University of Nigeria

Research Publications

OKEZIE, Stanley M.

PG/M.Sc./0027858 Author

Production of Cocoa Butter Substitutes/Equivalents from Palm Oil, Shea Butter and Dika Fat using Title Aspergillus niger Lipase

Biological Sciences Faculty Faculty

ent Biochemistry Departm

May, 2007 Date Date

Signature

PRODUCTION OF COCOA BUTTER SUBSTITUTESIEQUIVALENTS FROM PALM OIL, SHEA BUTTER AND DIKA FAT USING Aspcigoilltrs niger LIPASE.

OKEZIE, STANLEY MADUAKOLAM (PGIMSc/00/27858)

DEPARTMENT OF BIOCHEMISTRY UNIVERSITY OF NIGERIA NSUKKA

MAY, 2007 TITLE PAGE

PRODUCTION OF COCOA BUTTER SUBSTITUTES/EQUIVALENTS FROM PALM OIL, SHEA BUTTER AND DIKA FAT USING Aspergillus tziger LIPASE. CERTIFICATION PAGE

OKEZIE, MADUAKOLAM STANLEY, a post-graduate student of the Department of Biochemistry and with the Registration Number PG/M.Sc/00/27858, has satisfactorily completed his requirement for research work for the Degree of Master of Science (M.Sc) in Industrial BiochemistryA3iotechnology. The work embodied in this thesis is original and has not been submitted in part or fill1 for any other diploma or degree of this or any other University.

J Prof. I. C. Ononogbu Prof. 0. U. Njoku (Supervisor) (HOD)

Externd Examiner DEDICATION

This work is dedicated to the Almighty God, who in his infinite mercies and powers, have made it possible for me to complete this project. ACKNOWLEDGEMENT

I am particularly grateful to my Supervisor Prof. I.C. Ononogbu who not only made this work possible but also provided the guide. I also thank him for all his generosity and contribution which saw this work to completion. My most profound gratitude also goes to Prof. 0.11. Mioki~for his efforts and support in the entire project. Despite his tight schedules, he still found time to read, criticize and correct the entire project work. I must express my indebtedness to Mr. O.E. lkwuagwu ~vhoby his cosupervision, commitment and guide played a major role in the sirccess of this work. He was also helpful in many other ways too numerous to mention. For various kinds of' contributions and suggestions, I am equally gratefill to Prof. 0. Obidoa, Dr. O.F.C. Nwodo. Dr. F.C Chilaka, Dr. P.N. Uzoegbu, Dr. L.U.S. Ezeanyika, Dr. B.C. Nwanguma, Dr. 1.N.E Onwurah, Dr. E.O. Aiumanah, Dr. V. Ogugua, Dr. S. Eze, Dr. Onwubiko, Mr. P.A.C., Egbuna and Mr. C.U. Anyanwu. I am also grateful to the entire academic and technical staff members of the Depart~nent for all their efforts My colleagues are also warmly acknonfledged-particularly Chinedu Nwuke, Chibuzor Nwaka, Parker Joshua, Dr. Ikgene. Mr. Sabastine, Chukwu Jude. I,. Onyenweaku (Prof.) Mr. I. Iroha and a host of others. Special appreciation goes to Miss Perpetua Offor for all her efforts in the typing of this Project. Special thanks go to my parents, Mr. and Mrs. Okezie, my brothers and sisters, and also my friends. Lastly but by no mems the least, my special ack~iowIedgemntgoes to God Almighty whose plans and purposes must surely prevail.

Okezie S. M. B.Sc (Hons) Nig May 2007 ABSTRACT

Cocoa butter Substitutes/I~quivalents were prepared by enzymatic interesterification of fractions from partially refined palm oil, shea butter and dika fat. The partially retined oil samples were fractionated by crystallization in a solvent medium of 11-hexane. Aspergillus niger lipase immobilized by adsorption on keseilguhr was used as the interesterification catalyst. The lipase was found to have pH and temperature range stability of 6-9 and 25-55" C respectively. Optimal activity of the enzyme was found to be at 30mM oil concentration, pH of 7.2 and temperature of 40' C. The V,,, and K,, of the enzyme was found to be 462u/l and 47mM respectively. Temperature and pH Stability suggest an enzyme suitable for industrial application. Incubation of the oil fractions with the i~nrnobilizedenzyme For 3 days, yielded interesteritied products. Analysis of the products obtained was done using melting point and TLC analysis of their fatty acid concentration. Interesterified products of Palm mid fraction + shea butter crude and palm mid fraction + shea olein were found to have stearic and oleic acid content respectively that compared with that of cocoa butter. The melting point range of palm mid fraction+ shea stearin. palm mid fraction + shea olein, palm olein + shea butter crude, palm mid fraction + shea butter crude, and palm olein + dika olein were found to compare u~ll with the melting point range of cocoa butter. TABLE OF CONTENT CONTENTS PAGE Title page...... i . . Certification Page...... II ... Dedication...... ,111 Acknowledgement...... iv Abstract...... v Table of Contents ...... vi List of Tables...... vii List of Figure...... ix

CHAPTER ONE-INTRODUCTION AND LITERATURE REVIEW Introduction ...... I Lipids an overview...... 3 Fats and oil ...... 3 Nature composition and uses of selected fats and oils...... 6 Industrial Processing of Fats and Oils...... I I Characterization of fats and oils ...... I5 Lipases...... I9

CHAPTER TWO-MATERIALS AND METHODS Materials...... 26 Oil seeds ...... 26 C helnicals...... 26 Methods...... 26 Preparation of reagents...... 26 Extraction of oil from samples ...... 28 Partial refining of oil samples...... 28 Determination of % oil yield ....;...... 36 Solvent fractionation (crystallization) ...... 29 Characterization of ttie partially refined oil samples...... 29 Production and extraction of Fungal Iipase...... 33 2.2.8 Characterization of lipase ...... 34 2.2.9 Immobilization of the Iipase ...... 35 2.2.10 Interesterification reaction system ...... 36 2.2.1 1 Fatty acid analysis of hydrolysates ...... 36 2.2.12 Statistical analysis ...... 38

3.0 CHAPTER THREE-RESULTS 3.1 Physical properties of crude and partially refined oil samples ...... 39 3.2 Chemical properties of partially refined oil estracts and their Fraction ...... 46 3.3 Enzyme properties of A.spergillw nigw lipase ...... SO 3.4 Physical and chemical properties of intercsterified products ...... 55

4.0 CHAPTER FOUR-DISCUSSION Discussion ...... 58 Conclusion ...... 65 Suggestion ...... 66 References ...... 67 LIST OF TABLES Table 1.1 a Foods uses ol'fats and oils ...... 4 Table I . 1 b Industrial uses of fats and oils ...... 5 Table 1.2 Fatty acid composition of dika fat ...... 7 Table 1.3 Fatty acid composition of shea butter ...... 9 Table 1.4 Some specific lipases for the production of specific structured triacylgycerols ...... 24 Table 3.1 Yield of oil fiom different oil fruitstseeds ...... 39 Table 3.2 Acylglycerol fractions of palm oil at varying temperati~re and solvent omcentrations ...... 40 Table 3.3 Acylglycerol fractio~isof shea butter at varying temperature and solvent concentrations ...... 41 Table 3.4 AcylglyceroI fractions of dika fat at varying temperature and solvent clmcentrations ...... 42 Table 3.5 physical properties of partially refined (PR) palm oil and its fracticns ...... 43 Table 3.6 Physical properties of PR shea butter and its fractions ...... 44 'Table 3.7 Physical properties of PR dika fat and its fractions ...... 45 Table 3.5 Physical properties of PR cocoa butter ...... 46 Table 3.9 Chemical properties of PR palm oil and its fractions ...... 47 Tablc 3.10 Chcmical propct-tics of PR rcfincd shca butter and its fractions ...... 48 Table 3.1 1 Chemical property of PR dika fat and its fractions ...... 49 Table 3.12 Chemical property of PR cocoa butter ...... 49 Table 3.13 the melting / slip points of the interesterified products of fractions from palm oil, shea butter and dika fat ...... 54 Table 3.14 Oleic. stearic and palmitic acid contents of 5 Selected interesterified products ...... 56 LIST OF FIGURES Figure I Effect of substrate concentration on lipasc activity ...... 50 Figure 2 Line Weaver Burk plot of enzynle activity against substrate concentration ...... 51 Figure 3 Effect of pH on enzyme activity ...... 51 . . Figure 4 Effect of temperature on enzyme actwty ...... 52 Figure 5 Effect of quantity OF immobilization support on enzyme activity ...... 52 Figure 6 Standard curve for Oleic acid ...... 79 Figure 7 Standard curve for Palmitic acid ...... 80 Figure 8 Standard cul-ve for Stearic acid ...... 81 CHAPTER ONE 1.0 INTRODUCTION AND LITERATURE REVIEW

II INTRODUCTION. About 400 different botan~cspecies are known from whose seeds or fruits oil can be obtained. Oils from different origin at times could have very close similarities but no matter how close they may be it IS practically impossible to find two of them identical. Each has some distinctive chemical characteristics, infrequently physical, sometimes qualitative and more frequently quantitative differences. Saturated and unsaturated acids such as lauric, palmitic, stearic, oleic, linoleic, linolenic, myristic and arachidic acids are the major components of common seed oils (Klopfenstein and Walker, 1983). Other saturated and unsaturated acids are often present as major or minor components as In the cases of erucic acid in rapeseed oil and ricinoleic acid in castor oil.

For oils and fats, ~t is well known that the functional and nutritional properties of their triacylglycerols depend not only on their fatty acid profiles but also on the fatty acid distribution on the glycerol backbone (Macrae and Hammond, 7985; Macrae, 1989, 1992) Therefore, production of fats and oils with desired physical and chemical properties by replacing the fatty acid moieties of triacylglycerol with other fatty acids is of great importance and interest from an industrial viewpoint (Yokoreki gt aJ., 1982).

Fats and oils have a very wide application in the industries. Areas where lipids can play a significant role in industry are cooking, paints production, emulsifiers for tableting, body creams and oils, fat emulsions for parenteral nutrition, anti-oxidant, soap production, butter and margarine, cream and mayonnaise and waxes (Ononogbu, 2002). Triacylglycerol could therefore be regiospecifically modified or restructured to meet special functions for nutrition, food, pharmaceutical, industrial and other applications. This can often be done either chemically or enzymatically by interesterificatian. Chemical interesterification is an advanced technology in industry and only randomized products can usually be produced (Xu, 2000). Enzymatic interesterification of fats and oils with Sn-I, 3 specific lipases can lead to introduction of desirable fatty acids into the triacylglycerol molecule and production of preferred positional fatty acid arrangement in the triacylglycerol molecules (Young et al., 1994). Enzymatic inter-esterification has been employed by industry for the production of cocoa butter-like fats and human milk fat substitutes (Quinlan and Moore, 7993). Cocoa butter ranks among the most costly edible oil in the world comprising rnainly of 1,3-dipalmitoyl-2- oleoyl glycerol (POP), I-palmitoyl-2oleoyl-3-stearoyl glycerol (POS) and 1,3-distearoyl-2- oleoyl glycerol (SOS) (Owusu -Ansah 1994). With the use of 1,3-specific lipases, it is possible by inter-esterification to produce valuable cocoa butter substitutes/equivalents from cheaper starting materials (Macrea, l983., Bloomer d., 1990;Mojovic g

-.al I I993.,Shukla,1996.,Liu ,el a.,1997). The resultant products are known variously as cocoa butter substitutes (CBS) , cocoa butter equivalents (CBE) and cocoa butter replacements fats (CBR).To formulate cocoa butter substituteslequivalents, fats with similar triacylglycerol to cocoa butter are needed as starting materials. Palm oil and seed kernel from tropical trees possess them (Timm, 2QOO).

Objective The uncertainty in cocoa butter supply and the unpredictable changes in its price have compelled confection producers to seek for alternatives. The major trend in search of cocoabutter alternatives has been the modification of suitable vegetable fatshils into cocoa butter substitutes by lipase-catalysed inter-esterification. Studies In India (Sridhar gt a., 1991) and China (Zhang a a,, 1997) have'shown that cocoa butter substitutes can be successfully produced from suitable local vegetable oitslfats by lipase-catalysed inter- esterification.Cocoa butter substitutes have also been successfully produced by lipase catalysed inter-esterifrcation of the mid-fraction of palm oil (Mojovic et al 79931, a Nigerian local vegetable oil. However no work appears to have been done on lipase-catalysed inter- esterification of other fraction of palm oil and on other local vegetable oilslfats.The objective of this study therefore is the modification of selected local vegetable oilstfats notably ,palm oil,shea butter and dika fat into cocoa butter substitutes by lipase-catalysed inter- esterification.

The specific objectives of the project are: To ascertain the status of the various'fractions obtained by refining palm oil ,shea butter and dika fat as potential cocoa butter substitutes,using physico-chemical parameters. To isolate and characterize lipase from Aspergillus niger for inter-esterification of vegetable oil/fat fractions. To modify fractions of palm oil, shea butter and dika fat into cocoa butter substitutes/equivalents by inter-esterification using immobilized llpase from Aspergillus niger.

LITERATURE REVIEW I An overview of Lipids. Lipids are substances, which are soluble in organic solvents but very sparingly soluble in aqueous solvents (Ononogbu, 1990). They are a heterogenous group of compounds related more by their physical rather than by their chemical properties. Thus, lipids include fats, oils, steroids, waxes and related compounds. other lipids although present in relatively small quantities, play crucial roles as enzymes cofactors, e7ectron carriers, light-absorbing pigments, hydrophobic anchors, emulsifying agents, hormones and intracellular messengers (Nelson and Cox, 2000).

1.2.2 Fats and Oils Fats and oils are fatty esters of glycerol and high alcohols. The natural oils and fats are complex mixtures of mixed triacylglycerol with (frequently) small amounts of other lipids such as phosphatides, sterol esters, fatty acids and waxes, together with fat-soluble pigments such as carotenoids .The classical definition of fat is triacyglycerol that is derived from animal source while oil is the triasyglycerol that is derived from plant source (Ononogbu, 2002). Despite this definition, differences in physical 'properties still exist among fats and oils from different sources. Glycerol esters of short chain fatty acids give softer fats than those of long- chain fatty acids. Triacylg;ycerols containing substantial proportions of unsaturated fatty acids are usually liquid oil at normal ambient temperatures, while the more saturated fats and oils with substantial proportion of saturated fatty acids are usually solid.

Fats and oils have been used throughout the ages as food, fuels, lubricants and starting materials for other compounds. Currently, technological advances have played a decisive part in increasing the value and versatility of fats and oils in the edible and industrial market (Hatje, 1989). The major part (about 80%) of fats and oils produced are used for edible 4

purposes while the rest are processed in the industrial sector. Some examples of the wide uses of fats and oils are shown in the tables below. Table I.la Food uses of oils and fats. I Uses Type of oillfat Peanut oil ra round nut oil), Pistachio oil, Grape seed oil and Pine nut oil.

Cooking oil/ Frying oils Coconut oil, Canola oil, Olive oil, Corn oil, Palm oil, Safflower oil, sunflower ail, Almond oil, Pecan oil, Bottle gourd oif, Pumpkin oil, Water melon seed oiF, Amaranth oil, Artichoke oil, cohune oil, Babassu oil, false flax oil, Hemp oil, Kapok seed oil, Mustard oil, Perilla seed oil, Pequi oil, Pine nut oil, Poppyseed oil, Prune Kernel oil, Quinoa oil, Ramitil oil, tea oil, Argan oil, and Avocado oil. Groundnut oil, Grape seed oil, Rice bran Flavoring oil. Hazelnut oil, Walnut oil, Coriander seed oil, Wheat germ oil, Orange oil. Coconut oil, Borneo tallow nut oil, Cohune oil, Cocoa butter, Babassu oil. Palm oil. Confectionary & Baking fat Palm oil, Almond oil, Pecan oil, Cocoanut oil. Acai oil, Black currant seed oil, Evening Margarine prime rose oil, Wheat germ oil.

Supplement -- Table 1.1b Industrial uses of FatslOils Uses Type of FatlOil Pharmaceuticals Almond oil, Cashew oil, Cocoa butter

Medicinal & Antiseptics Borage seed oil, Borneo tallow nut oil, Carobpod oil, Coriander seed oil, Perilla seed oil, wheat germ oil, Amur cork tree fruit oil, Brucea havanica oil, Burdock oil, Castor oil, Chaulmoogra oil, Mowrah fat, Snow ball seed oil, Sea buckthorn oil, Lemon oil, Almond oil, Cashew oil, Shea butter Hazelnut oil, Cotton seed oil, Olive oil, Cosmetics & Skin care Coconut oil, Acai oil, Amaranth oil, Avacado oil, Borneo tallow oil, Cohune oil, Grape seed oil, Perilla seed oil, Poppy seed oil, thistle oil, Apple seed oil, Burdock oil, Candle nut oil, Carrot seed oil, Lemon oil, Neem dl, Rose hip seed oil, Shea butter Mowrah butter, Mango oil, Tea oil, Kapok seed oil, Borneo tallow nut oil, Palm oil, Palm kernel oil. Soaps and Cleaning products. Dammar oil, Lin seed oil, Poppy seed oil, Stillingia oil (Chinese vegetable tallow oil), Tung oil, Vernonia oil, Safflower oil, Walnut Paints & Varnishes oil. Castor oil, Cuphea oil, Snow bail seed oil, Bladder pod oil, Crambe oil Vernonia oil. Crambe oil Chemicals Olive oil, Ramtil oil, Damrnar oil, Mowrah Lubricants butter. Amur cork tree fruit oil, Neem oil. Candles & Lighting Orange oil, Tonka bean oil, lemon oil, .Balanos oil. Insecticides Palm oil, Sunflower oil, Artichoke oil, Perfumes & Fragrances Ben oil, false flax oil. Castor oil, Algae oil, Copaiba oil, Honge oil, Jatropha oil, and Biofuel & Biodiesel. Jojoba oil,

In recent years, competitive products derived from petroleum have been developed for industrial purposes (Hatje 1989).

4.2.3 Nature, Composition and 'Uses of Selected Fats and Oils Palm Oil Oil palm (Ellaies guineenses) belongs to the family Aracaceae. The oil palm tree originated in West Guinea and was introduced to other parts of Africa, South East Asia and Latin America (Ononogbu, 2002). The fruit of the oil palm grows in bunches, each bunch weighing up to 20kg and containing of the order of 1500 fruit (Young a.,1994). Palm oil is obtained from the fibrous and pulpy mesocarp that surrounds the nut or kernel, which is enclosed by a woody shell. It is one of four major oils in the world (Ong, 1989). Palm oil is made up of triacylglycerols and the component fatty acids include palmitic ac~d (40.8%), stearic acid (4.7%) oleic acid (41.1%) and finoleic acid (8.8%) (Ononogbu, 2002). It also contains other minor components such as tocopherols (Vit E) and tocotrienols, betacarotene, beta-sistosterols and squalene, which are of nutritional importance (Ong, 1989). Palm oil is a semi-solid fat and as such, it needs no hydrogenation prior to use in food applications (Gordon 1993). It is often used as a camponent in such products as margarine and biscuit fat blends. Palm oil may be fractionated into several components. With single- stage fractionation, a low cloud point palm dein which can be used as a substitute for soft oil is obtained and a stearin of melting point 44-50'~ which is used as a margarine or shortening hard stock (Porirn, 1981). Double solvent fractionation in which the olein from a first separation is refractionated, produces a mid-fraction that is used in the production of cocoa butter substitutes (Yomg a,,1994). (b) Dika Fat Dika fat is the oil produced from the seeds of bush mango tree (~ingiagabonensis). The wild mango belongs to the family Anacardiaceae and is of the lrvingia species. It is found in southern Senegal and Zaire (Keay, 1989). It is also found in the lowland warm humid forest zone of West and Central Africa. Irvingia gabonensis assumes greater importance in Nigeria and Cameroon than in other parts of humid lowlands of West Africa (Bennuah gt d.,1994). This is because its seeds are used as traditional soup condiments and as a seasoner for various dishes. The ripe fruit is eaten for nutrients, flavour and aroma and is claimed to whiten the teeth (Umoh and Oke, 1978). The fatty matter obtained from the seeds is termed "dika butter" in Gabon (Eka, 1980). The seeds have been shown to yield varying percentages of fatty material ranging from 55-70% (Ononogbu, 2002), 71.97% (Okolo, 1994), 54-67% ['Urnoh and Oke,1978) to 68-75% (Onwuka g aJ., 1984). lrvine (1961) had obtained 54-67% fat while Dalziel (1955) had recorded fatty extract of 71.95%. Amino acid analyses shows that the seed is very deficient in the sulphur amino acids methionine and cystine but high in lysine, leucine, isoleucine and threonine (Abaelu

-et 1.,1980). Okolo (1994) reported that the seed contains 8.56% protein. In/ingia gabonensis seed fat contain triacylglycerol as its major component and phospholipid as a minor component (Eka 1980). Njoku & al., (1997) obtained 65.7% gum, 0.05mglg gossypol, 14.8% phospholipid, 8.08mgtg sterol and 0 7 mgtg p-carotene Analysis of the fatty acid composition shaws that there exist eight fatty acids as shown in the table 1.2. Table 1.2 Fatty acid composition of Dika Fat. Fatty acid % total Lipid Lauric acid 40.40 Myristic acid 42.75 Palmitic acid 6.56 Oleic acid 1.88 Capric acid 1.60 Stearic acid 2.21 kinoleic acid 2.0 Linolenic acid - - Unidentified fractiorl 3.60 Source: Eka (1980) (c) SheaButter Shea butter is obtained from the tree Butyrospermum paradoxurn. The shea butter tree is of the Sapotaceae family and is found growing wild in West Africa and Uganda (Gordon, 1993). In Nigeria, it is "ound mostly in the Middle Belt area where it grows wild in the bush. The fruit of shea butter tree is sweet while the nut contains the kernel, which is the source of a natural fat known as 'shea butter'. Shea butter seed is rich in oil, which is about 50-60.7% (Gordon, 1993). Traditionally, shea butter is extracted by first pounding the kernels in wooden mortar with pestle followed by grinding with stone to obtain an oily paste. The paste is boiled in water for about one hour and the oil floating on the surface of the water is skimmed off. In the Northern part of Nigeria shea butter is commonly used as a frying and cooking oil but that is not the case in the Southern part of the country. However, it is known to some people of Nigeria for its local medicinal value in the treatment of cough and minor bone dislocation (Badifu, 1989).1t also used as an embrocation. In terms of proximate chemical composition, shea butter is reported to consist of crude protein 4.1%, moisture 7.3%, crude fibre 7.5%, ash 3 6% and carbohydrates 4.0% (Farinu, 1986). The fat had 51.7% and 48.3% level of saturation and unsaturation, respectively (Badifu and Abah, 1998). Toxicological studies of shea butter indicated the absence cf gossypol and mycotoxins, while chemical analysis revealed the presence of phospholipids, sterols and glycolipids (Njoku gt aJ., 2000). Shea butter has one of the highest unsaponifiable matter content of any vegetable oil (Gordon,1993). The unsaponifiable matter comprises of a mixture of terpenoid hydrocarbons. Shea butter can be processed and solvent-fractionated to give hard and soft fats. The hard stearin fraction can be used as a constituent of chocolate or a component in cocoa butter equivalent formulation. It can also be used in the cosmetic industry. Stearic acid and oleic acid were found to be the major fatty acids'present in shea butter as shown on Table 1.3 below Table 1.3 - Fatty acid composition of shea butter. Fatty Acid Composition (% total methyllesters) Palmitic 4.2 Stearic 47.5 Oleic 41.8 Linoleic 6.5 Total saturates 51 -7 Total unsaturates 48.3 Source : Badifu and Abah, (1998).

(d) Cocoa butter Cocoa butter is a unique simple fat obtained from pressing ground, roasted, decorticated cocoa beans (Gordon, 1993). In the liquid state, it has a bright yellow colour but yellowish whlte in solid form (Catsberg and Kernpen van Dommelen, 1990). It has a typical composi'tion of 26% palmitic acid, 36% stearic acid, 33% oleic acid and 3% linoleic acid, although there are slight variations in composition depending on the geographical source (Owusu -Ansah, 1994).

Cocoa butter IS used extensively in the making of chocolate and in other confectionery products, and to a limited extent, in the pharrnaceutlcal industry as theobrorna oil (Gordon, 1993). It has also been used in cosmetic formulas for making suntan products, toilet soaps, suppositories, ointments, creams, lotions and for lipstick (Zoumas and F~nnegan,1979) It 1s particularly suitable for these purposes because ~t has a low but sharp melting point, it is brittle and fractures readily and whilst it is not greasy to the touch, it melts completely in the mouth (Gordon, 1993). These properties are a reflection of its triacylglycerol composition comprising mainly of I-palmitoyl-2oleoyl-3-steamy1 glycerol (POS), 1,3-distearoyl-2-oleoyi glycerol (SOS) and 1,3-dipalmitoyl-20leoy1 glycerol (POP). In all the triacylglycerols present in cocoa butter, the 2-or central hydroxyl group on the glycerol backbone of the triacylglycerol molecule is esterifled with an unsaturated acid (Macrae, 1983). (e) Cocoa Butter Substitutes and Equivalent The price of fats has varied considerably over the years but cocoa butter is normally one of the most expensive. The high and variable cost has stimulated an extensive search for fats that could be used to replace or substitute cocoa butter in the industries.

Attempts have therefore been made to produce substitutes mimicking the characteristics of cocoa butter using cheaper and available oil sources (Liu a aJ., 1997). These are known variously as cocoa butter replacement fats, cocoa butter substitutes, and cocoa butter equivalents. Cocoa butter substitutes do not chemically resemble cocoa butter and are compatible with cocoa butter only within specified limits. In contrast, Cocoa butter equivalents are chemically similar to cocoa butter and can replace cocoa butter in any proportion without deleterious physical effects (Zoumas gt 4.,1979). Cocoa butter substitutes and equivalents differ greatfy with respect to their method of manufacture, sources of fats and functionality. Processes for producing such product groups include fractionation, hydrogenation and inter-esterification (Hellstrom, 2000). Except for fractionation, which has been fairly successfully used (Chakrabarty eJ aJ., 1983), the other chemical processes have drawbacks, which preclude their exactness in duplicating the characteristics of cocoa butter. Chemical inter-esterification leads to randomization and therefore, does not yield exactly the major triacylglycerols of cocoa butter. Hydrogenation is unpreventably associated with trans isomer formation, which affects the melting 'characteristics of the final product and thus making it difficult to mimic the melting properties of cocoa butter.

The use of 1,3- specific lipases capable of modifying the 1,3-fatty acid composition of oils through inter-esterification has made it possible to produce cocoa butter-like product resembling cocoa butter in both composition and functional characteristics. There has been strict control and variations in the acceptance of cocoa butter substitutes and equivalents. The legislatio~sconcerning their use in food products vary from country to country. However, cocoa butter equivalents produced by lipase inter-esterification are granted GRAS (generally recognized as safe) status (Anon, 1988). Thus more research in this area is hereby recommended as such areas of interest has been shown to be a major income earner for developing countries as has. been proved in Malaysia, India and Indonesia where such product groups are currently being produced and exported to Europe and other advanced countries of the world. 1.2.4 Industrial Processing of Fats and Oils The quality of a fat or oil product is influenced by many factors. In the case of vegetable raw materials, these include seed or plant type, growth conditions, pests and rnicroarganisms, chemical and weed contaminant, harvesting and storage (Young @ d.,1994). Therefore, the processes of extraction, handling of the crude oil extract, refining, modifying, processing, packaging and distribution also affect the quality of fats and oil products.

Oil bearing fruits such as oil palm has h~ghmorsture content and is favourable to enzymic action (Young gt a.,j994). Consequently, they must be processed promptly after harvesting to minimize degradation of the oil Other groups of oil bearing seeds with lower moisture content are less liable to deterioration and may be stored under suitable conditions for long periods (Young gt aJ., 1994). It has been suggested that the maximum permissible figure for safe storage of oil seeds generally is 14% af moisture calculated on the non-oleaginous portion of the seed (Young gt gl.,1994). However, even lower moisture contents are required to prevent the growth of moulds and the safe limit is the moisture content attained when the material is in equilibrium with an atmosphere of 70% relative humidity (Young @ 4 , 1994) Not only the enzymes within the seed cause deterioration of the seed in storage but also by the external effect of insects, yeasts, mould and bacteria, especially, on split or damaged seeds (Young gt a.,1994).

There exist a wild range of differences in physical characteristics of oil seeds, nuts and fruits. Therefore, both rudimentaqt and sophisticated systems have been developed to extract their oils and process them to finished products. The common objective is to maximize the yield of fat or oil from the materials, minimize the damage to the fat or oil, and produce components as free as possible from undesirable impurities and produce a residual oil cake of the greatest possible value (Carr, 1989). The preparation of or1 bearing materials prior to extraction, is in all cases, concerned, first, with separation of extraneous matter and secondly, transforming the material into a state that will give a high yield of good quality oil meat (Young gt A,,1994).

(a) Oil Extraction Conventional methods for extracting oils involve three basic approaches, namely, physical, chemical and a combination of both (Owusu-Ansah, 1994). The method employed depends generally on the oil content of the raw materials. Physical method is employed for oil seeds of high oil content, e.g., groundnut and palm kernel. While chemical method is used primarily for oil seeds, of low oil content, e.g., soybeans and rice bran (Owusu-Ansah, 1994).

The methods used could affect the physical and chemical properties of the oil or fat to a considerable extent. In selecting solvent for extraction, the solubility of the fat or oil in the solvent, solvent toxicity and the intended use of the oil are of utmost importance. Petroleum ether, hexane, methanol and chloroform are frequently used (Christie, 1982).

Enzymes have found use in oil extraction. The application of enzyme in oil extraction can be categorized into enzyme-assisted pressing; enzyme enhanced solvent extraction and enzyme assisted aqueous extraction (Owusu-Ansah, 1994). In all these approaches, the enzymes are used to break the cell walls of the oil-bearing materials to release the oil.

(b) Refining The further processing of edible oils after extraction from the raw materials is concerned with refining and modification (Young gt a,,1994). Refining treatment is needed to remove or reduce as far as possible, those contaminants of the crude oil, which will adversely affect the quality of the end product and the efficient operalion of the modification process. Two methods are in use for the refining of oils and fats. These are termed physical and chemical methods. The difference between the two methods lies in the means by which free fatty acids are removed from the oil (Young gt a., 1994). In the physical method the fatty acids are distilled off while in the chemical method, the fatty acids are neutralized using alkaline reagent thus forming soaps, which are removed from the oil by phase separation.

(c) Bleaching Pigments such as caroteroids, chlorophyll, gossypol and related compounds and the products of degradation and condensation reactions that occur during the handling, storage and treatment of the extracted oils are removed by bleaching. It was later realized that activated adsorbents, in particular are responsible for removing, at least partially, other impurities such as soaps, trace metals, phosphatides and compounds containing sulphur (Mag, 1990). Oxidation levels are also reduced by breakdown of the hydroperoxide primary oxidation products on the adsorbent surface followed by adsorption of the carbonyl compounds that are the secondary oxidation products (Mag, 1990). The process is usually carried out by treating the oil with adsorbents such as special clays and charcoals at high temperature (100-1 1O'C) and under reduced pressure (Haraldsson,1983). The operation, however, provides a lighter coloured oil and prepares it for subsequent processing.

(d) Modification The modification processes are used to widen the applicability of oils and fats and also to make products of acceptable oxidative and flavour stability from unstable or relatively unstable raw materials. Tie process consists of hydrogenation, inter-esterification and fractionation (Young gt gl.,1994). They can be used alone or in combination with each other and they can be used on single oils or on fat blends.

(e) Hydrogenation Most of the fatty acids of which the triacylgfycerols of naturally occurring oils and fats are composed are unsaturated. Hydrogenation process is used to provide the direct addition of hydrogen into the unsaturated double bonds of the fatty acid chains within neutral oils. When hydrogen is added to fatty acid double bond, it becomes saturated with consequent increases in the oxidative stability and melting point of the oil of which it is part (Young eta., 1994). The process involves reacting gaseous hydrogen, liquid oil and nickel or copper catalyst by mechanical agitation at a specific temperature (150 - 180'~)and pressure (3-5 atm) in a closed reaction vessel. The reaction is directed by changes in conditions affecting mass transfer of hydrogen to catalyst surface and of oil to and from the surface (Young gt aJ., 1994). The reaction end polnt is controlled by determining the refractive index, which relates relatively closely to the iodirle value. Iodine value of zero is not desirable for product will be brittle and unpalatable (Carr, 1989).

(f) Fractionation Fats and oils are mixtures of triacylglycerols having different fatty acid compositions. They have melting points spanning from -50 to 80%, each oil having its own melting range. The melting range limits the use of a particular oil or fat (Young gt d.,1994). Fractionation is a thermomechanical process by which the raw materials are separated into two or more portions which widens the use of the oil. Thermomechanical separation processes include distillation and crystallizaticn. Distillation is commercially unsuitable for the fractionation of triacylglycerol mixtures because of their low vapour pressures and because of their relative instablility at high temperatures (Young gt gl,,1994). Separation can however be effected by crystallization. Crystallization of the oil can be done using three methods, namely, the dry process, the lanza or lipof-ac process and the solvent process (Young & a., 11994). The difference in efficiency between the three techniques lies in the degree of contamination of the stearin by the olein after separation. Due to the high viscosity of the crystalline slurry, separation of the olein from the stearin is difficult. In solvent process, the oil is crystallized using mainly acetone or hexane as the solvent. In this way the viscosity is reduced thereby improving crystallization and filterability (Fritz, .1998). Fractionation process can be used for extending the applicability of fats.

(g) Inter-esterification. Natural fats are mixtures of triacylglycerols in which the acyl groups are wsually distributed in a random manner. Under the influence of an appropriate catalyst, the acyl groups are redistributed first intramolecularly and then intermolecularly until a wholly random distribution is finally achieved. Inter-esterification is the name given to this process in which the arrangement of fatty acid in a triacylglycerol molecule is changed. Chemical inter- esterification leads to a random distribution of the fatty acids on the glycerol molecules. This is known as random inter-esterification. Catalysts usually employed are sodium hydroxide, sodium methoxide and a sodium potassium alloy at 0.2-0.4% level (Young gt aJ., 1994). The redistribution of acyl groups leads to a change in physicochemical properties of the triacylglycerol mixture.

An extension of the process has been introduced by the use of lipases as catalyst for oil and fat inter-esterification reactions (Macrae,1983). Its advantage over the more conventional procedures lies in the additional control of product composition (Xu, 2000). Inter-esterification procedures are used industrially to improve the physical properties of lard, to produce cocoa butter substitutes from cheaper oils {usually combined with hydrogenation and fractionation), to produce fats containing acetic acid and to produce oils of appropriate melting behaviour with a minimum content of polyene acids (Young gt aJ., 1994). 1.2.5 Characterization of Fats and Oils The characteristics of fats and oils are dependent on the physico-chemical properties, which help in the identification and analysis of the individual fats and oils.

(a) Physical Properties The physical properties of 'ats and oils are determined or influenced largely by the nature of their constituent fatty acids although the presence of minor components can also be important. Thus, fats which contain a large proportion of saturated fatty acids, tend to be solids at room temperature. Nevertheless, variations, due to climatic and seasonal conditions and variety of vegetable oil affect the physical properties (Kochhar, 1986). Some of the physical properties are briefly discussed below: -

Organoleptic Properties: Pure fats and oils and their constituent fatty acids are generally colourless, tasteless and odourless. Any colour, odour and taste in fats and oils are due to non-triacylglycerol components such as pigments (chlorophyll, carotenoids), oxidation products (aldehydes and k~tones),unsaturated fatty acids and sterols (Ononogbu, 2002). The colour, odour and flavour of an oil are of considerable commercial importance. Pale bright colours, for example, is desired in refined oils, as this is regarded as the criterion of quality and purity, and also facilitate production of finished products with desired colour shade (Kaimal and Lakshminarayana, 1993).

Solubility and Miscibility: In general, fats and oils are insoluble in water, since they contain predominantly non-polar groups. Solubility is determined to a large extent by the constituent fatty acids. The long chain fatty acids are insoluble in water, while those of the short chain and unsaturated fatty acids are greater in solubility. Saturation and increasing chain length tend to decrease solubility (Christie and Williams, 1982). The solubility of fats in alcohol varies with the nature of the combined fatty acids. The unsaturated and the short chain acids being more soluble than those of the long chain fatty acids (Macrae, 1992). The hydroxy fatty acids themselves are insoluble in petroleum ether. However, at very high temperature and pressure, fats dissolve in large amount in water (Christie and Williams, 1982).

Melting Point I Slip Point: Natural fats are complex mixtures of glycerides and have no sharp melting point since they are mixtures of trialylglycerols each of which is different in molecular weight and often also in degree of unsaturation. Natural oils have zones of melting as one triacylglycerol after another loses it's crystalline form (Hawthorn, 1981). To obtain reliable results for the melting point of whole fats, it is necessary to follow carefully a standardized procedure. One common method used is the slip melting point method. This is the temperature at which a column of fat of specified length rises in an open capillary tube suspended in a water bath of gradually increasing temperature (Gordon, 1993). Stabilization of the fat before measurement of the slip melting paint is extremely important.

Density 1 Specific Gravity: The relative density or specific gravity of an oil is defined as the ratio of the apparent mass, determined by weight in air, of a given volume of the oil at tOcto that of the same volume of water at 20'~;whereas the apparent density (at tOc) is the apparent mass in grams, determined by weighing in air, of Iml of fat at t•‹C (Gordon, 1993). Density and specific gravity are used to convert volumes to weights. They are used to calculate the weight of oils in large tanks. For the liquid fats, the density varies directly with the degree of unsaturation and indirectly with the molecular weight (Applewhite, 1980). Specific gravity is usully measured at 15'~. In tropical regions this temperature is usually unobtainable and thus it can be measured at any temperature tOcup to the temperature of boiling water and then corrected as shown in the formula below (Onyechi, 1987). Specific gravity at (15'~)= specific gravity at t')~(tk - 15'~)x 0.000069.

Refractive Index: Refractive index is helpful in the identification of oils, It can be determined either by means of the Zeiss refractometer, which reads on an arbitrary scale, or by means of the Abbe refractometer, which gives the true refractive index. Refractive Index is the quotient of the sine of the incident angle of light in air and the sine of the angle of refraction in the substance. It is easily determined on small samples and depends on the molecular structure, molecular weight, degree of unsaturation, temperature and the presence or absence of hydroxy-acid, conjugated bonds and free fatty acid, all contribute to the magnitude of its value (Fedeli and Jacini, 1971). Refractive index increases with molecular weight and has an approximately linear increasing relationship kith degree of unsaturation (F. A 0, 1997). Determination is done at 20% for liquid fats and 40•‹c for soiid fats. A correction factor can be included where it is not possible to work at the stipulated temperature. Refractive index = R + 0 Where R = refractometer reading Where 0 = correction factor Ash Content: The ash content is the residue, which remains after a weighed amount of fat is heated in a crucible to the ignition of the fat. The fat is left to burn, and the crucible is heated to a dull red heat until no more change is observed. The ash content is calculated from the intial weight of sample and the difference in weights of the empty crucible and the crucible together with the ash (Gordon, 1993). As oils and fats are organic substances, they should burn completely and leave no residues. Measurement of the inorganic residue after ignition is therefore a useful test for the determination of inorganic impurities (Gordon, 1993)

(b) Chemical Properties Acid Value: This is the number of milligrams of KOH required to neutralize lg of oil or fat. It indicates the amount of free fatty acids present {Ononogbu, 2002). The presence of free fatty acids in an oil or fat is an indicator of previous lipase activity, other hydrolytic action or oxidation (Gordon, 1993). Free fatty acids content varies with the source of the oil and level of refining. It can occur in refined oils at about 0.1% (wtw) up to as much as 15% in crude oil, but typically about 5% in crude oils (Hammond, 1993).

Iodine Value: Iodine value is the number of grams of iodine that combines with 'I009 of oil or fat. It gives the degree of unsaturation of the fat or oil (Ononogbu, 2002). This is based on the fact that halogen addition occurs at unsaturated bonds until these are completely saturated. Not all unsaturated bonds are alike in reactivity, and those near a carboxyl group hardly absorb iodine (Gordon, 1993). Such fatty acids are however rare. When the double bonds are conjugated, they react more slowly than non-conjugated double bonds (Gordon, 1993).

Several methods for determining iodine value are available. Those in common use being the tests of Wijs, Hanus and Rosenmundkuhnhem (Gordon, 1993). The differences between the methods stated are in the halogenating agents. Fats and oils can be classified by their iodine values. The iodine values of edible oils range from about 7 to over 200(Simpson and Corner- Orgarzally, 1986). Oils with value below 70 are usually referred to as fats because they are solid at room temperature. Another grouping which reflect their iodine value is: drying (higher than 150), semi-drying (between 100-150), non-drying (between 70-100) and fats (below 70) (Simpson and Corner- Orgarzally, 1986).

Saponification Value: This is the number of milligram of potassium hydroxide required to neutralize the fatty acids resulting from complex hydrolysis of lgof oil or fat. It is a measure of both free and combined acids. The esters of low molecular we~ghtfatty acids require more alkali for saponification, so that the saponification value is inversely proportional to the mean molecular weight of the fatty acids in the triacylglycerols present (Gordon, 1993). Because many oils have similar saponification values (Gordon, 19833, the test is not universally useful in establishing identity or indicating adulteration and should always be considered along with the iodine value for these purposes.

Peroxide Value: Peroxide value is the milliequivalent of peroxide oxygen per lOOg of fat. It is used to indicate the degree to which a fat has been oxidized (Ononogbu, 2002). Oxidation of an unsaturated oil or fat takes place via the formation of hydroperoxides. The hydroperoxides subsequently decompose into secondary oxidation products, the majority of which have unpleasant odour and flavours (Badifu and Abah, t998). Although hydroperaxides themselves have no off-flavours, they are an important aspect of rancidity development and are determined as the peroxide value. Peroxide value is usually less than 10 per gram of a fat sample when the sample is fresh (Gordon, 7993).

Unsaponifiable Matter: Unsaponifiable matter refer to the whole quantity of substances present in the oil or fat which after saponification by potassium hydroxide and extraction with a specified solvent, are not soluble in aqueous alkali and non-volatile under the condition of test.

The unsaponifiable matter of a fat includes, sterols, higher aliphatic alcohols, pigments, hydrocarbons as well as any foreign organic matter non-volatile at 100Oc (eg mineral oils) (Christie and Williams, 1982). Refined oils contain lower amounts of unsaponifiable matter than unrefined oils. Its d4ermination can be useful in indicating contamination and adulteration of the oil with a mineral oil or other non triglyceride contaminant (Young gt a., 1.2.6 Lipases Lipases are enzymes that catalyze the hydrolysis of stored triacylglycerols releasing fatty acids (Nelson and Cox, 2000). They belong to the class of serine hydrolases and therefore do not require any cofactor for activity, (E~gtved,1992). Under natural conditions lipases catalyses the hydrolysis of ester bonds at the interface between an insoluble substrate phase and the aqueous phase in which the enzyme is dissolved (Verger 1997, Schmid aJ., 1998). Under certain experimental conditions, such as in the absence of water, they are capable of reversing the reaction (Klibanov, 1997). The reverse reaction leads to the esterification and formation of glycerides from fatty acids and glycerol.

Lipases are widely distributed in animals, plants and micro-organisms, Among all these sources, it is the microbial lipases that find immense application. This is because microbes can be easily cultivated and their lipases can catalyse a wide variety of hydrolytic and synthetic reactions(Xu, 2000). Although pancreatic lipases have been traditionally used for various purposes, it is now well established that microbial lipases are preferred for commercial application due to their multi-fold properties, easy extraction procedure and unlimited supply (Macrae and Hammond1985).

Although widespread in nature, lipases have only recently become available in large quantities for industrial purposes (Xu, 2000). Some of the possible applications have been established within the last 15-20 years due to the discovery in which lipases could be used in microaqueous organic systems at high temperatures up to 70-80'~ (Zaks and Klibanov, 1984). The structural elucidation of lipase has since been investigated (Brady gJ a,,1990). Lipases tend to have a three dimensional structure. All lipases consist of a catalytic triad of His-Ser-Asp (Glu) (Saxena gt a,,1999). In addition, an oxyanion hole is formed, stabilizing the charge distribution and reducing the ground state energy of the tetrahedral intermediate (Derewenda, 1994). The catalytic triad of all lipases is buried under a "lid" of a surface loop, which undergoes a conformational change to open a channel for the active site to be accessible to substrates. This repositioning of the lid is caused by interfacial activation (Wong, 1995). Studies on lipase structure-function relationship have also been extensively carried out (Blow, 1991; Derewenda and Derewenda, 1991; Peters a aJ., 1996; Cygler and Schrag 1997; Svendsen gt d.,19%'; Zuegg @ d.,1997; Wei d.,1998). In nature, the lipases available from various.sources have considerable variation in their reaction specificities. From the fatty acid side, some lipases have affinity for short chain fatty acids (acetic, butyric, capric, caproic, caprylic, etc.). Some have preference for unsaturated fatty acid (oleic, linoleic, linolenic etc) while many others are non-specific and randomly split the fatty acids from traicylglycerol (~acrae,1983). From the glycerol side, the lipases often show positional specificity and attack the fatty acids at Sn-l or 3-carbon position of glycerol or at both positions' but not the fatty acid at the Sn-2 position of the glycerol molecule (Villeneuve a a,,1995).

(a) Microbial Lipases Microbial lipases particularly those of bacterial and fungal sources have been well studied (Sharma a aJ., 2001). However, the fungal lipases have been well studied compared to bacterial lipases. Lipases obtained from microorganisms have been shown to be commercially useful. They have been used to catalyse a number of reactions including esterification, transesterification, regioselective acylations of glycols and methols and synthesis of peptides (Chowdary gt d.',2001).

Bacterial Lipases: Bacterial lipases are glycoproteins, but some extracellular bacterial lipases are lipoproteins. Wrinkler et al., (1979) reported that enzyme production in most bacteria is affected by certain polysaccharides. Most bacterial lipases reported so far are constitutive and are non-specific in their substrate specificity and a few bacterial lipases are thermostable (Macrae and-Hammond ,1985).

Fungal Lipases: Fungal lipases are exploited due to their low cost of extraction, thermal stability, pH stability, substrate specificity and activity in organic solvents (Lawson et al., 1994). The chief producer of commercial lipases are: Aspergillus niger, Candida cylindracae,

Humicola lanuginosa, Mucor mehei, . Rhizopus arrhizus, R.delemar, R.japonicus, R.niveus and R.oryzea (Godfredson, 1990).

Among mucorales, the lipolytic enzymes of the moulds Mucor hiemalis, M.miehie, M.lipolyticus, M.pusillus, Rhizopus japonicus, R. arhizus, R.delemar, R.nigricans, R.nodosus, R.microsporus and R.chinesis, have been studied in great details (Lazar and Schroder, 1992). So far, lipozyme IM, in which Rhizomucor miehei lipase was immobilized on an exchange resin, has been the only commercial immobilized lipase for the production of specific structured triacylglycerols. Most studies,either screening or direct applrcations,have shown that lipozyme IM is the most optimal lipase in terms of reactivity and stability (Lee and Akoh, I996 Xu gt aJ., 1998a; Soumanou gt a,,1998;). The thermophilic Mucor pusillus, is well known as a producer of thermostable extracellular lipase. Stereospecificity studies on lipase have shown that lipases from Rhizopus species show maximum activity towards medium-chain fatty acids. Some have also been shown to have some degree or specificity towards Sn-2 or Sn-I and Sn-3 positions (Eigtved,1992; Villeneuve, 19%; Wong, 1995). Candida antarctica A lipase was specif~c for Sn-2 (Xu, 2000). The intracellular and extracellular lipases of Aspergillus niger are 1,3, -(regio) specific (Okumura gt aJ., 1976).

(b) Microbial Lipase Production Lipids are insoluble in water and need to be broken down extracellularly into their more polar components to facilitate absorption if they are to function as nutrients for the cell Therefore, majority of the lipases are secreted extracellularly [Saxena et a1.,1999) Environmental factors, such as temperature, pH, nitrogen, carbon and lipid sources, agitation and dissolved oxygen concentration have been shown to influence hpase production (Elibol and Ozer, 2001). Besides this, free fatty acids, hydrolysable esters, bile salts and glycerol also stimulate lipase production (Saxena gt d.,1999).

(c) Lipase Assay and Properties Lipases are known to hydrolyse triacylglycerols to form free fatty acids and glycerol. Therefore, the assay methods involve spectrophotometry, titrimetry, radiolabelling assay, fluorimetry, surface tension, and estimation of free fatty acids by high performance liquid chromatography (Brune and Gotz, 1992). Tributyrin plate assay and titrimetry (spectrophotometry) are the most commonly used methods for screening of lipase producers and estimation of Iipase activity respectively.

Lipase Purification: Lipases have been purified from fungal and bacterial sources by different methods involving ammonium sulphate prec~pitation, gel filteration and ion exchange chromatography. In recent years, affinity chromatographic technique is being used as this technique decreases the number of steps necessary for lipase purification as well as increases specificity. More currently, reversed mrcellar and two-phase systems, membrane processes and immuno-purification are belng used fot purification of lipases (Saxena gt &I.,1999). pH Optima: The lipases that have been studied show profound stability around pH 6.0-7.5 with considerable stability at acidic pH up to 4 and alkaline pH up to 8. Extracellular lipase of Aspergillus niger, Chromobacterium viscosum and Rhizopus Sp are active at acidic pH (Sharma & aJ., 2001). An alkaline lipase active at pH 11.0 has been isolated from Pseudomonas nitroducens (Watanabe gt a., 1977). A lipase from Penicillium expansum showed a maximum activity at pH of 9 and at 45'~and was stable at broad pH range of 6- 10 (Stocklein gt gl.,1993).

Temperature Optima and Thermal Stability: The pancreatic lipases lose activity on storage at temperatures above 40oC (Saxena gt aJ., 1999), but some microbial lipases of Aspergillus Niger, Rhizopus japonicus and R. viscosum are stable at 50•‹C (Sharma A,, 2001). Lipases of thermotolerant Humicola lanuginosa and P.nitroreducens are stable at 60'~and 70•‹c,respectively (Saxena gt aJ., 1999). Candida gigantea llpase had half-life for inactivation of 35.7, 46.4 and 22.9 minutes, respectively, at 45'~,50'~ and 55'~(Tombs, 1991). Purified lipase from Aspergillus terreus retained 100% of its activity at 60'~for 24 hr

Activation and lnactivation of Lipases: Cofactors are not required for the expression of lipase activity (Saxena a aJ., 1999). Divalent cations such as calcium generally stimulate their activity. Thls has been postulated as being based on the formation of calcium salts of long chain fatty acids (Macrae and Hammond, 1985., Godfredson, 1990). Several agents have been shown to activate or inactive the activity of lipase from different sources. The lipase activity, is inhibited drastically by co2', ~i~+,Flg2+,sn2+. In addition, 2n2', VIg2+, EDTA and SDS slighthly inhibited lipase activity (Saxena gj a,, 1999). In H. lanuginosa S-38, sulfhydryl reducing agents like dithiothreitol did not alter the lipase activity but did render it more susceptible to heat inactivation (Ghosh g aJ., 1996). Inactivation is accelerated by the addition of urea (Ghosh gj a.,1996). Reducing compounds (cysteine,2-mercaptoethanol), chelating agent (EDTA, 0-phenanthroline) and thiol group inhibitors (p-chloromercuric benzoate, monolodoacetate) did not show a detectable effect on lipase from Mucor pusillus, suggesting that lipase is not a metalloenzyme and it does not require either free -SH group or an intact S-S-bridge for its activity (Saxena gt aJ., 1999). (d) Substrate Specificity Specificity of lipases is controlled by the molecular properties of the enzyme, structure of the substrate and factors affecting binding of the enzymes to the substrate (Saxena a &I.,1999). Substrate specificity is often crucial to the application of lipases for analytical and industrial purposes. The microbial lipases can be placed in three groups accordmg to their specificity. The first group shows no marked specificity both as regards the position on the glycerol molecule which is attacked, and the nature of the fatty acid released. These lipases catalyse the complete breakdown of the triacylglycerols to glycerol and fatty acids together with diacylglycerols and monoacylglycerols as intermediates in the reaction. These intermediates do not accumulate since they are hydrolysed more rapidly than the triacylglycerols (Macrae, 1983). The second group of lipases catalyses the release of fatty acids regiospecifically from the outer 1 and 3 positions of acylglycerols (Xu, 2000). These lipases hydrolyse triacylglycerols to give free fatty acids 1,2,(2,3) -diacylglycerols and 2- rnonoacylglycerol ((Xu, 2000). Because 1,2,(2,3,) -diacylglycerols and especially 2-monoacylglycerol are chemically unstable and undergo acyl migration to give 1,3, diacylgiycerols and 1, (3) monoacylglycerol, respectively, prolonged incubation of a fat with a 1,3-specific lipase will give complete breakdown of some of the triacylglycerols with the formation of glycerol (Ghosh gt &,1996). Many extracellular microbial lipases such as those from Aspergillus niger show 1,3- (regio) specificity (Okumura gt aJ.,1976). The third group of lipases catalyses the specific release of a particular type of fatty acid from triacylglycerol molecules. Most lipases attack triacylglycerols as readily as partially esterified glycerides, but some have been shown to attack monoacylglycerol and diacylglycerols more rapidly than triacylglycerols and have been described as partial glycerol ester hydrolases (Saxena gt aJ., 1999). The table below shows the specificity of some lipases towards Sn-2 , Sn-I and Sn-3 positions and preference for long, medium and short chain fatty acids. Table 4 Some specific lipases for the production of specific structured triacylgycerols. Lipase source Fatty acid Regio specificity Sn specificity

Aspergillus niger S, M, L 1,3>>2 Candida lipolytica S, M, L 1,3>2 Humicola Ianuginose S, M, L 1,3>>2 Mucor javanicus M,L=+S 1,3>2 Rhizomucor miehei S > M, L 1 >3~>2 Pancreatic S>M,L 1,3 Rhizopus dekmar M,L>>S 1,3>>2 Rhizopus niveus M, L>S 1,3>2 Rhizopus arrhizus S,M>L I ,3.

Abbreviation: - L - long-chain fatty acids; S-short-chain fatty acids; M- medium chain fatty acids. Source: Xu (2000).

(e) tipases as lnter-esterification Catalysts The use of lipases as catalysts in organic synthesis is of much advantage. Beside their specificities, other obvious merits are efficacy of lipases under mild reaction conditions, utility in "natural" reaction systems and products, reduced environmental pollution, availability of lipases from a wide range of sources, ability to improve lipases by genetic engineering and in special situations the use of lipases for the prbduction of particular biomolecules (Xu 2000). The tolerance of lipases from aqueous buffer systems through biphasic emulsions and microemulsions, to organic solvent also offers much advantage. Lipase catalysed reactions are reversible (Macrae,1983). Hydrolysis and resynthesis of triacylglycerol therefore occur when lipases are incubated with oils and fats. This gives rise to inter-esterified products irnder conditions in which the amount of water in the reaction system is restricted (Klibanov, 1997). For this reaction, different oils and fats that contain different major fatty acids at the Sn-2 position of their triacylglycerols should be choosen (Xu, 2000). Also free fatty acids and ethyl ester could be used as acyl donors (Haling, 1987., Bloomer gt a.,1991; Quinlan and Moore, 1993; Lee and Parkin, l998., Pinsirodom & aJ, 1999). Immobifization of Lipase for Inter-esterification : The stability of lipases is one of the important characters for consideration in their industrial application. This is due to their high cost. Thus strategies for recycling of lipases used in inter-esterification are being developed. Enzyme immobilization have made this possible. After immobilization, lipases normally are more stable than their free states. Immobilized lipases can be used at high temperatures especially in microaqueous systems. The stabilization of lipases by protein/genetic engineering and enzyme modification has been studied (8allesteros & g.,1998).

Enzymes from different sources vary in their effectiveness with different immobilization techniques. Therefore there is need to match enzymes, support systems and substrates for optimal inter-esterification reaction rates (Bloomer gt a,,1990). Carrier choice 1s dependent on factors, such as, mechanical strength, microbial resistance, thermal stability, hydrophobic/hydrophilic characteristics, ease of regeneration, loading capacity and cast (Owusu-Ansah, 1994). immobilization strategy is dictated by factors such as enzymatic activity, effectiveness of utilization, toxicity of the immobilization reagents, deactivation and regeneration characteristics, cost of immobilization procedure and the desired final propertres of the immobilized lipases (Bailey and Ollls, ?986).The methods of adsorption, entrapmenlt and ion-exchange binding has found wide application for lipases used for inter-esterification reactions (Owusu-Ansah, 1994)

Inter-esterification Reaction Processes: Most studies for the systhesis and production of inter-esterified triacylglycerol products have used stirred tank reactors or simple glass vessels for the reactions (Schmid et al., 1998, Xu et al., 1998). However, some available reports show that packed-bed reactors are more promising for industrial developments with immobilized lipases (Jung and Bauer,1992; Mu gt a.,1998;and Xu gj d.,1999). Membrane technology has been introduced for the production of structured triacylglycerol [Basheer gt 4.) 1995; Balcao gt aJ., 1996 and 1998):Also Tubular-flow continuous reactors and continuous fluidized-bed reactor are more recent methods employed for industrial development(Chee gt 4.,2001 and Natalia 4.,2005) CHAPTER TWO MATERIALS AND METHOD

2.1 MATERIALS. 2.1 .1 Oil Seeds Fresh oil palm (Ellaies guineenses) fruits were purchased from Nsukka market. Fresh bush mango (I~ingiagabonensis) seeds were purchased from Abakaliki, Ebonyi State. The nut of shea butter tree (Butyrospennum paradoxumj were purchased from Markudi, Benue State. Fresh cocoa (Theobrorna cacao) pods were purchased from Aba, Abia State.

All the plant materials collected were identified and confirmed by Mr A Ozioko of Bio- resources Development and Conservation Programme (BDCP) Nsukka. They were then used for the subsequent experiment for preparation of fatsloils namely, palm oil, dika fat, shea butter and cocoa butter, respectively.

2.1.2 Chemicals All the chemicals used in this study were of the purest grade commercially available and were obtained variously from Merck, Germany, BDH Chemicals Ltd, Poole, England; May and Baker Ltd, England; Riedel-De Haen Ag Seilze-Hannover, Germany; Hopkin and Williams, Essex, England.

2.2 METHODS 2.2.1 Preparation of reagents The reagents used for enzyme analysis were prepared according to the methods described by Schmidt & a,,(1 974). 2.0% peptone 2.0g of peptone (oxoid) were dissolved in 100ml of deionized water. 0.5% yeast extract 0.59 of yeast extract (oxoid) was dissolved in 100ml of deionized water 0.5% NaCl 0.5g of NaCl was dissolved in 100ml of deionized water. 1.O% glucose 1.Og of glucose was dissolved in 100rnl of deionized water. Phenolphthalein indicator I.Og of phenolphthalein was dissolved in 100ml of ethanol. O.IN KOH 5.69 of KOH were dissolved in 100ml of deionized water. Is0/0 potassium lodide l5.Og of KI were dissolved in 100rnl of deionized water. 1.O% starch indicator 1.0g of starch was prepared with hot water into gel and made up to 100ml w~thde~onized water. 10.0% potassium lodide 10.09 of potassium lodide were dissolved in 100ml of deionozed water. Olive oil Suspension 5.09 olive oil + 5.09 gum Arabic was blended'with 95ml 0.89% NaCl solution in a mixer for 1Omins Deoxycho'late (IOmM) 0.4146g of deoxycholate was dissolved in 100ml,of deionized water Triethanolamine buffer (1M pH 8.5) 18.6g of triethnolamine hydrochloride was dissolved in 70ml distilled water and adjusted to pH 8.5 with 16ml of 5N NaOH, this was diluted to 100rnl with deionized water Copper reagent 18.6g of triethanolamine hydrochloride was dissolved in 70ml distilled water and 6.45g Cu(N03)2.3H20 was dissolved in 100ml of deionized water. Both solutions were mixed and adjusted to pH 7.5 with 18.5ml.5N NaOH and diluted to 2QOml with deionized water. Diethyldithiocarbarnate (IImM) 250mg of sodium diethyldithiocarbamate was dissolved in secondary butanol and made up to 100ml. Stearic acid standard solution (50uM) 142.25rng of stearic acid was dissolved in 100rnl chloroform and Iml of this solution was diluted with 100ml of chloroform. Incubation mixture 50 volumes of olive oil suspension was mixed with 5 volumes of deoxycholate solution and 45 volumes of triethanolamine buffer. 2.2.2 Extraction of oil from Samples Palm oil: 2kg portion of the palm fruits so purchased were pounded in a mortar to separate the pulp from the nuts. After separation, oil was extracted from the mashed pulp using 2 litres of chloroform and methanol, 2: 1 vlv.

Shea butter: The nuts of the shea butter tree were cracked to release the kernet. 3009 portion of the kernel was dried in the oven at 50'~for 2days. The dried kernel was milled and the oil extracted with 1.5 litres of petroleum ether (40-60•‹C) for 3hrs using a Soxhlet apparatus.

Dika fat: The fleshy parts of the bush mangoes were cut off using knife to reveal the nut. The nuts were then cracked using stones to release the seeds. A 400g portion of the seeds was dried in the oven at 50'~for 2days. The dried seeds were milled and the oil extracted with 1.5 litres of petroleum ether (40-60•‹C)for 3hrs using a soxhlet apparatus.

The oil samples obtained were separated from the extracting solvents by distillation at 5OoC. The samples were then stored in the refrigerator until required for analysis.

2.2.3 Partial Refining of the Oil Samples The oil samples were partially refined by bleaching according to the method used by P.0.S pilot plant,Saskatoon, Canada as outlined below. Filtration: The oil samples were filtered by passing them through a Whatman no 4 filter paper immersed in a big funnel and kept inside the oven.The oven temperature for filtration of palm oil extract was 40'~while shea butter and dikafat where each filtered at oven temperature of 50'~. Bleaching: 459 of Fuller's earth were transferred into 3009 of the filtered oil inside a round bottom flask. The mixture was thoroughly shaken manually and the flask was set up on an aluminium plate containing paraffin (oil bath} and which, in turn, was placed on a hot plate. The flask was connected to a vaccum pump through a trap placed on ice blocks. The temperature of the paraffin was raised and maintained at 80•‹c for 30 minutes with the flask being shaken manually at intervals of 10 minutes. The flask was allowed to cool for 10 minutes and the mixture was filtered through a Buchner funnel whose sides were treated with grease before the Whatman filter paper was inserted. 2.2.4 Determination of % Oil Yield. The crude oil yield was determined by the IUPAC method (1979). A (log) portion of the dried fatty material was extracted using a soxhlet extractor for a period of six hours with petroleum ether (40--60'~) and allowed to stay for a period of 16hr. The solvent was then evaporated and the residual oil was dried to a constant weight in an oven at 50•‹C.The oillfat content was expressed as the percentage of the fatty material.

2.2.5 Solvent Fractionation (Crystallization) Fractionation of the partially refined oil samples was carried out using n-hexane as described by Gunstone gt a.,(1994).

A 50g alliquot of the oil sample was dissolved in n-hexane. The solution was slowly reduced to its crystallizing temperature using LKB(BROMMA)7000 ULTRORAC FRACTION COLLECTOR made in Germany, as determined through preliminary tests. Crystallization was allowed to continue for 16hr and the olein-stearin fractions were separated through filtration at the same temperature conditions: palm oil (1 -- ~OC),shea butter (6 --lZ•‹Cj, and dika fat (18-- 24%). The solvent volume was varied while the amount of oil was kept constant for each temperature under study to obtain different olein-stearin fractions. The results were expressed as a percentage of the quantity of oil used.

2.2.6 Characterization of the Partially Refined Oil Samples Characterization was carried out to assess the quality of the oil extracts, which are measurable by suitable physical and chemical techniques.

Colour and Odour The colour and odour of the samples were physically evaluated by visual inspection and smelling.

Specific GravitylDensity: A 50ml specific gravity bottle was first washed with a detergent (Omo), rinsed with water and then petroleum ether. The bottle was dried in the oven at 50•‹c, allowed to cool and weighed empty. The bottle was then filled with distilled water and the weight of the bottle and water noted. The bottle was kept in the oven to dry.After the bottle was cooled, the oil sample to be analyzed was used to fill the bottle and the weight also rioted. The specific gravity of the oil was calculated using the formula below Specific gravity - weiqht of xcm3 of oil Weight of Xcm3 of water Where X = Volume af sample rased in cm3

Refractive Index: For this measurement, the Abbe refractometer (Bellingham +Stanley Ltd No A82316 made in England) was used. The instrument was reset before use, using a light compensator (water at about 25'~).A smear of the sample was then placed on the lower prism of the instrument and after closing, light was passed through by means of an angled mirror. The reflected light appeared in form of a dark background. With five adjustments the telescope tubes were moved until the black'shadow appeared central in the cross wire- indicator. The refractive index was read off. The temperature of measurement was at 30%

Viscosity: This measurement was conducted at 45'~using universal torsion viscometer (Gallenkamp Vs-010 and Vs-020 made in England). The viscometer cylinder was simply immersed into oil contained in a cup. The pointer was released and its position on the calibrated dial was noted. This was repeated to get a second reading. The average was used to calculate viscosity using the viscosity graph.

Melting PointlSlip Point: 10ml portion of oil was drawn into a thin wall capillary tube, Imm in diameter. One side of the tube with the oil was sealed in a small flame without burning the oil and the tubes were stored overnight in..a refrigerator. The tube was attached to a thermometer, suspended in a water bath, and heated slowly while the water was being stirred manually and gently. The melting point was taken as the temperature at which the oil becomes completely transparent and liquid when observed through with a magnifying glass.

Acid Value: A quantity (20ml) of ethanol (95%) was mixed with 20ml of diethyl ether, Iml of phenolphthalein indicator (1% in ethanol) was added. The mixture was neutralized by adding with 0.1N KOH from a burette and warmed. A 50ml quantity of this solution was added to 29 of the oil weighed into a 250ml conical flask. The mixture was boiled for about 5 minutes and titrated while hot with a standard aqueous solution (standardized by titration with standard HCI solution) of potassium hydroxide while shaking vigorously during the titration until a pink colour, which persisted for 15seconds appeared. Acid value = 56.1 xVxN W Where V = volume of KOH N = normality of KOH W = weight of oil sample.

Iodine Value: The Wijs method as described by Hendriske and Hawood (1986) was used. The Wijs reagent (iodine trichloride solution) was prepared by dissolving 29 of iodine trichloride in 50ml of glacial acetic acid and mixing with 2.25g of iodine dissolved in 100rnl glacial acetic acid. The mixture was then made up to 250ml with glacial acetic acid, stored in a brown glass bottle, and kept out of light until use. A 0.5g portion of the oil was weighed and transferred into a 250ml glass-stoppered bottle. Then 15ml of Wijs solution was addedto dissolve the oil and a further 25ml of Wijs solution was added from a burette. The flask was closed and the content mixed by manually shaking the flask. This was allowed to stand in the dark for 30 minutes. After standing, 20ml of 15% potassium iodide solution was added and the bottle stoppered and shaken thoroughly and the sides of the bottle and the slopper were washed with 100ml of recently boiled and cooled water. Then the solution was titrated with a standard O.1N sodium thiosulphate solution, the reagent being added with constant shaking until the yellow colour of the iodine has almost disappeared. Before continuing the titration, 2ml of 1% starch indicator was added. When the blue colour had almost disappeared, the bottle was stoppered and shaken vigorously for the remaining iodine in the organic layer to pass into the water layer. Two blank determinations with the same quantities of reagents were carried out at the same time and under the same conditions.

Iodine value = 12.69 x M (v-v') m Where M = molarity of sodium thiosulphate m = mass of oil in grams V = volume of sodium thiosulphate used for the blank v1 = volume of sodium thiosulphate used for the sample

Peroxide Value: The Wheeler method as described by Hamilton gt a.,(1992) was used. A 1.Og portion of the oil was dissolved in 25ml of a solvent mixture consisting of 60% glacial acetic acid and 40% of chloroform and lml of 10% saturated solution of potassium iodide was added. The flask was shaken and allowed to stand in the dark for 5 minutes and 75ml of distilled water added. The mixture was then titrated with 0.1N standard solution of sodium thiosulphate using 2ml of 1% starch solution as indicator. A blank determination was carried out at the same time. Peroxide value = (V, - Vb) N x 100 W Where V, =volume of sodium thiosulphate used in test sample (ml) Vb = volume of sodium thiosulphate used in blank (ml) N = the normality of sodium thiosulphate W = mass of sample used in grams.

SAPONIFICATION VALUE: the method described by Njoku @ gl.,2000 was used as follows: A 2.0g portion of Ithe oil was weighed into a 250ml conical flask. 25ml of 0.5M alcoholic caustic potash solution was added. The flask was connected to a reflux condenser and the mixture heated on a hot plate for 1 hour (by which time the sample was completley saponified as indicated by the absence of any oil matter and apperance fo clear solution).

After the flask and the condenser were cooled, the inside of the condenser was washed down with 1Om1 of hot ethanol neutral to phenolphthalein. Iml of phenolphthalein indicator solution was added and solution titrated with 0.5M standard hydrochloric acid. A blank determination using the same quantify of potassium hydroxide was carried out at the same time.

Saponification Value = 56.1 x (V - v') M W Where M = molarity of potassium hydroxide v - titre volume of the blank v1 - titre volume of the test sample W = mass of oil used in grams.

Unsaponifiable Matter: The method described by Njoku gt aJ.,(2000) was used. After the titration for the saponification value, the neutralised liquid was made alkaline with Iml of aqueous 3N potassium hydroxide solution, This was then transferred to a separatory funnel and washed in water. While still warm, the solution was extracted three times each with 50ml of diethyl ether. Each ether extract was poured into another separatory funnel containing 20mI of water. The combined ether extract was shaken first with 20ml of distilled water and then vigorously twice with 20ml of distilled water.

The ether extract was further washed twice with 20ml of aqueous O.5N potassium hydroxide solution and with 20ml of distilled water until washings were no longer alkaline to phenolphthalein. The ether extract was then poured into a weighed flask and evaporated to dryness at 80'~. Unsaponifiable matter = 1OOml/m WhereM, = mass (in grams) of the residence M = mass (in grams) of oil.

2.2.7 Production and Extraction of Fungal Lipase Isolation of fungus: Pure isorates of Aspergillus niger used for this research was collected from the Mycology Laboratory of the Department of Microbiology, University of Nigeria, Nsukka (identity confirmed by Mr. C. U Anyanwu of the same department).

The pure isolates were reactivated using Sarbroud Dextrose Agar(S.D.A) medium. Culture plates were sterilized together with the S.D.A medium in an autoclave for 15min at 12I0C and I5psi (Omar et al., 1987). The sterilized medium was poured into the sterilized plates and was left overnight to gel properly. The plates containing the gelled medium were then inoculated with pure Aspergillus niger strain. This was incubated at room temperature for 3 days for the organism to grow.

Production of fipase: Lipase production was done according to the method of Chandler et al.,(1980). The reactivated Aspergillus niger was inoculated rnto 50ml af the sterilized growth media (pH 7.0) composed of 2% peptone (oxoid), 0.5% yeast extract (oxoid), 0.5% NaCI, 1% glucose and 0.1% olive oil .The incubation was done at room temperature for 72 hr for the production of lipase. Enzyme Extraction: Lipase extraction was done according to the method described by Omar gt aJ., (1987). The medium used for lipase production was centrifuged for 30 minutes at 5,000xg and 10'~using refrigerated centrifuge IEC B-20A centrifuge (DAMON - England), to separate the mycelial growth. The cell-free extract was the source of lipase enzyme. To 50ml portion of this extract was added 40mI of cold acetone. After standing overnight in a refrigerator, the precipitate formed was collected by centrifugation for 30 minutes at 10,000xg and 10'~. The resulting precipitate was washed twice with cold acetone and then recentrifuged for IOmin at 10,000xg and 10'~.The final precipitate was dissolved in 0.5 M sodium phosphate buffer (pH 7.0) and the undissolved materials were removed by centrifugation at 10,000~g and 10'~for 1Omin.

2.2.8 Chacterization of Lipase Most commercial applications of lipases do not require highly pure enzyme (Sharma aJ., 2001) Excessive purification is expensive ad reduces overall recovery of the enzyme (Chisti, 1998) therefore, partial purification of the Aspergilus niger enzyme by acetone precipitation could be considered adequate for its use as interesterification catalyst in this study. The loading capacity of the support is an impoprtant factor to be considered in the use of enzyme for intersterrification (Owusu - Ansah, 1994). In order to match enxyme and support system for optimal interesterification reaction, the study of the effect of quantity of support on the activity of the immoblized enzyme wascarried out.

The lipase produced was partially characterized for pH, temperature and substrate concentration as follows: T

Temperature optimum:The optimum temperature for activity was determined by incubating the enzyme and substrate in a shaking water bath which was maintained at different temperature levels of 25'~-85'~.~hevolume of enzyme and substrate concentration used are as described in assay for activity below. pH 0ptimum:The optimum pH for activity was determined by preparing buffers of different pH levels as described by Rassadin (1989).The buffers prepared are Phosphate-citrate buffer(O.lM,pH 5.0-8.0), Tris-buffer (O.lM,pH 7.1-9.2) and Glycine buffer(0.2M ,pH3.6-5.8). Assay of lipase activity as described below was done using the various pH ranges as the suspending medium. Substrate optimum: Different substrate (olive oil) concentrations were prepared ranging from O.6mM-80mM.The individual substrate concentrations were used to prepare incubation mixtures, which were treated according .to the method used for the assay for activity,described below:

Assay for Activity: This was done according to the method described by Schmidt g d., (1974). A 1.0ml aliquot of the incubation mixture was added to a O.Iml sample of the enzyme in a centrifuge tube. The tube was stoppered and incubated in a shaking water bath (30'~). After exactly IOmin, the tube was immersed in a water bath at 80% for I min to stop the reaction. 50mI chloroform and 25ml of copper reagent were added to the mixture, then stoppered and shaken for 20 minutes in a shaker. The tubes were centrifuged for Sminutes at 5,000xg and 10'~.The aqueous phase was sucked off with a polyvinyl chloride capillary. The chloroform layer was gently pipetted into clean tubes. The colour was developed with solution V and the extinction was measured at 440nm on a Unicam sp 500 spectrophotometer. Volume activity was calculated with the formula.

Volume activity = E li~a,, x 250 x F

WhereV = sample volume F = sample diffusion factor (where necessary) E - enzyme

2.2.9 Immobilization of the Lipase

The enzyme was immobilized on silica gel type 60 for TLC, by adsorption as described by Macrae (1983).

A 0.59 portion of the inorganic (Silica gel type 60,lO-40uM pore size.E.Merck,Darmstadt,Germany) support was poured 5mls of deionized water to make slurry. This was put in the refrigerator and allowed to stay overnight. The slurry was then introduced into a 2.5ml buffered solution (pH 7.2) of the lipase. To this was poured cold acetone to a final concentration of 80% vlv. The mixture was kept in the refrigerator and allowed to stay overnight. The precipitated enzyme coats the inorganic particles. The lipase coated-particles were collected by filtration, dried at room temperature and then used for the study. Determination of Loading Capacity of the Support System: To different quantities (0.2- 2.09) of the support slurry was poured 2.5ml of the enzyme solution. Immobilization was completed as described above. The activity for each was determined as previously described.

2.2.10 Inter-esterfication Reaction System.Inter-esterification was done according to the method of Macrae (1983)

The inter-esterfication reaction was carried out in a batch sysfem. The oil fractions were mixed in a 1:j ratio with 39 of the immobilized iipase. The mixture was stirred at ?OOrpm at a temperature of 35'~for 3 days using Gallenkamp orbital incubator,England. The solvent medium was n-hexane. After the reaction, the enzyme was removed by filtration through Whatman No. 4 filter paper. The filtrate was washed several times with water. The organic layer was dried over anhydrous Na2S04 and then the solvent evaporated under reduced pressure using Sargent-Welch Vacuum Pump.no.8805,serial-5212. 7300 North Linder Avenue,Skokie ILLINOISl60026.U.S.A. The product obtained was taken to be the inter- esterified product

2.2.1 1 Fatty Acid Analysis of Hydrolysates.

Hydrolysis of the Products: The fatty acids in the selected inter-esterified products were released by ester hydroysis as described by Badifu and Abah (1998). A known quantity (29) of the inter-esterfied product was weighed into a 250ml round bottom flask. To this was added 20ml of the methylating reagent comprising methanol, benzene and sulphuric acid (20:10:lv/v). The mixture was refluxed for about 90 minutes. The solution was transferred into a 250ml separatory funnel and 25ml of n-hexane was added followed by 25ml of saturated sodium chloride solution. The mixture was shaken vigorously. The lower layer was drawn off while n-hexane layer was then filtered through Whatman No. 1 filter paper into a 50ml beaker. The solution was reduced on a steam bath in a fume cupboard to a final volume of about 5ml.

The fatty acids were liberated from the methyl-esters by dropping 20ml of 10% dilute sulphuric acid into the hydrolysates (El-Zanati and Khedr 1991). The released fatty acids were then extracted with 20ml of n-hexane, washed several times with deionized water and dried over anhydrous Na2S04. The samples were stored in bijour bottles until required.

Urea Fractionationllnclvsion: Urea was incorporated into the fatty acids on the basis that linear fatty acids form inclusion compounds with urea. The method is that of Schlenk (1954). Iml portion of each fatty acid sample stored in bijour bottle was measured into a 50ml beaker. To this was added 5mI of hot methanollurea mixture (30:5 mllg). The solution was allowed to cool to room temperature at which urea inclusion compounds crystallized readily. This was done to the standards and the samples to be analysed.

TLC Studies: Glass plates 20x 20 cm were coated with 0.5mm of slurry of 509 silica gel G in 100ml of distilled water using a Shandon spreader. 0.16% Na2C03was incorporated into the silica gel. The plates were allowed to dry at room temperature for 2 hours and then activated in the oven at 110'~for 2 hours.

The urea inclusion compounds of the standards and samples were spotted at a distance of 1.5cm from the lower edge of the plate with a micropipette, dried in a cold air stream and placed in a Shandon TLC tank saturated with a solvent system of hexane- diethyl ether - acetic acid (80:20:1 vlvlv). The plates were allowed to develop by ascending technique for Ihr 30min. The plates were dried at room temperature and components separated were visualized by iodine vapour (Christie and Williams, 1982). After the spots corresponding to the fatty acid standards were identified, they were circled. The circled portions were scrapped into test tubes. 5ml of chloroform was added to each test tube, vigorously shaken and filtered through whatman filter paper No 4. The filtrate was evaporated to dryness at 40' C under reduced pressure using Sargent - Nelch Vacuum Pump Model = No 8805, serial, 5212, 7300 North linder avanue, skokie Illinois, 60026, USA. To each of the test tubes were added 5ml of deionized water to split the urea inclusion compounds and release the fatty acids. The liberated fatty acids were extracted from the aqueous phase with an extracting mixture of 15ml chloroform and methanol (21 vh). After shaking thoroughly, the mixture was poured into a separatory funnel and allowed to stand for 10min. The chloroform layer was dried over anhydrous Na2S04.This'was further treated according to Ducombe's modification of the method for the determination of long chain fatty acids (Ducombe, 1977). The method is based on the formation of copper complex of fatty acids which is then measured calorimetrically. The solution above was partially evaporated to about 5ml quantity at 40' C in a water bath, after which lOml of phosphate buffer (pH 7.0, O.-lM) was added. The mixture was shaken for about 90 seconds using a Gallenkamp test tube shaker .The tubes were allowed to stand for 15min at room temperature. The lower aqueous layer was removed using polyvinyl chloride capillary. The upper chloroform layer was transferred into another dry clean test tube and 5ml of copper triethanolamine solution was added. The tube was shaken thoroughly using a shaker KARL KOLB VORTEX-GENIE, West Germany, for IOmin and allowed to stand for 15min. The lower layer was removed with a pipette and the chloroform layer filtered with Whatman No 4 filter paper into dry clean test tubes. Two drops of sodium diethyldithiocarbamate was added and the absorbance of the resultant colour read at 440nm against a blank in a UNICAM series-SP 500 spectrophotorneter.

2.21 2 Statistical analysis The data were analysed using the Student's t-test and two-way ANOVA at 5% level of significance. Values reported are averages only. CHAPTER THREE

RESULTS The results obtained in this study which are shown in this chapter include, the results of crude oil yield, fractionation studies of the partially refined (PR) oil samples, physical and chemical characterization of the PR oil samples and their fractions, partial characterization of the Aspergillus niger lipase and characterization of the inter-esterified oil samples.

3.1 PHYSICAL PROPERTIES OF CRUDE AND PARTIALLY REFINED OIL SAMPLES

3.1 .I The yield of crude oil from the different oil fruitslseeds

The oil yield in percentage and in grammes per kilogramme is shown in table 3.1 below. The result shows that dika fat had the highest oil yield, while cocoa butter had the least oil yield. The yield of palm oil, dika fat and shea butter from their oil bearing materials, was found to exceed 50% but that of cocoa butter did not: The oil yield in gram per kilogram of the oil seeds shows that dika seeds produced more oil than all the other samples while palm fruits had the lowest production of oil. The yield of palm oil, shea butter and dika fat was found to vary significantly from the yield of cocoa butter (pe0.05).

Table 3.1: the yield of oil from the different oil fruitlseeds

Yields Oil or fat Significant glkg oil seed % level Palm oil 220 k 21.2 56.9sfi 0.28~ PeO.05

Shea butter 513f 16.4 . 51.3k 1.56b Pe0.05 Dika fat 684 + 23.1 68.45 f 1.48b Pe0.05

Cocoa butter 442 f 20.2 44.2-L 0.28a

Tab~~latedvalues are means $' SE (N=3) Means having the same superscript as cocoabu!ter are no1 significantly different ( P >O 05) 3.1.2 Acylglycerol fractions of the partially refined (PR) oil extracts. The result of the acylglycerol fractionation of the PR oils are summarized on tables 3.2, 3.3 and 3.4, respectively

Table 3.2 Acylglycerol fraction of Palm Oil at varying temperatures and solvent concentrations. Table 3.2a % Acvlqlycerol fraction at Oil /solvent 1:1 1: 2 1:3 1 :4 15

Palm Olein 19..975 4 1 .38a 23.775 f 1.38" 39..500i 1.38" 5 1 .025f 1.38' 59.450+ 1.38"

Palm Stearin 80.025 1 1.37-7.225 ? 1.37" 60.500 f 1 .37b 48.975 + I .37" 40.600 k 1 .37d

Tabulated values are Means f SE (N = 3) Valucs with tlu same superscript arc not signiliuantly d~lli-rentat (P>0.05)whereas those with direrent superscript are significantly different (p>0.05)

Tahle3.2h

'XI Acylglycerol fraction at

loc 3"~ jOc 7Oc

Palm Olein 28.26 + 1 ,73a 34.021t: 1 .73b 42.52k 1.73' 50.18+ 1 .73d

Palm Stearin 7 1 ~4 + 1.22a

Tabulated values are Means f SE (N = 3)

Values w~ththe same superscript nre not signiticantl!, different at (1'>0.03) whereas those with diffeent superscript are signiticantlv different (P

Table 3.2a and 3.2b shows the means of Olein and Stearin fractions of palm oil at varying

temperatures and solvent concentrations. The result shows that as the temperature was illcreased from

1•‹cto 7' C, the olein fractions increased while the stearin fractions decreased. Also as the solvent

ratio was increased, there was an increase in otein production and a decrease in stearin production. The

highest stearin fraction of about 88.2% and lowest olein yield of 1 1.8% were obtained at lo C and I : I oiI/solvent, while the highest olein fraction of about 70% and lowesi stearin fraction of 29.8% were obtained at 7' C and 1:5 oil/solvent. The nearest equilibrium fraction of olein and stearin was observed at lo C and 1 :5 oil/solvet~t.Statistical analysis showed a correlation between the temperattlre and solvent changes. This indicates that the differences in olein- stearin separation are statistically significant (P<0.05).

Table 3.3 Acylglycerol fraction of Shea Butter at varying temperature and solvent concentrations

Table 3.3a % Acylglycerol fraction at

Oil /Solvent I:I 112 1:3 114

Shea Stearin 94.075 f 4.7Za 85. I25 + 4.7Zab 71.475+ 4.7zbC 64.75 + 4.72' 61.425 + 4.72'

Tabulatcd values are Means k SE (N = 3)

Values with the same supcrscrip~are not significantly different at (P>O.Oj) whereas thosc with different strpencript are significantly differen1 (P<0.05).

Table 3.3b % Acylglycerol fraction at

6'~ 8Oc l OOC 12Y Shea olein 6:46 + 4.3fja l5.16+ 4.36" 33.40+ 4.36' 43.10~4.36"

Shea stearin 93.54 f 4.22" 84.84 f 4.22" 66.80 f 4.22~~56.30 .' 4.22'

Tabulated values are h,leans k SF. (N = 3) Values w~rhthe same supzrscrlpt are not significantly difrerenr at (PX.05)whereas thosc with direrent superscript arc significantly d~n'crcnt(P;O.Oj)

Table 3.3a and 3.3b shows the means of Olein and Stearin fractions of shea butter at varying temperatures and solvent concentrations. For shea butter, increasing temperature from 6% to 12'~showed a progressive increase in olein fraction and decrease in stearin fraction. Increase in sovent ratio also produced an increase in otein production but a decrease in stearin production. The highest shea butter stearin fraction of 98.7% and lowest olein fraction of 1.3% were obtained at 6'~and 1:l oil/solvent. The highest sheabutter olein fraction (67.5%) and lowest stearin fraction (32.5) were both obtained at 12'~and 1:5 oillsolvent. At 12'~and 1:3 oil/solvent, approximately 50% and 50% stearin was produced. Statistical analysis showed a correlation between the temperature abd solvent changes. This indicates that the differences in olien- stearin separation are statistically significant (P<0.05).

Table 3.4 Acylglycerol fraction of Dika Fat at varying temperatures and solvent concentrations.

Table 3.4a % Acylglycerol fraction at

Oil/ solvent 1:l 1 :2 1 :3 1:4, 1:5

Dika olein 2.700k 3.28" 5.925 + 3.2ga 18.675 + 3.2gb 25.95 rt 3.2~~43.45 + 3.28'

Dika Stearin 97.300k 3.28" 94.075 k 3.28" 81.325-t 3.2~~74.050-t 3.2gb 56.550+ 3.28'

Tabulated \aloes arc hdcans k SE (N = 3)

Valucs ~\~ththe same superscript are not signilicanlly d~fferrnra1 (I'N.115) \\hereas those wthdilr'crent superscript are signiticant!y ditrerent (P

Table 3.4b % Acylglycerol fraction at 1 20•‹c 22Oc 24"~ Dik olein 7.96 f 2.93" 15.54 2 2.~)3~" 23.38 t-~.93~ 30.48f 2.93'

Dika stearin 92.04 f 2.93-4.46 + 2,Nab 76.62f 2.93b" 69.52 + 2.93'

Tabulated values are Means k SE 1N = 3).

Values with the same superscript are not sign~licantlydifferent at (P>OOS) n41eri.a~tlwsc with different sctperscript are significantly difkrent (P

Table 3.4a and 3.4b shows the means of Olein and Stearin fractions of dika fat at varying temperatures and solvent concentrations. The result shows that as the temperature is increased (from 18'~to 24'~) different proportions of olein-stearin separation were achieved. Also increase in solvent volume produced more increase in olein-stearin separation at all the temperature levels studied.The highest stearin (99%) and lowest olein (1%) fractions were obtained at 18'~1:l oil/solvent, while the highest olein (59.5%) and lowest stearin (40.5) was produced at 24'~and 1:5 oil/solvent. The closest equilibrium fractions of 43.6% olein and 56.4% stearin were obtained at 24% and 1:4 oil/solvent. Statistical analysis showed a correlation between the temperature and solvent changes. This indicates that the differences in olien- stearin separation are statistically significant (P<0.05).

3.1.3: Physical properties of partially refined (PR) oils and their fraction. The results of the PR oil samples and their fractions are summarized on tables 3.5, 3.6, 3.7 and 3.8, respectively.

Table 3.5: Physical properties of PR palm oil and it's fraction Property Palm oil Palm olein Palm midfraction Palm stearin Physical state Semi liquid Liquid Liquid Solid

Colour Reddish Yellow yellow Yellow Odour Indistinguishable Indistinguishable Indistinguishable Indistinguishable Specific gravity 0.909-t 0.001 a 0.915 -t 0.003b 0.889-t 0.001b 0.897 -t 0.001b Melting point 24-30 <20 25-28 34-38 Refractive index 1.465 + 0.015a 1.468 + 0.006a 1A67 + 0.004a 1.460-t 0.014b Viscosity (cp) 45' 149f 1.41a 144f 0.41b 154kO.lb 158k2.12b

T;tbulakd values are Mcaris 2 SD (N = 3)

Means having the same letter with palm oil along the same column are not significantly different (P>0.05) whereas those having letter from palm oil are significantly different (Pc0.05)

Table 3.5 above shows the physical properties of partially refined palm oil. As can be seen from table, bleaching changed the colour of crude palm oil from red to yellow. The olein fraction was found to have the highest specific gravity and refractive index. Melting point of palm oil increased in the stearin fraction but decreased in olein and palm mid fraction (PMF). Palm olein could not crystallaize under the available means of temperature analysis. Palm stearin had the highest viscosity than all the other fractions. The specific gravity of the partially refined (PR) palm oil differs significantly (Pc0.05) from those of all its fractions. The viscosity of PR palm oil sample differed significantly (P0.05) from those of its fractions. The refractive index of PR palm oil did not differ significantly (P>0.05) with those of its olein and mid fraction but did differ with that of its stearin fraction. 45

Table 3.6: Physical properties of (PR) shea butter and its fractions

Property Shea butter Shea olein Shea stearin Physical state at 30•‹c Solid Semi-liquid Solid Colour Milky yellow Bright yellow Milky yellow Odour Indistinguishable Indistinguishable Indistinguishable Specific gravity 0.923 + 0.006a 0.92950.013b 0.906-t-0.01 1 b Melting point 34 - 38 28-32 38-40 Refractive index 1A67 + 0.008a 1.473f0.008b 1.46350.007a Viscosify (cp) 45' 7 52 + 2.83a 150+O,la 154 + 1.49a

Tabulated values are Means k SD (N = 3) Means having the same letter with shea butter along the same colwnn are not significantly different (P>0.05}whereas lhose having letter from shea httetl are significantly different (Pc0.05)

Table 3.6 above shows the physical properties of PR shea butter and its fractions.The colour of shea butter changed from milky yellow to a bright yellow olein and milky yellow stearin when fractionated. The melting point range of PR shea butter decreased remarkably in the olein but increased slightly In the stearin fraction, The olein fraction of PR shea butter had higher specific gravity and refractive index than the PR shea butter and its stearin fraction. The stearin fraction had the highest viscosity of all the other samples. The refractive index (RI) of PR shea butter differed significantly (Pc0.05) with the RI of its olein fraction but did not differ with that of its stearin fraction. The specific gravity of PR shea butter differed significantly (P4.05) with those of its olein and stearin fractions. Table 3.7 Physical properties partially refined (PR) dika fat and its fractions.

Property Dika fat Dika olein Dika stearin Physical state at 30•‹c Solid Semi-liquid Solid Colour Yellow Bright yellow Yellow Odour lndlstinguishable Indistinguishable Indistinguishable Specific gravity 0.901 + 0.015a 0.913 + 0.808b Q.892k0.01 3b Melting point 42-45 35-38 42-46 Refractive index 1.477 + . 003a 1.477 + 0.001a 1.478+ 0.001a \fiscosity (cp) 45Oc 170rtr 2.83a 160-114.24b 170f 0.la

Tabulated values are Means k SD (N = 3) Means having the same letter with dika fat along the same column are not significantly different (P>0.05) whereas those having letter from dika fat are significantly different (Pc0.05)

Table 3.7 above shows the physical properties of PR dika fat and its fractions. The result shows that with dika fat, fractionation did not produce any appreciable change in colour. The olein fraction has the highest specific gravity and lowest viscosity. The melting range of PR dika fat decreased remarkably in the olein but did not differ with its stearin fraction. The specific gravity of the PR dika fat and its olein fraction did not differ.But specific gravity of PR dika fat differed significantly (P4J.05) with the specific gravity of its stearin fraction. The refractive index of the dika fat did not differ significantly (P>0.05) with those of its fractions. The viscosity of PR dika fat did not differ with that of its stearin fraction but differed significantly(P<0.05) with that of its olein fraction. Table 3.8: Physical properties of partially refined (PR) Cocoa butter.

Property PR Cocoa butter Physical State at 30•‹c Solid Colour Light yelfow Odour Indistinguishable Specific gravity 0.921 + 0.01 1 Melting point 30-34Oc Refractive Index 1.46305 0.003 Viscosity 150 f2.83

Tabulated values are Means k SD (N = 3)

Table 3.8 above shows the physical properties of PR cocoa butter. Here, the values of whole cocoa butter is given and was not fractionated. This is because; the study aims to compare the extent of resemblance of oils from other sources to whole cocoa butter and not its fractions. Therefore, the values of fractions from cocoa butter are not required for this study. From the results above, it shows that the specif~cgravity of cocoa butter did not differ significantly with those of PR shea butter, dika olein and palm olein, but differed significantly (P<0.05) from those of PR dika fat, PR palm.oil, palm stearin, palm mid fraction and shea butter olein. Also the refractive index of PR cocoa butter did not differ significantly (P405) with those of PR palm oil, PR shea butter, palm stearin and shea butter stearin but differed significantly (P4.05) with the refractive index of PR dika fat, palm olein, shea butter olein and dika fat olein. It can also be seen that the viscosity of PR cocoa butter did not vary significantly with those of PR palm oil, PR shea butter, shea butter olein, shea butter stearin, but did differ significantly (Pe0.05) with the viscosity of dika olein, palm olein, palm stearin, palm mid fraction dika steain, PR dika fat.

3.2 Chemical properties of partially refined oil extracts and their fraction

The result of the study of the chemical properties 0' the partially refined extracts of palm oil, shea butter, dika fat and cocoa butter sampleg and their respective fractions as determined by various parameters are sctmmarized on Tables 3.9-3.12 respectively Table 3.9: Chemical property of partially refined (PR) palm oil and its fractions

Paremeter Palm oil Palm olein Palm mid Palm stearin fraction Acid value 21.5+ 1.13a 23.9+ 0.99b 18.2-t.0.42b 15.65 0.71b Peroxide value 0.15 + 0.02a 3.0+ 0.28b 0.050.001b 0.01+ 0.001 b Iodine value 54.1 + 0.71a 60.6+0.42b 50.8k 0.001b 46.3k 0.2Bb Saponification value 200.4k 3.4Oa 206-t 0.4.24b I 94.0f2.83b 192.1 5 2.83b

Unsaponifiable matter 0.36k 0.03a 0.20f 0.02b 0.21 f 0.03b 0.44fr 0.014b

Tabulated values are Means k SD (N = 3) Means not having the same letter with pa:m oil along the same column are significantly different (PX.05)whereas those having the same letter with palm oil are not significantly different (Pc0.05)

Table 3.9 above shows the chemical properties of PR palm oil and its fraction. The result shows that palm olein had the highest acid value, peroxide value, Iodine value and saponification value, but the lowest unsaponifiable matter content. Palm stearin was found to have the lowest acid value, peroxide value, iodine value and saponification value but had the highest unsaponifiable matter content. The acid value, saponification value, iodine value, and peroxide value of the PR palm oil were found to differ significantly (Pc0.05) with those of its fractions. However, the unsaponifiable matter content of PR palm oil did not differ significantly (P>0.05) with that of the palm mid fraction but did differ significantly (Pc0.05) with those of palm olein and palm stearin. Table3.10: Chemical properties of partially refined shea butter and its fraction

Parameter Shea butter Shea olein Shea stearin Acid value 1.96+ 0.20a 2.24+ 0.03b 1.40 f. 0.28b Peroxide value 4.0 rt 0.28a 5.5k0.14b 2.54 0.01b Iodine value 48.2 + 0i71a 49.4 -t 0.92~1 34.010.71 b Saponification value 168.352.97 168.3+ 2.69a 176.7+ 2.12b Unsaponifiable matter 1.82+0.1la 1.202A0.14b 2.40*0.07b

Tabulated values are Means $_ SD (N = 3) Means not having the same letter wlth shea butter along the same column are significantly different (P>0.05) whereas those having the same letter with shea butted are not significantly different (P<0.05)

Table 3.10 above shows the chemical properties of shea butter and its fraction, The result shows that shea butter olein had the highest acid value and iodine value but lowest peroxide value and unsaponifiable ratter content. The shea butter stearin fraction had the highest peroxide value, saponification value and unsaponifiable matter content. However, its acid value and iodine value were the lowest. The acid value, peroxide value and unsaponifiable matter of the PR shea butter differed significantly (Pe0.05) with those of shea olein and shea stearin. The iodine value and saponification value of PR shea butter did not differ significantly (P=-0.05)with that of shea olein but differed significantly (P4l.05) with that of shea stearin. Table 3.11: Chemical property of PR dika fat and its fraction Parameter Dika fat Dika Olein dika stearin Acid value 3.09 1 0.08a 1.40 + 0.1 7b 2.81 + .25b Peroxide value 1.9 + 0.04a 2.40_+0.02b 0.7+ 0.05b Iodine value 4.95 + 0.27a 7.2 1 0.44b 2.98 + 0.23b Saponification value 208.9+ 5.4a 221.6+ 4.3b 196.4 f 8.6b Unsaponifiable matter 0.44+ 0.07a 0.27f 0.03b 0.49fO.14b

Tabulated values are Means + SD (N = 3) Means not havmg the same letter w~thd~ha fat along the same column are s~gn~ticantlyd~fferenl (P

Table 3.11 above shows the chemical properties of PR dika fat and its fractions. The result indicates that the partially refined dika fat had the highest acld value. Uika olein was also found to have the highest peroxide value, iodine value and saponification value but had the lowest unsaponifiable matter content. Dika stearin had the lowest peroxide value but had the highest unsaponifiable matter content. The acid value, peroxide value, iodine value, saponification value and unsaponifiable matter contents of the partially refined dika fat was found to vary significantly (Pc0.05) with those of dika olein and dika stearin.

Table 3.12: Chemical property of partially refined (PR) cocoabutter Parameter PR cocoa butter Acid value 2.3 + 0.06 Peroxide value 2.8k 0.03 Iodine value 36+_2.07 Saponification value 196rt: 1.57 Unsaponifiable matter 1.2 fr 0.03

Tabulated values are Means 4 SO (N = 3)

The result above shows the chemical properties of PR cocoa butter with out its fractions. The fractions were not also included because of the reason already given earlier on. Analysis showed that the acid value of PR cocoa butter varied significantly (P<0.05) with those of PR dika fat, PR palm oil, PR shea butter, dika olein, palm mid fraction, palm olein, dika stearin, stearin and shea stearin. However the acid value of PR cocoa butter did not vary significantly (P>0.05) with that of shea olein. The peroxide value of the PR cocoa butter was also found to vary significantly (Pc0.05) with those of all the other oil samples and their fractions. The iodine value of the PR cocoa butter varied significantly (P<0.05) with those of all the other oil samples and their fraction except shea stearin. The saponification value of the PR cocoa butter did not vary significantly with that of dika stearin but varied significantly (Pe0.05) with those of the other oil samples and their fractions. The unsaponifiable matter of PR cocoa butter did not vary significantly (P~0.05)with that of shea olein, but did vary significantly (Pc0.05) with those of the other oil samples and their fractions.

3.3 Enzyme properties of Aspergillus niger lipase. The results of studies of the effect of substrate concentration, pH, temperature and quantity of immobilization support on the lipase activity are presented on figures 1-5 respectively.

FIG. 1 EFFECT OF SUBSTRATE CONCENTRATION ON LIPASE ACTIVITY Intercept a -2.167 x 1 0-3

- Slope = 0.102

1/[sl FIG. 2: LINE WEAVER BURK PLOT OF ENZYME ACTIVITY AGAINST SUBSTRATE CONCENTRATION

FIG 3 EFFECT OF pH ON ENZYME ACTIVITY FIG 4. EFFECT OF TEMPERATURE ON ENZYME ACTIVITY

0.25 0.5 0.75 1 1.25 1.5 1.75 2 QUANTITY OF SUPPORT (9) FIG 5 EFFECT OF QUANTITY OF IMMOBILIZATION SUPPORT ON ENZYME ACTIVITY .

Figure I represent the effect of increasing substrate concentration on enzyme activity. The figure shows that the activity of the enzyme increased with increase in substrate concentration .The opimal activity was observed at 30mM oil concentration. Figure 2 shows the Lineweaver-Burk plot of enzyme activity against concentration. The data (Fig 2 ) gives the Km and Vmax values of the enzyme as 46.7918mM and 461.554u/l,respectively.one unit of enzyme activity is defined as the amount of fatty acid liberated per unit time (IOmin) under optimum conditions.

Figure 3 above represents the effect of pH on the enyme activity. Result shows that activity of the enzyme increased progressively with increasing pH (Fig3) from pH 6.2 to pH 7.2.Thereafter, there was a progressive declineln effect, the optimum pH of the enzyme is 7.2.Activity then decreased progressively showing loss of enzyme activity.Figure 4 above shows the effect of increase in temperature on lipase activity. The optimum temperature of the enzyme (Fig 4) was found to be 40•‹C after which there was a decline and complete loss of enzyme activity at 60•‹C. Figure 5 above shows the effect of quantity of inorganic support on enzyme activity. Optimal enzyme activity was obtained with 0.5g of immobilization support material. Values above 0.5g resulted in progressive decrease in enzyme activity. 3.4 Physical and chemical properties of inter-esterified products. The melting 1 slip points of 21 inter-esterified samples and fatty acid composition of five selected interesterified sam~lesare given on Tables 3.13 and 3.14, respectively.

Table 3.13: The melting I slip points of the inter-esterified products of fractions from palm oil, shea butter and dika fat. Sample Melting pointlslip point (OC) Cocoa butter 30-34 Pam mid fraction + shea stearin 34 - 36 Palm mid fraction + shea olein 30 - 32 Palm mid fraction + shea crude 32 - 34 Palm mid fraction + Dika olein 35 - 36.5 Palm mid fraction + Dika stearin 37 - 38 Palm mid fraction + Dika crude 36 - 38 Palm olein + shea crude 31 - 33 Palm olein + shea stearin 34 - 35 Palm olein + Dika olein Palm olein + Dika stearin Palm olein + Dika crude 35 - 37 Palm stearin + shea stearin 36 - 38 Palm stearin + shea olein 35 - 36 Palm stearin + shea crude 34 - 36 Palm stearin + Dika olein 35 - 36.5

Palm stearin -I-Dika stearin 36 - 38 Palm stearin + Dika crude 36 - 37 Shea olein + Dika olein 35 - 37 Shea butter olein + Dika crude 37 - 38 Shea butter stearin + Dika olein 36 - 37 Shea butter crude + Dika crude 38 - 40

Table 3.13 above shows the slip melting point of the intere-esterified products. As can be seen from table, palm mid fraction + shea olein, palm mid fraction + shea butter crude, palm olein + shea butter crude, palm olein + dika olein, palm mid fraction + shea stearin, palm mid fraction + dika olein, palrr olein + shea stearin, palm stearin + shea butter crude and palm stearin + shea olein all had melting ranges that compares to that of the cocoa butter sample used in this work.Namely:30-32,32-34,30-32,34-36,35.5-36.5,34-35,34-36,35- 36,respectively. products.

Name of product. Oleic acid Stearic Palmitic acid mgllOOml acidmgl100ml mg/100ml Palm mid Faction + shea stearin 1.8742k 0.08b

Palm mid Fraction + shea olein 1.6299k0.12a

Palm mid fraction + shea crude 1.92082 0.09b

Palm olein + shea butter crude 2.1 170k 0.09b

Palm olein + dika olein 0.5334k 0.03b

Cocoa butter crude 1.6322+ 0.07a

Tabulated values are Means k SD (N = 3) Means having the same letter with cacoa butler along the same vertical axis are not significantly different (~~0.05)whereas means having different letter are significantly different (Pc0.05)

Table 3.14 above shows the oleic, stearic and palmitic acid content of selected interesterified products. The result shows that palm olein + PR shea butter had the highest oleic acid content while palm olein + dika olein produced the lowest. PR cocoa butter had the highest stearic acid content while palm olein + dika olein produced the lowest. Palm midfraction + shee crude had the highest palmitic acid content while palm oleint + dika olein had the lowest. Statistical analysis showed that the oleic acid content of the PR cocoa butter did not differ significantly (P>0.05) from that of palm mid fraction + shea olein. In addition, the stearic acid content of PR cocoa butter did not differ from that of palm mid fraction + shea stearin. However, the palmitic acid content of PR cocoa butter differed significantly (P<0.05) from that produced by all the other interesterified products CHAPTER FOUR DlSCUSSlON

4.1 The status of partially refined palm oil, shea butter, dika fat and their fractions as potential cocoa butter substitutes. All the samples used had good yield of oil that are comparable to other oil seeds. The % yield of palm oil is comparable to 55% reported by Gordon (7993) and 50-65% reported by Ononogbu (2002). The % yield of dika fat compares well with the range of 55-70% reported by Achinewhu (1998) and 68-75% obtained by Onwuka et a/. (1984). The % yield of shea butter is comparable to 50-60% reported by Achinewhu (1998) and higher than 42-45% reported by Ononogbu (2002). Farinu (1986) also reported a % yield of up to 60.7% for shea butter. Cocoa butter had a % oil yield that compares well with 43.8-46.2% reported by Ojeh (1980). The % oil yield of these four samples is also comparable to other oil seeds,namely,castor-50%, cotton seed-30%, linseed-40%, palm kenel-50%, rapeseed-40•‹h, sun flower 35%, groundnut-50%, (Carr, 1989).

The yield in gram per kilogram obtained for palm oil, shea butter, and dika fat, places them as highly ranked oil bearing materials. Their values are high and comparable to those of other highly rated oil producing seeds such as 500g for castor oil, 620g for copra oil, 4209 for mustard oil, 370g for rapeseed oil, 500g for sesame oil 320g for sunflower oil and 420g for pam fruit (Adlof and Duchateau, 2006). However, it is pertinent to note that oil yields vary widely from one geographical location to another and from one variety to another. The yield of palm oil, shea butter and dika fat were high compared to cocoa butter and justify the study on their possible use as substitute for cocoa butter.

The differences in the temperature ranges under which fractionation of the oil samples were carried out is very remarkable and could be attributed to the level of saturation and unsaturation of the fatty acids in the three oil samples. Unsaturated acids with low melting points crystallize at lower temperatures while saturated acids can be crystallized at higher temperatures (Gunstone @ aJ., 1994). The fractionation at varying temperatures and solvents was done to obtain a good olein-stearin separation that is suitable for this study. Good olein- stearin separation obtained with the lowest amount of solvent was chosen as optimal. This agrees with an earlier result on Monodora myristica oil (Njoku gi.,1996) and rice bran oil (El-zanati and Khedr, 1991) which stated that.fractionation at lowest solvent ratio should be chosen as optimal due to high cost associated with solvents.

The specific gravity of PR palm oil and its fractiox compares favourably with 0.898-0.907 reported for palm oil by Umoh (1998). The specific gravity of PR shea butter and its fractions compares well with 0.927 obtained by Badifu and Abah (1998), but slightly lower than 0.967 reported by Njoku et a/. (2000). The specific gravity of PR dika fat and its fractions compares well with that obtained by Eka (1980) but low compared to 0.968 reported by Njoku & d., (2000). Cocoa butter gave a specific gravity value that is comparable to 0.915 reported by Zoumas and Finnegan (1979). The specific gravity values obtained in all the PR oil samples and their respective fractions fits into the range of 0.8-1 reported by Gordon (1993) for most vegetable oils. There were generally, higher specific gravity values for the olein fractions and lower values for the stearin fractions compared to the PR oil samples. Such could imply that the olein fraction of oils will be weightier than their stearin and crude samples.

The refractive index of PR palm oil and its fractions differed slightly from 1.453-1.459 reported by Achinewhu (1998) for palm oil. The PR shea butter and its fractions have refractive index values which compares well with 1.462 reported by Njoku gt d.,(2000) and 7.463-1.467 recorded by Gordon (1993). The PR dika fat and its fractions gave refractive index values that slightly compares to 1-473 reported by Njoku and Ugwuanyi (1997) for dika fat. The refractive index value obtained for PR cocoa butter is higher than the 1.456 reported by Ojeh (1980) and slightly higher than 1.456-1.458 reported by Zoumas and Finnegan (1979). Higher refractive index values were observed in the olein fractions compared to the stearin fractions. This could be attributed to thk differences in saturation and unsaturation of these two fractions, This also agrees with that stated that refractive index falls with increase in melting point of oils (FAO, 1997). The higher refractive index also reported in this study coutd be attributed to the partial refining of these oil samples as it has been shown that refined oils have slightly higher refractive index values compared to their crude samples (Ojeh, 1980).

The viscosities of PR shea butter,dika fat, and those of their stearin fractlons were very close. But a reverse of this trend was observed for PR palm oil. The observed differences in viscosity trend among these oil samples could be due to the temperature at which the measurements were taken (45'~).This temperature far exceeds the melting point of palm oil and therefore there will be very low intermolecular attraction among it molecules compared to PR shea butter and dika fat which both have higher melting points. The closeness of the viscosity values obtained from PR palm oil, PR shea butter, shea butter olein and shea butter stearin to that of cocoa butter is very remarkable. The viscosity values obtained from these oil samples and their fractions could be considered high when compared to those of other oils such as 37.82cst for rapeseed oil, 34.90cst for safflower oil, 32.60cst for soybean o~l, 34.70 for sun flower oil, 29.30cst for castor seed oil (Quick,l989). These differences in viscosity measurements are mainly due to the method applied in this study and due to the temperature used for the study as viscosity is highly affected by temperature.

Apart from the melting point of palm olein which could not be determined due to its inability to crystallize under the available means of temperature analysis, melting point range of the partially refined oil samples varied when fractionated to give a lower melting and a higher melting stearin fractions. The melting point range difference between olein and strearin fractions was much higher in palm oil compared to shea butter and dika fat. The melting point range of shea olein, palm mid fraction and palm stearin are comparable to that of cocoa butter. The high values obtained from the oil samples studied could be attributed to the relatively high proportions of saturated fatty acids.

The acid values of PR palm oil and its fractions are high when compared with 2 to 15 recorded for crude palm oil by Umoh (1998). The acid value of PR shea butter and its fractions compares with 1.54 reported by Njoku gt aJ., (2000). The acid value of PR dika fat and its fractions are low conipared with 9.25 obtained by Eka (1980) and 15.4 obtained by Njoku and Ugwuanyi (1997). The acid value obtained for PR cocoa butter is close to the value of 2.1 reported by Zoumas and Finnegan (1979). The acid values of the PR samples of palm oil, shea butter and dika fat were found to differ significantly from those of their fractions. This indicates significant variations in their free fatty acid contents. The olein fractions of all the samples were found to have highest acid content while the stearin fractions had the lowest free fatty acid values. The peroxide values of all the PR oil samples differed significantly from those of their fractions. Highest peroxide values were obtained in the olein fractions. This indicates that the olein fractions are most susceptible to oxidation compared to the PR oil samples and the stearin fractions. The peroxide value obtained for PR cocoa butter compared well with 2.57-3.20 recorded by Ojeh (1980). The peroxide value of PR cocoa butter differed significantly from those of other PR oil samples and their fractions. All the oil samples generally had low peroxide values. Such low peroxide values are indicative of low susceptibility to oxidative rancidity and thus high stability. The stability of these oils could be due to the presence of a high-level of saturated fatty acids (Badifu and Abah, 1998). Stability could also be attributed to the presence of antioxidants [Kar and Mital, 1981; Ong, 1989). The iodine values of the PR palm oil and its fractions are comparable to the range of 46-60 reported by Umoh (1998) and also comparable to 53.2-54.0 for crude palm oil, 57.0-60.0 for palm olein and 43.0-47.3 for palm stearin recorded by Ong (1983). The iodine values of the PR shea butter and its olein fraction compares well with 50 and 45.62 reported by Badifu and Abah (1998) and Njoku a aJ., (2000), respectively, for crude shea butter. However, its stearin fraction does not compare well with the above values. PR dika fat had iodine values that are low when compared with 19 obtained by Njoku and Ugwuanyi (1997) but comparable to 3.83 obtained by Eka (1980). The iodine value of PR cocoa butter is comparable to the range of 33-36.0 reported by Ojeh (1980) and 25.7-44.0 recorded by Zoumas and Finnegan (1979) for crude cocoa butter. The iodine values of the PR sample of palm oil, shea butter and dika fat were found to differ significantly from those their fractions except between PR shea butter and its olein fraction. In addition, the iodine value of the PR cocoa butter did not compare well with that of shea stearin.The differences between iodine values of PR oil samples and those of their fractions suggest differences in concentration of saturated and unsaturated fatty acids in the different PR oil samples and their fractions. Dika fat had the lowest iodine number in all the samples and this suggests a high level concentration of saturated fatty acids. Iodine number increases as the level of unsaturation increases and, vice versa.

The saponification value of PR palm oll and fractions compares well with the range of 196- 209 recorded by Eka (1980). The saponification value of PR shea butter and its fractions are slightly low when compared with 186.64 obtained by Njoku gt gl.,(2000) but compares well with the 169.64 reported by Badifu and Abah (1998). The saponification value of crude dika fat is very high when compared with the 101 reported by Njoku gl.,(1997) but lower than 230 reported by Dalziel (1955). However Eka (1980) reported a saponification value of 224.8 for crude dika fat. The saponification value Of cocoa butter was higher than 183.1-188.3 obtained by Ojeh (1980), but compares well with 192-196 recorded by Zoumas and Finnegan (1978). The saponification values of the PR oil samples differed significantly from those of their fractions except between PR shea butter and it's olein fraction. The similar saponification values of these oil samples would suggest similar molecular weight fatty acids (Jacobs, 1958).

The unsaponifiable matter content of PR palm oil and its fractions compares with 0.3-1.0 reported by Achinewhu (1998). PR shea butter and its fraction gave unsaponifiable matter contents that relates well with 1.2 recorded by Badifu and Abah (1998) and Njoku gt a,, (2000), but low when compared with 4.8 reported by Gordon (1993). Unsaponifiable matter content of PR dika fat gave values that are high compared to 0.12 obtained by Njoku and Ugwuanyi (1997) but compare with 0.45 reported by Eka (1980). PR cocoa butter gave an unsaponifiable matter content that falls into the range of 1.10 -1.34 reported by Ojeh (1980) but high when compared with 0.35 obtained by Zoumas and Finnegan (1979).The partially refined oil samples and their fractions had significant differences in their unsaponifiable matter content except between PR palm oil and its mid fraction (pmf). Cocoa butter also had similar unsaponifiable matter content with shea olein. The stearin fractions generally had higher unsapponifiable matter content. This suggests a higher concentration in high molecular weight sterols and phosphatides, which could be present in these oil samples (Ibemesi 1992).

From the above, it can be inferred that the chemical and physical properties of these fats samples could differ significantly when fractionated except in few cases which have been pointed out. Also some of the PR oll samples and their fractions namely- PR shea butter, dika olein, palm mid fraction, PR palm oil add shea butter stearin have physico-chemical properties that are comparable to those of cocoa butter. This suggests the possible use of fractions from palm oil and shea butter as cocoa butter substitutes, (Ong, 1989; Gordon, 1993; Young gt &I.,1994; Xu, 2000).

4.2 Assay of the Immobilized Inter-esterification Aspergillus niger lipase The 2.5ml aliquot of enzyme preparation used for this study was found to have optimal activity at 0.5g quantity of support material. The slightly lower activity recorded at 0.25g could be due to inadequate amount of the carrier, Inadequate amount of support could create competition for the avalaible space for the immobilization (Owusu-Ansah, 1994). The optimal activity recorded at 0.59 could be as a result of adequate amount of carrier to accommodate most of the enzyme in solution. This makes more of the enzyme available for activity and avoids being lost during filteration. The lower activity observed at 0.79-29 could be due to high quantity of carrier thereby creating increased surface for immobilization of the given quantity of enzyme preparation used. These values differ from that of 3.2g reported by Macrae (1983) using 250mg of lipase prepared from Rhizopus delemar. Yokozeki et al., (1982) reported even lower values of 0.259 using 5mg of Rdelemar lipase immobilized on celite. This difference in values could be attributed to quantity of enzyme preparation used and type of support material employed.

The optimal pH for activity observed for Aspergillus niger lipase tended towards the alkaline range. This is comparable to a report by Ohnish~a a,,(1994) who observed an alkaline lipase having optimal pH of 7.5 and 10 for Aspergillus oryzae . A.terreus was also found to show good pH tolerance at pH 3.0-12.0 with optima at 5.5 and 10 (Yadav et al., 1998). However, the value is slightly higher than that obtained from other Aspergillus spp such as pH 5.3 obtained for A.repens (Kaminishi gj d:,19991, pH 5.5 for A. Wentii (Chander gt aJ., 1980). The result obtained also compares with other fungal lipases such as pH of 7.0 reported as optimal for Penicillin cyclopium (Chahinian gt a., 2000), pH optimum of 8.0 for P. aeruginosa (Chartrain a aJ., 1993) and pH of 9.0 for P, expansum (Stocklein gt aJ., 1993).As lipase catalysed reaction are sensitive to the pH of the reaction systern,the pH optima of 7.2 observed for Aspergillus niger lipase used in this study guided the reaction pH to be adjusted so as to enable the lipase to function at its best state.This will enhance the rate of inter- esterification reaction. Thermal stability is a desirable characteristic of lipases especially those for industrial applications. This is because the productivity of the reaction can be greatly enhanced by operating at a relatively high temperature (Janssen a., 1994). Aspergillus niger lipase showed activity at a temperature range of 25-55' C with optimum at 40' C. Such stability at a temperature range is very useful for industrial application as it can withstand sudden changes in operational temperature. The optimal temperature for Aspergillus niger is high compared with 27' C obtained as optimum for Aspergillus repens by Kaminishi aJ., (1999). However, it compares well with other fungal lipases such as 40'~ obtained as optimum temperature for Penicillium cyclolpium (Chahinian gt a., 2000), 45% for Penicilljum expansum (Stocklein gt a.,1993). Sekero gt d.,(2004) obtained even higher values of 60'~from a lipase produced from Candida antarctica. The stability observed in Aspergillus niger lipase may have been enhanced by immobilization. Arroyo gt a. (j999) and Hiol a d. (2000) stated that native enzyme and the covalently immobilized enzyme preparation appeared to follow different modes for thermal deactivation. Thermal stability of lipases is related with their structure (Zhu g gl.,2001).The temperature optimum of 40•‹C for the inter-esterification lipase from A.niger will greatly enhance the rate of inter esterification and thus lead to quicker production of the inter-esterified products.Such low energy demand is also of great advantage in the industries for it circumvents the high cost associated with reactions requiring high energy imput.The graph of V, against So had the form of a rectangular hyperbola. This is consistent with experimental findings for many enzyme- catalyzed reactions (Palmer, 1985). The parameters of Vmax and Km allows us to evaluate the kinetic efficiency of enzymes. Vmax which relates with kcat signifies the turn over number of the enzyme .It is equivalent to the number of substrate molecules converted to product in a given unit of time on a single enzyme molecule when the enzyme is saturated with substrate (Nelson and Cox, 2000).The Vmax obtained is lower than 520 u/l obtained for B thermoleovorans 1D-1 (Lee gt gl.,1999). These differences in V maxvalues can be attributed to the total concentration of enzyme present. Km give the substrate concentration that produces half maximal velocity .Therefore when the substrate concentration is equal to the km ,the initial velocity is half maximal .This shows that there are still some free enzymes in solution.Addition of more substrates in such circumstances could lead to generation of more product.ln order to maximize the catalytic activity of the lipase, the Km value was used to measure the appropriate amount of oil used for inter-esterification by a specific lipase concentration so as to effectively enhance the production of inter-esterified products.The Km value of 47 suggests only half maximal velocity of the enzyme preparation used.This value is higher than that of 12mM obtained for P. Iepacia lipase (Pencreach and Baratti 1996).High Km of Aspergillus niger lipase also suggests a good affinity of the enzyme for its substrate. This shows wide ranges in Km values and agree with that earlier stated by Fullbrook (1996). Km values of enzymes has also been shown to differ with different substrates (Hatzinikolaou

-et -.Ial 1999).

4.3 Inter-esterification products of Aspergillus niger lipase Owusuh-Ansah (1994) stated that cocoa butter should be devoid of solid fat at 37' C. Sharma a &I.,(2001) stated a melting point range of approximately 30-36' C for cocoa butter. Therefore, interesterified products, which fall under the melting point range of cocoa butter used for this study, could be said to compare well generally with cocoa butter. Products such as palm mid fraction + shea stearin, palm mid fraction +dika olein, palm olein + shea stearin, palm stearin + shea olein, palm stearin + shea crude and palm stearin + dika olein come under this category. However the fact that these products have melting point range within that of cocoa butter does not guarantee their suitability as cocoa butter equivalents. Cocoa-butter equivalents should have chemical composition that is closely related to that of cocoa butter. However, they can be regarded as being under cocoa butter substitutes fats but further work needs to be done in order to determine their suitability as cocoa butter substitutes. Such parameters as solidification, carbon number, solid fat index, dilatation & other chemical characteristics still need to be checked. Charkrabaty et al (1983) have studied and produced valuable cocoa butter substitutes from sal fat and mango fat.

Cocoa butter has a relatively simple triaqlgTycerol composition that results in unique properties. The typical triacylglycerol composition is POP 16%, POS 35%, SOS,26%, PO0 4%, S006%, PLP,2%, PLS4%, PLO,I0h and others 4%, (Shukla 1996). This shows that it contains 80% of POS, SOS and POP triacyfglycerol. According to Dimick and Manning (1987)' Ivory coast cocoa butter, one of the finest in the world, has a composition of14.8%POP, 45.4% POS and 28.8% SOS. In terms of its fatty acid composition, Owusuh- Ansah (1994) stated that cocoa butter has a typical composition of 26% palmitic acid, 36% stearic acid, 33% olec acid and 3% lindeic acid. Xu (2000) also reported a typical fatty acid composition of 24.4% palmtic acid, 37.0% stearic acid, 33.6% oleic acid, 3.4% linoleic acid and 1.6% of others. From the above one observes that cocoa butter generally has stearic acid as it's highest amount of fatty acid, closely followed by oleic acid and then palmitic acid. Amongst the interesterified products, such trend as observed in cocoa butter was not found. However the closeness of oleic acid content of cocoa butter and palm mid fraction + shea olein is very remarkable. Also the closeness of the stearic acid content of cocoa butter and that of palm mid fraction + shea crude is also remarkable. None of the interesterified products had palmitic acid content that compares statistically with that cocoa butter.lnter- esterified products of paml mid fraction+shea stearin and palm olein+shea butter crude had stearic acid values that are close to that of cocoa butter.The oleic acid concentration of cocoa butter was even lower than that obtained for palm mid fraction+shea stearin and palm olein+shea butter crude.Such high oleic acid content is desirable for a fat to be considered as a cocoa butter equivalent fat.This is because cocoa butter has oleic as one of its major fatty acids.lt also known that oleic acid occupies all the 2-Sn position of cocoa butter's triacylglycerol(Macrae,1983). From the above, it can be suggested that the inter-esterified products of Palm mid fraction+Shea butter stearin,Palm mid fraction+shea olein,Palm mid fraction+Shea butter crude and Palm olein+Shea butter crude all hold promises as cocoa butter equivalents based on their close fatty acid concentration..

It is pertinent to note that the melting characteristics and fatty acid concentration were used to assess the closeness of the products obtained to cocoa butter. In this regard products such as palm mid fraction + shea stearin, palm mid fraction + shea olein, palm olein + shea butter crude, palm mid fraction + shea butter crude, palm olein + dika olein compared well with cocoa butter in melting points. Only palm mid fraction + shea butter crude had fatty acid contents that compared well with cocoa butter. Also palm olein + dika olein gave fatty acid values that are seemingly too low compared with that observed for cocoa butter and therefore may not be a good cocoa butter equivalent. However further work is still required on these samples for a more conclusive report. This could best be achieved by using GLC to check their fatty acid composition, carbon number and also the distribution of the fatty acids on the triacylgycerol molecules of the products obtained. However, due to unavailability of the equipment and other constraints, this could not be achieved. However this serves as an indicator to show that blending of fractions from palm oil, shea butter and dika fat could produce products which resemble cocoa butter in melting characteristics and fatty acid concentration as best observed in the result obtained in this study.

CONCLUSION This project has shown that palm oil, shea butter and dika fat can be separated into stearin and olein fractions which differ significantly in physical and chemical properties such as iodine number, peroxide value, viscosity, acid value, melting point and saponification value. The quantity of solvents required for fractional crystallization of saturated fat such as dika fat, and shea butter can be reduced by fractionating at lower temperatures. Aspergillus niger lipase was found to be stable over a pH range 6-9 and temperature range of 25-55'~. It also had pH and temperature optimal of 7.2 and 40'~respectively. Lipase catalyzed inter- esterification of fraction from palm oil, shea butter and dika fat produced fats that compares with cocoa butter in melting point and fatty acid concentration. SUGGESTIONS The fatloil used in this study are used locally for food uses. Palm oil and shea butter are also known for some industrial uses. The use of these oils as substitute for cocoa butter could put pressure on the demand for these various uses. However, in consideration of the high value attached to cocoa butter it is believed that conversion of these oils into cocoa butter substitute/equivalents will yield value added products and thus contribute to the improvement of the economy of the tropical countries. PQ- $4 c?lsnu mPtT *.- ,%

However more studies still need to be carried out on these products such as determination of the solid fat index, monitoring of the rate of ester interchange during inter-esterification, determination of the fatty acid and triacylglycerol composition of the oil fractions before inter- esterification and that of the interesterified product. Finally the blending of various amounts of the cocoa butter substitutes and cocoa butter need to be done to determine the extent of compliance or deviation from the original qualities observed in cocoa butter. These could not be done due to lack of equipment like .HPLCIGLC and unavailability of standard triacylglycerols.

Lipase catalysed production of specificaSly structured triacylglycerol in industry is not yet widely implemented and the technology is still being developed. Nevertheless it is expected that lipase application for the modification of oils and fat will increase in the industries in the near future due to numerous advantages it offers over traditional chemical methods. REFERENCES

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0 20 40 6 0 80 100 120 Final Concentration (uMIL)

Fig.?; Standard Curve for Palmitic Acid Concentration (uM/L)

Fig. 8:Standard Curve for Stearic Acid