Mycelium-bound lipase from a locally isolated strain of Aspergillus flavus link: Pattern and factors involved in its production

Aspergillus flavus produces a lipase (EC 3.1.1.3) which is partly bound to the mycelium during growth. The production of the mycelium-bound lipase is concomitant with growth, and declines when growth ceases. Maximum productivity of the enzyme is obtained when the culture is incubated at 30°C, an initial culture pH of 6·5 and with 2% (w/v) each of corn oil and yeast extract as carbon and organic nitrogen source. Yeast extract affects not only the production of lipase but also the secretion of proteases into the culture medium. Production of the latter enzymes, which inactivate the free lipase, is enhanced by adding yeast extract (1–2%, w/v) to the culture medium. However, at 5% (w/v) yeast extract concentration, proteolytic activity is not detected and consequently, the activity of free lipase may easily be measured. Free lipase activity can easily be detected when 0·001 mol dm⁻³ EDTA is added to the culture medium. The presence of the chelating agent enhances the production and maintains the stability of the extracted mycelium-bound lipase.     

Reactive extraction of acylglycerides using <Emphasis Type="Italic">Aspergillus flavus</Emphasis> resting cells

A one-pot TAG extraction and FAME formation using fungal resting cells and oilseeds at moderate temperature are described. The final yield of methyl esters is increased by the sequential addition of water and methanol. The process can be carried out either with solvents or in a solvent-free system. When a solvent-free medium is used, the final yield will increase if soft oilseeds are used.     

Production of structured lipids containing essential fatty acids by immobilizedRhizopus delemar lipase

An attempt was made to produce structured lipids containing essential fatty acid by acidolysis with 1,3-positional specificRhizopus delemar lipase. The lipase was immobilized on a ceramic carrier by coprecipitation with acetone and then was activated by shaking for 2 d at 30°C in a mixture of 5 g safflower or linseed oil, 10 g caprylic acid, 0.3 g water and 0.6 g of the immobilized enzyme. The activated enzyme was transferred into the same amount of oil/caprylic acid mixture without water, and the mixture was shaken under the same conditions as for the activation. By this reaction, 45–50 mol% of the fatty acids in oils were exchanged for caprylic acid, and the immobilized enzyme could be reused 45 and 55 times for safflower and linseed oils, respectively, without any significant loss of activity. The triglycerides were extracted withn-hexane after the acidolysis and then were allowed to react again with caprylic acid under the same conditions as mentioned above. When acidolysis was repeated three times with safflower oil as a starting material, the only products obtained were 1,3-capryloyl-2-linoleoylglycerol and 1,3-capryloyl-2-oleoyl-glycerol, with a ratio of 86∶14 (w/w). Equally, the products from linseed oil were 1,3-capryloyl-2-α-linolenoyl-glycerol, 1,3-caprylol-2-linoleoyl-glycerol, and 1,3-capryloyl-2-oleoly-glycerol (60∶22∶18, w/w/w). All fatty acids at the 1,3-positions in the original oils were exchanged for caprylic acid by the repeated acidolyses, and the positional specificity ofRhizopus lipase was also confirmed to be strict.     

Effects of oilseed storage proteins on aflatoxin production by Aspergillus flavus

Potential involvement of seed storage proteins in susceptibility to aflatoxin contamination was assessed with in vitro tests. Initially, two oilseed storage proteins [cottonseed storage protein (CSP) and zein] were compared with bovine serum albumin (BSA) and collagen. Supplementation of a complete defined medium with either oilseed storage protein resulted in significantly more aflatoxin production by Aspergillus flavus than supplementation with either BSA or collagen. Little or no aflatoxin was produced when either BSA, CSP, or zein was employed (at 0.5%) as both the sole carbon and the sole nitrogen source. Media with collagen (0.5%) as the sole nitrogen and carbon source supported aflatoxin production similar to the complete defined medium. Although lower than levels observed with defined medium, aflatoxin production increased with both increasing CSP concentration (0 to 2.0%) and increasing zein concentration (0 to 6.0%) when these proteins served as both the sole carbon and sole nitrogen source. Denaturing polyacrylamide gel electrophoresis and protease activity assays indicated that fungal acquisition of protein carbon was probably via hydrolysis mediated by the 35 kD metalloprotease of A. flavus. Media lacking nitrogen but containing sucrose (5.0%) and supplemented with either zein (1.7%) or CSP (2.0%) supported three- to eightfold more aflatoxin production than the complete defined medium. The results suggest seed storage proteins, when present with an accessible carbon source, may predispose oilseed crops to support production of high levels of aflatoxins by A. flavus during seed infection.     

Synthesis of fatty hydroxamic acids catalyzed by the lipase ofMucor miehei

Biotechnological synthesis of a new class of amphiphilic molecules—fatty hydroxamic acids—was carried out using the lipase ofMucor miehei by reacting hydroxyl amine with the fatty acids in their free or methyl ester form. Concurrently with enzymatic synthesis, chemical synthesis of hydroxamic fatty acids has also been developed by adapting methods that already existed for water-soluble acids. Different parameters were studied to determine the optimum operating conditions: temperature, molar ratio of reagents, quantity of biocatalyst and length of reaction. A general method, whatever the type of fatty acids used, is described.     

Utilization of acid oils in making valuable fatty products by microbial lipase technology

Microbial lipase-catalyzed hydrolysis, esterification, and alcoholysis reactions were carried out on acid oils of commerce such as coconut, soybean, mustard, sunflower, and rice bran for the purpose of making fatty acids and various monohydric alcohol esters of fatty acids of the acid oils. Neutral glycerides of the acid oils were hydrolyzed byCanadida cylindracea lipase almost completely within 48 h. Acid oils were converted into fatty acid esters of short- and long-chain alcohols like C₄, C₈, C₁₀, C₁₂, C₁₆, and C₁₈ in high yields by simultaneous esterification and alcoholysis reactions withMucor miehei lipase as catalyst. Acid oils of commerce can be utilized as raw materials in making fatty acids and fatty acid esters using lipase-catalyzed methodologies.     

Plackett-burman design for determining the preference of Rhizomucor miehei lipase for FA in acidolysis reactions with coconut oil

The preference of lipase (EC 3.1.1.3) from Rhizomucor miehei in the incorporation of 11 FA, ranging from C10∶0 to C22∶6, into coconut oil TAG during acidolysis was studied by applying the Plackett-Burman experimental design. Enzymatic acidolysis reactions were carried out in hexane at 37°C for 48 h with coconut oil (0.1 M) and a mixture of 11 FA at a TAG to FA molar ratio of 1∶1. Lipase was used at the 5 wt% level. The incorporation of FA into coconut oil TAG was determined by GC. The lipase showed preference for long-chain saturated FA for incorporation into coconut oil TAG. The FA with 18 carbon atoms showed a high incorporation rate (18∶1>18∶1>18∶3). The lipase showed the least preference for the incorporation of 12∶0, which occurs in maximal concentration (46%), whereas the most preferred FA, 18∶0, occurs at a very low concentration (<2%) in coconut oil. The overall preference of lipase for the incorporation of different FA into coconut oil TAG was 18∶0>18∶2, 22∶0>18∶1, 18∶3, 14∶0, 20∶4, 22∶6>16∶0>12∶0≫10∶0.     

Modification of butterfat by selective hydrolysis and interesterification by lipase: Process and product characterization

Butterfat was chemically modified via combined hydrolysis and interesterification, catalyzed by a commercial lipase immobilized onto a bundle of hydrophobic hollow fibers. The main goal of this research effort was to engineer butterfat with improved nutritional properties by taking advantage of the sn-1,3 specificity and fatty acid specificity of a lipase in hydrolysis and ester interchange reactions, and concomitantly decrease its level of long-chain saturated fatty acid residues (viz., lauric, myristic, and palmitic acids) and change its melting properties. All reactions were carried out at 40°C in a solvent-free system under controlled water activity, and their extent was monitored via chromatographic assays for free fatty acids, esterified fatty acid moieties, and triacylglycerols; the thermal behavior of the modified butterfat was also assessed via calorimetry. Lipase-modified butterfat possesses a wider melting temperature range than regular butterfat. The total saturated triacylglycerols decreased by 2.2%, whereas triacylglycerols with 28–46 acyl carbons (which contained two or three lauric, myristic, or palmitic acid moieties) decreased by 13%. The total monoene triacylglycerols increased by 5.4%, whereas polyene triacylglycerols decreased by 2.9%. The triacylglycerols of interesterified butterfat had ca. 10.9% less lauric, 10.7% less myristic, and 13.6% less palmitic acid residues than those of the original butterfat.     

Trans fatty acid composition and tocopherol content in vegetable oils produced in Mexico

The purpose of this study was to evaluate the trans fatty acid (TFA) composition and the tocopherol content in vegetable oils produced in Mexico. Sample oils were obtained from 18 different oil refining factories, which represent 72% of the total refineries in Mexico. Fatty acids and TFA isomers were determined by gas chromatography using a 100-m fused-silica capillary column (SP-2560). Tocopherol content was quantified by normal-phase high-performance liquid chromatography using an ultraviolet detector and a LiChrosorb Si60 column (25 cm). Results showed that 83% of the samples corresponded to soybean oil. Seventy-two percent of the oils analyzed showed TFA content higher than 1%. Upon comparing the tocopherol contents in some crude oils to their corresponding deodorized samples, a loss of 40–56% was found. The processing conditions should be carefully evaluated in order to reduce the loss of tocopherols and the formation of TFA during refining.     

Soybean phytoalexin,glyceollin, prevents accumulation of aflatoxin B1 in cultures ofAspergillus flavus

The soybean phytoalexin, glyceollin, suppresses the accumulation of aflatoxin B₁ in cultures ofAspergillus flavus. At concentrations of 6.25g/ ml and 62.5g/ml, glyceollin causes 70% and 95% decreases in the maximum observed levels of aflatoxin B₁, respectively. In contrast to the dramatic effect on aflatoxin B₁ levels, these concentrations have little effect on fungal growth. For example, at 62.5g/ml in liquid culture, glyceollin causes a barely discernible lag in the beginning of growth and a 11.5% decrease in maximum fungal mass. When the same concentration of glyceollin is added to the colony margin on semisolid medium, an inhibition zone is formed and then overgrown in one day. Glyceollin appears to act by inhibiting aflatoxin B₁ synthesis, since the rate of aflatoxin B₁ breakdown is not increased in fungal cultures that have been grown in the presence of glyceollin. Glyceollin does accumulate in viable soybean seeds that have been infected withAspergillus flavus. Such seeds accumulate aflatoxin B₁ at one-third the rate of non-glyceollin-producing, nonviable seeds. These results suggest that the synthesis of glyceollin in infected seeds may explain, at least in part, why aflatoxin contamination of soybeans is not a common problem.     

Use of thermostable Fusarium heterosporum lipase for production of structured lipid containing oleic and palmitic acids in organic solvent-free system

A newly developed 1,3-positionally specific thermostable lipase from Fusarium heterosporum (named R275A lipase) was immobilized on Dowex WBA for the production of structured lipid by acidolysis of tripalmitin (PPP) with oleic acid (OA). The immobilized catalyst was fully activated by pretreatment at 50°C in a PPP/OA mixture containing 2% water. The pretreatment caused concomitant hydrolysis, but the hydrolysis was repressed using a substrate without water in the subsequent reactions. The optimal reaction conditions were determined as follows: A mixture of PPP/OA (1∶2, w/w) and 8% immobilized lipase catalyst was incubated at 50°C for 24 h with shaking at 130 oscillations/min. The acidolysis reached 50% under these conditions, and the contents of triolein, 1,3-dioleoyl-2-palmitoyl-glycerol, 1(3),2-dioleoyl-3(1)-palmitoyl-glycerol, 1(3),2-palmitoyl-3(1)-oleoyl-glycerol, 1,3-dipalmitoyl-2-oleoyl-glycerol, and PPP in the reaction mixture were 8, 36, 4, 28, 1, and 6 mol%, respectively. The stabilities of immobilized R275A lipase catalyst and two immobilized catalysts containing Rhizopus delemar or Rhizomucor miehei lipases were compared under the conditions mentioned above, with the catalysts being transferred to fresh substrate every 24 h. The half-life of the R275A lipase catalyst was 370 d, which was significantly longer than those of Rhizopus and Rhizomucor lipase catalysts.     

Deodorization of vegetable oils: Prediction of trans polyunsaturated fatty acid content

Kinetics of the formation of trans linoleic acid and trans linolenic acid were compared. Pilot plant-scale tests on canola oils were carried out to validate the laboratory-scale kinetic model of geometrical isomerization of polyunsaturated fatty acids described in our earlier publication. The reliability of the model was confirmed by statistical calculations. Formation of the individual trans linoleic and linolenic acids was studied, as well as the effect of the degree of isomerization on the distribution of the trans fatty acid isomers. Oil samples were deodorized at temperatures from 204 to 230°C from 2 to 86 h. Results showed an increase in the relative percentage of isomerized linolenic and linoleic acid with an increase in either the deodorization time or the temperature. The percentage of trans linoleic acid (compared to the total) after deodorization ranged from <1 to nearly 6%, whereas the percentage of trans linolenic acid ranged from <1 to >65%. Applying this model, the researchers determined the conditions required to produce a specially isomerized oil for a nutritional study. The practical applications of these trials are as follows: (i) the trans fatty acid level of refined oils can be predicted for given deodorization conditions, (ii) the conditions to meet increasingly strict consumer demands concerning the trans isomer content can be calculated, and (iii) the deodorizer design can be characterized by the deviation from the theoretical trans fatty acid content of the deodorized oil.     

Triglyceride specificity ofCandida cylindracea lipase: Effect of docosahexaenoic acid on resistance of triglyceride to lipase

Tuna oil was hydrolyzed withCandida cylindracea lipase. After 70% hydrolysis of the oil, the docosahexaenoic acid (DHA) content in the glyceride mixture [a mixture of TG (triglyceride), DG (diglyceride) and MG (monoglyceride)] was twice that of the original oil. DHA-rich TG and DG were observed, but DHA-rich MG was absent.C. cylin-dracea lipase seemed to have a “triglyceride specificity,” and it favors TG without DHA over TG containing DHA. In accordance with this hypothesis, TG containing a mixture of oleic acid (OA) and DHA was synthesized and then hydrolyzed withC. cylindracea lipase. TGs in the hydrolysis product were fractionated and analyzed quantitatively by high-performance liquid chromatography. Four kinds of TGs were obtained. TG with three molecules of OA was hydrolyzed most easily. Increasing the DHA content of TG resulted in less hydrolysis of TG. The results suggested thatC. cylindracea lipase had a TG specificity for the whole structure of TG in preference to the individual ester bonds; OA coexisting with DHA in TG was resistant toC. cylindracea lipase due to the TG structure.     

Enzymatic esterification of (−)-menthol with fatty acids in solvent by a commercial lipase from Candida rugosa

Esterification of (−)-menthol with fatty acids in isooctane was successfully catalyzed using a commercial lipase, Lipase AY “Amano” 30 from Candida rugosa in original powder form. The esterification reactions were performed to elucidate the effects of temperature, enzyme load, molar ratio of (−)-menthol/fatty acid, and fatty acid type, keeping the (−)-menthol concentration at 200 mM. At the optimal conditions for (−)-menthol esterification, determined at a (−)-menthol/lauric acid molar ratio of 1∶1 and 35°C [1.5 g enzyme/g (−)-menthol, 0.1 g molecular sieves], the molar conversion of (−)-menthol after 48 h reached 93%. After 24h, the lowest and the highest molar conversions of fatty acids at 2∶1 molar ratio were obtained with myristic acid (71%) and margaric acid (98%), respectively. After 48 h, the molar conversions of lauric acid at molar ratios 2∶1, 1∶1, and 1∶2 were 98, 93, and 49%, respectively.     

Borago officinalis oil: Fatty acid fractionation by immobilized Candida rugosa lipase

γ-Linolenic acid (Z,Z,Z-6,9,12-octadecatrienoic acid), a very important polyunsaturated fatty acid is found in the free fatty acid fraction prepared by the hydrolysis of borage oil. Our aim was to enrich this fraction in γ-linolenic acid using selective esterification. Candida rugosa lipase was used as catalyst after immobilization on the following ion-exchange resins: Amberlite IRC50, IRA35, IRA93, and Duolite A7, A368, A568. In every case, immobilization modified the lipae’s specificity: palmitic, stearic, oleic, and linoleic acids were preferentially esterified compared to γ-linolenic acid, thus allowing a γ-linolenic acid enrichment of 3.0.     

Conversion of vegetable oil to biodiesel using immobilized Candida antarctica lipase

Biodiesel derived from vegetable oils has drawn considerable attention with increasing environmental consciousness. We attempted continuous methanolysis of vegetable oil by an enzymatic process. Immobilized Candida antarctica lipase was found to be the most effective for the methanolysis among lipases tested. The enzyme was inactivated by shaking in a mixture containing more than 1.5 molar equivalents of methanol against the oil. To fully convert the oil to its corresponding methyl esters, at least 3 molar equivalents of methanol are needed. Thus, the reaction was conducted by adding methanol stepwise to avoid lipase inactivation. The first step of the reaction was conducted at 30°C for 10 h in a mixture of oil/methanol (1:1, mol/mol) and 4% immobilized lipase with shaking at 130 oscillations/min. After more than 95% methanol was consumed in ester formation, a second molar equivalent of methanol was added and the reaction continued for 14 h. The third molar equivalent of methanol was finally added and the reaction continued for 24 h (total reaction time, 48 h). This three-step process converted 98.4% of the oil to its corresponding methyl esters. To investigate the stability of the lipase, the three-step methanolysis process was repeated by transferring the immobilized lipase to a fresh substrate mixture. As a result, more than 95% of the ester conversion was maintained even after 50 cycles of the reaction (100 d).     

Lipase‐catalyzed alcoholysis of vegetable oils

Fatty acid alkyl esters were produced from various vegetable oils by transesterification with different alcohols using immobilized lipases. Using n‐hexane as organic solvent, all immobilized lipases tested were found to be active during methanolysis. Highest conversion (97%) was observed with Thermomyces lanuginosa lipase after 24 h. In contrast, this lipase was almost inactive in a solvent‐free reaction medium using methanol or 2‐propanol as alcohol substrates. This could be overcome by a three‐step addition of methanol, which works efficiently for a range of vegetable oils (e.g. cottonseed, peanut, sunflower, palm olein, coconut and palm kernel) using immobilized lipases from Pseudomonas fluorescens (AK lipase) and Rhizomucor miehei (RM lipase). Repeated batch reactions showed that Rhizomucor miehei lipase was very stable over 120 h. AK and RM lipases also showed acceptable conversion levels for cottonseed oil with ethanol, 1‐propanol, 1‐butanol and isobutanol (50‐65% conversion after 24 h) in solvent‐free conditions. Methyl and isopropyl fatty acid esters obtained by enzymatic alcoholysis of natural vegetable oils can find application in biodiesel fuels and cosmetics industry, respectively.     

Acidolysis of Tripalmitin with Oleic Acid Catalyzed by a Newly Isolated Thermostable Lipase

The catalytic efficiency of a lipase from Bacillus stearothermophilus MC7 (lipase MC7) was evaluated in acidolysis of tripalmitin with oleic acid to yield dioleoylpalmitoylglycerol, a structured triglyceride used in health food. The immobilized enzyme exhibits good operational thermostability with a half-life of 50 days at 60 °C in a solvent-free system. The degree of conversion exceeded 50% after 48 h. The side reaction of hydrolysis was suppressed. However, the monosubstituted product was prevalent in the product mixture. Tested in a broad range of solvents, lipase MC7 showed tolerance towards medium polarity.     

Pre-Ripening damage to cottonseed byaspergillus flavus is not influenced by seed coat permeability

Infection of cottonseed (Gossypium hirsutum L.) byAspergillus flavus and associated production of aflatoxins are problems in the arid portions of the United States cotton belt. The hard seed (impermeable to water) characteristic confers resistance to these problems in ripened cottonseed. Experiments were done to determine if low seed coat permeability to water or impermeability protect developing seeds from deterioration and aflatoxin formation. No differences were observed in the degree of seed deterioration in the various cotton lines that could be attributed to seed coat permeability. It is likely that, because the impermeable or low permeability phenotypes are expressed only upon seed ripening, these characteristics afford no protection to cottonseed against attack byA. flavus during seed development. To whom correspondence should be addressed at USDA/ARS, Dept. of Botany and Plant Pathology, Michigan State University, East Lansing, MI 48824-1312.