, C. S. (2001). NADP-malic enzyme from plants: a ubiquitous enzyme involved in different metabolic pathways. FEBS Lett 490, 1–6.[CrossRef][Medline]
Eichinger, L., Pachebat, J. A., Glockner, G., Rajandream, M. A., Sucgang, R., Berriman, M., Song, J., Olsen, R., Szafranski, K. & other authors (2005). The genome of the social amoeba Dictyostelium discoideum. Nature 435, 43–57.[CrossRef][Medline]
Evans, C. T. & Ratledge, C. (1983). Biochemical activities during lipid accumulation in Candida curvata. Lipids 18, 630–635.[Medline]
Evans, C. T. & Ratledge, C. (1984). Phosphofructokinase and the regulation of the flux of carbon from glucose to lipid in the oleaginous yeast Rhodosporidium toruloides. J Gen Microbiol 130, 3251–3264.
Evans, C. T. & Ratledge, C. (1985). Possible regulatory roles of ATP : citrate lyase, malic enzyme and AMP deaminase in lipid accumulation by the oleaginous yeast Rhodosporidium toruloides CBS 14. Can J Microbiol 31, 1000–1005.
Galagan, J. E., Calvo, S. E. & Borkovich, K. A. (2003). The genome sequence of the filamentous fungus Neurospora crassa. Nature 422, 859–868.[CrossRef][Medline]
Galagan, J. E., Calvo, S. E., Cuomo, C., Ma, L. J., Wortman, J. R., Batzoglou, S., Lee, S. I., Basturkmen, M., Spevak, C. C. & other authors (2005). Sequencing of Aspergillus nidulans and comparative analysis with A. fumigatus and A. oryzae. Nature 438, 1105–1115.[CrossRef][Medline]
Hillgartner, F. B. & Charron, T. (1998). Glucose stimulates transcription of fatty acid synthase and malic enxyme in avian hepatocytes. Am J Physiol 274, E493–E501.[Medline]
Kendrick, A. & Ratledge, C. (1992). Desaturation of polyunsaturated fatty acids in Mucor circinelloides and the involvement of a novel membrane-bound malic enzyme. Eur J Biochem 209, 667–673.[Medline]
Kulkarni, G., Cook, P. R. & Harris, B. G. (1993). Cloning and nucleotide sequence of a full-length cDNA encoding Ascaris suum malic enzyme. Arch Biochem Biophys 300, 231–237.[CrossRef][Medline]
Li, Y., Adams, I. P., Wynn, J. P. & Ratledge, C. (2005). Cloning and characterization of a gene encoding a malic enzyme involved in anaerobic growth in Mucor circinelloides. Mycol Res 109, 461–468.[CrossRef][Medline]
Mackenzie, D. A., Wongwathanarat, P., Carter, A. T. & Archer, D. B. (2000). Isolation and use of a homologous histone H4 promoter and a ribosomal DNA region in a transformation vector for the oil-producing fungus Mortierella alpina. Appl Environ Microbiol 66, 4655–4661.
Michaelson, L. V., Lazarus, C. M., Griffiths, G., Napier, J. A. & Stobart, K. A. (1998). Isolation of a 
5-fatty acid desaturase gene from Mortierella alpina. J Biol Chem 273, 19055–19059.
Ohlrogge, J. B. & Jaworski, J. G. (1997). Regulation of fatty acid synthesis. Annu Rev Plant Physiol Plant Mol Biol 48, 109–136.[CrossRef][Medline]
Ramli, U. S., Salas, J. J., Quant, P. A. & Harwood, J. L. (2005). Metabolic control analysis reveals an important role for diacylglycerol acyltransferase in olive but not in oil palm lipid accumulation. FEBS J 272, 5764–5770.[CrossRef][Medline]
Rangasamy, D. & Ratledge, C. (2000). Genetic enhancement of fatty acid synthesis by targeting rat liver ATP : citrate lyase into plastids of tobacco. Plant Physiol 122, 1231–1238.
Ratledge, C. (1997). Microbial lipids. In Biotechnology, 2nd edn, vol. 7, pp.135–197. Edited by H. Kleinkauf & H. Dohren. Weinheim, Germany: VCH.
Ratledge, C. (2004). Fatty acid biosynthesis in microorganisms being used for Single Cell Oil production. Biochimie 86, 807–815.[Medline]
Ratledge, C. (2005). Single Cell Oils for the 21st Century. In Single Cell Oils, pp. 1–20. Edited by Z. Cohen & C. Ratledge. Champaign, IL: AOCS Press.
Ratledge, C. & Gilbert, S. C. (1985). Carnitine acetyltransferase activity in oleaginous yeasts. FEMS Microbiol Lett 27, 273–275.
Ratledge, C. & Hopkins, S. (2006a). Lipids from microbial sources. In Modifying Lipids for Use in Foods, pp. 80–113. Edited by F. Gunstone. Abington, UK: Woodhead Publishing.
Ratledge, C. & Hopkins, S. (2006b). Applications and safety of microbial oils. In Modifying Lipids for Use in Foods, pp. 567–585. Edited by F. Gunstone. Abington, UK: Woodhead Publishing.
Ratledge, C. & Wynn, J. P. (2002). The biochemistry and molecular biology of lipid accumulation in oleaginous microorganisms. Adv Appl Microbiol 51, 1–51.[Medline]
Rognstad, R. & Katz, J. (1979). Effects of 2,4-dihydroxybutyrate on lipogenesis in rat hepatocytes. J Biol Chem 254, 11969–11972.
Rothermel, B. A. & Nelson, T. (1989). Primary structure of the maize NADP+-dependent malic enzyme. J Biol Chem 264, 19587–19592.
Schipper, M. A. A. (1976). On Mucor circinelloides, Mucor racemosus and related species. Stud Mycol 12, 1–40.
Shimomura, I., Shimano, H., Korn, B. S., Bashmakov, Y. & Horton, J. D. (1998). Nuclear sterol regulatory element-binding proteins activate genes responsible for the entire program of unsaturated fatty acid biosynthesis in transgenic mouse liver. J Biol Chem 273, 35299–35306.
Song, Y., Wynn, J. P., Li, Y., Grantham, D. & Ratledge, C. (2001). A pregenetic study of the isoforms of malic enzyme associated with lipid accumulation in Mucor circinelloides. Microbiology 147, 1507–1515.
Sourdioux, M., Brevelet, C., Delabrosse, Y. & Douaire, M. (1999). Association of fatty acid synthase gene and malic enzyme gene polymorphisms with fatness in turkeys. Poult Sci 78, 1651–1657.
Thelen, J. J. & Ohlrogge, J. B. (2002). Metabolic engineering of fatty acid biosynthesis in plants. Metab Eng 4, 12–21.[CrossRef][Medline]
van Heeswijck, R. & Roncero, M. I. (1984). High frequency transformation of Mucor with recombinant plasmid DNA. Carlsberg Res Commun 49, 691–701.[CrossRef]
Velayos, A., Blasco, J. L., Alvarez, M. I., Iturriaga, E. A. & Eslava, A. P. (2000). Blue-light regulation of phytoene dehydrogenase (carB) gene expression in Mucor circinelloides. Planta 210, 938–946.[CrossRef][Medline]
Wierenga, R. K., Terpstra, P. & Hol, W. G. J. (1986). Prediction of the occurrence of the ADP-binding β––β-fold in proteins, using an amino acid sequence fingerprint. J Mol Biol 187, 101–107.[CrossRef][Medline]
Wolff, A. M., Appel, K. F., Petersen, J. B., Poulsen, U. & Arnau, J. (2002). Identification and analysis of genes involved in the control of dimorphism in Mucor circinelloides (syn. racemosus). FEMS Yeast Res 2, 203–213.[Medline]
Wongwathanarat, P., Michaelson, L. V., Carter, A. T., Lazarus, C. M., Griffiths, G., Stobart, A. K., Archer, D. B. & MacKenzie, D. A. (1999). Two fatty acid delta 9-desaturase genes, ole1 and ole2, from Mortierella alpina complement the yeast ole1 mutation. Microbiology 145, 2939–2946.
Wynn, J. P. & Ratledge, C. (1997). Malic enzyme is a major source of NADPH for lipid accumulation by Aspergillus nidulans. Microbiology 143, 253–257.
Wynn, J. P. & Ratledge, C. (2000). Evidence that the rate-limiting step for the biosynthesis of arachidonic acid in Mortierella alpina is at the level of the 18 : 3 to 20 : 3 elongase. Microbiology 146, 2325–2331.
Wynn, J. P., Kendrick, A. & Ratledge, C. (1997). Sesamol as an inhibitor of growth and lipid metabolism in Mucor circinelloides via its action on malic enzyme. Lipids 32, 605–610.[Medline]
Wynn, J. P., Hamid, A. A., Midgley, M. & Ratledge, C. (1998). Widespread occurrence of ATP : citrate lyase and carnitine acetyltransferase in filamentous fungi. World J Microbiol Biotechnol 14, 145–147.[CrossRef]
Wynn, J. P., Hamid, A. A. & Ratledge, C. (1999). The role of malic enzyme in the regulation of lipid accumulation in filamentous fungi. Microbiology 145, 1911–1917.
Wynn, J. P., Hamid, A. A., Li, Y. & Ratledge, C. (2001). Biochemical events leading to the diversion of carbon into storage lipids in the oleaginous fungi Mucor circinelloides and Mortierella alpina. Microbiology 147, 2857–2864.
Yang, Z., Floyd, D. L., Loeber, G. & Tong, L. (2000). Structure of a closed form of human malic enzyme and implications for catalytic mechanism. Nat Struct Biol 7, 251–257.[CrossRef][Medline]
Yang, Z., Zhang, H., Hung, H. C., Kuo, C. C., Tsai, L. C., Yuan, H. S., Chou, W. Y., Chang, G. G. & Tong, L. (2002). Structural studies of the pigeon cytosolic NADP+-dependent malic enzyme. Protein Sci 11, 332–341.[CrossRef][Medline]
Zhang, Y. (2005). A biochemical and molecular study of the roles of malic enzyme in lipid accumulation in Mortierella alpina and Mucor circinelloides. PhD thesis, University of Hull.
Received 20 September 2006;
revised 11 December 2006;
accepted 15 February 2007.
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