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PURIFICATION AND CHARACTERIZATION OF
LIPASE (EC.3.1.1.3) FROM THE SEEDS OF
Cucumeropsis mannii (WHITE MELON).
BY
EZEMA, BENJAMIN ONYEBUCHI.
PG/M.Sc/09/51506.
DEPARTMENT OF BIOCHEMISTRY,
UNIVERSITY OF NIGERIA, NSUKKA.
JULY, 2012.
UNIVERSITY OF NIGERIA, NSUKKA.
DEPARTMENT OF BIOCHEMISTRY.
PURIFICATION AND CHARACTERIZATION OF
LIPASE(EC.3.1.1.3) FROM THE SEEDS OF Cucumeropsis mannii
(WHITE MELON).
BY
EZEMA,BENJAMIN ONYEBUCHI
PG/M.Sc/09/51506.
BEING A PROJECT REPORT SUBMITTED TO THE SCHOOL
OF POSTGRADUATE STUDIES IN PARTIAL FULFILMENT OF
THE REQUIREMENTS FOR THE AWARD OF MASTER OF
SCIENCE (M.Sc) DEGREE IN BIOCHEMISTRY,
UNIVERSITY OF NIGERIA,NSUKKA.
SUPERVISOR: DR.S.O.O. EZE.
2
DEPARTMENT OF BIOCHEMISTRY,
UNIVERSITY OF NIGERIA,NSUKKA.
CERTIFICATION
Mr Ezema,Benjamin.O, Reg. No: PG/M.Sc/09/51506, a Postgraduate student of the
Department of Biochemistry, University of Nigeria, Nsukka, has satisfactorily
completed the requirements of research work for the Award of Degree of Master of
Science (M.Sc) in Biochemistry. The work incorporated in this dissertation is original
and had not been submitted in full or in part, for any other diploma or degree of this
or any other University.
……………………………… ………………………………
Dr.S.O.O. Eze Prof. L.U.S. Ezeanyika
(Supervisor) (Head of Department)
…………………………………………
External Examiner
DEDICATION
3
This work is dedicated to my mother, Mrs. Ezema Evelyn (Nee Ugwu) of blessed
memory for her motherly care and love before her departure from the earth.
ACKNOWLEDGEMENT
My special thanks goes to God almighty that made this project work possible. This
research could not have been possible without the encouragement and assistance of
numerous individuals to whom I owe my gratitude. I am deeply indebted to my
supervisor Dr. S.O.O. Eze for critically supervising and reading this work. I will also
remain grateful to Dr. Parker .E. Joshua who was always handy to help and advice me
in the course of this research work.
4
My gratitude goes to the teaching staff of Biochemistry Department who in one way
or the other contributed ideas and solutions to the challenges encountered during this
research, especially Professors. L.U.S. Ezeanyika (Head Department of
Biochemistry), O.F.C. Nwodo, I.N.E. Onwura, F.C. Chilaka, O.U. Njoku, P.
Uzoegwu and E.O. Alumanah.
Others are Dr. Onwubiko, Dr. V.N. Ogugua, Dr. (Mrs.) Anosike, Dr. O.C. Enechi
and Dr. Ubani. I also thank Mr Obina Ojeh,Ozioko Paul,Ukonu Christian,Mmadueke
Ebubechukwu,Joy, Florence and all the other colleagues I worked with in the
laboratory for their friendly relationship and assistance.
I am also grateful to the members of my family, Amoge, Ngozi, and Barr F.O.
Ukwueze for their encouragements all through the days of this research.
5
ABSTRACT
Lipase (triacylglycerol acylhydrolase, EC. 3.1.1.3) was isolated from the endosperm
of 4 days germinated seeds of Cucumeropsis mannii. Homogenate of the
Cucumeropsis mannii endosperm prepared on the 4th
day of germination was
fractionated by centrifugation at 5000xg for 15mins to yield a fat layer (lipid
bodies), supernatant (water soluble fraction) and pellets. Lipase activity was found
to be high in the lipid bodies, followed by the water soluble fraction and least in the
pellets. Solubilization of the lipase in the lipid bodies was investigated using two
detergents (Tween 80 and Triton X-100). Best solubilization was obtained at 1.95%
(w/v) concentration of Tween 80. The supernatant (water soluble fraction) obtained
after solubilization and centrifugation of the extract was used as the crude lipase.
The crude enzyme was purified through a 3 step purification procedures; combined
(NH4)2SO4 and cold acetone precipitation, followed by dialysis and sephadex G-200
gel filtration column chromatography. Two times gel filtration column
chromatography of the enzyme after dialysis gave two peaks with lipase activities
indicating the presence of two forms of lipases in the seed (alkaline and acid
lipases). Studies on the two lipases showed that alkaline lipase which was
designated lipase A have optimum pH of 7.5 while the acid lipase designated lipase
B has optimum pH of 5.9. The alkaline lipase was found to be stable between pH
6.5 and 8.0 while the acid lipase was found to be stable between pH 4.5 and 6.0.
The two lipase fractions showed optimum temperature of 370C and were stable at
temperatures up to 450C for 1 hr. Ca
2+ was observed to be a very good activator and
Pb2+,
a potent inhibitor of the two lipases. At the end of the final stage of
purification, purification folds of 5.4 and 11.0 were obtained for the alkaline and
acid lipase respectively. Kinetic studies on the two lipase fractions showed that the
alkaline lipase have Vmax of 142.8U and Km of 13.3g/L while the acid lipase have
Vmax value of 166.7U and Km value of 15.7g/L of the substrate (Olive oil).
TABLE OF CONTENTS
6
Title page
Approval
Dedication i
Acknowledgement ii
Abstract iii
Table of content iv
List of figures vii
List of tables
viii
List of plates ix
List of appendices x
CHAPTER ONE: INTRODUCTION 1
General introduction 1
1.0 Brief history of enzyme 2
1.1 Meaning of enzyme 2
1.2 Lipases 2
1.3 Three-dimensional structure of lipase 3
1.4 Classification of lipases 4
1.5 Sources of lipases 5
1.5.1 Seed lipases 5
1.6 Role of seed lipases 6
1.7 Specificity of seed lipases 7
1.8 Oil seed lipases 8
1.9 Factors affecting lipases 10
1.9.1 Moisture 10
1.9.2 Temperature 10
1.9.3 Effect of pH 10
1.9.4 Activators 10
1.9.5 Inhibitors 10
1.10 Applications of lipases 11
1.10.1 Application of lipase in food industries 11
1.10.2 Application of lipase in detergent industries 11
1.10.3 Application of lipase in oil and fat industries 12
7
1.10.4 Application of lipase in fine chemical industries
12
1.10.5 Application of lipase in biodiesel production
12
1.11.0 Cucumeropsis mannii
13
1.11.1 Nutritional composition of Cucumeropsis mannii
13
1.11.2 Scientific classification of the plant Cucumeropsis mannii
14
1.11.3 Production and processing
16
1.11.4 Uses of Cucumeropsis mannii
16
1.12 Aim of the research
17
1.13 Objective of the study 17
CHAPTER TWO: MATERIALS AND METHODS
18
2.0 Materials
18
2.1 Plant material
18
2.2 Chemicals
18
2.3 Equipments
18
2.4 Methods
18
2.4.1 Reagent preparation
18
2.4.2 Seed germination
20
8
2.5.0 Germination parameters
20
2.5.1 Germination energy
20
2.5.2 Water sensitivity
21
2.5.3 Average root length
21
2.6.0 Lipase extraction
23
2.7.0 Preparation of substrate emulsion
23
2.8.0 Lipase assay
23
2.8.1 Measurement of lipase activity during seed germination
24
2.8.1 Localization of Cucumeropsis mannii lipase
24
2.8.2 Solubilization of lipase
25
2.9.0 Purification of Cucumeropsis mannii lipase
27
2.9.1 Isolation of lipase
27
2.9.2 Solubilization of lipase
27
2.9.3 Precipitation of lipase
27
2.9.4 Ammonium sulphate precipitation
27
2.9.5 Cold acetone precipitation
28
2.9.6 Dialysis
28
9
2.9.7 Gel filtration column chromatography
28
2.10.0 Characterization of Cucumeropsis mannii lipase
28
2.10.1 Effect of pH on activity and stability lipase
28
2.10.2 Effect of temperature on activity and stability of lipase
29
2.10.3 Effect of metal ions and EDTA on activity of lipase
29
CHAPTER THREE: RESULTS
30
3.0 Results
30
3.1 Germination Parameters
30
3.2 Result of variation of lipase activity and protein with
Period of germination in days
30
3.3 Result of the comparison of lipase activity in the coated and uncoated
Seeds of C. mannii 32
3.4 Result of the comparison of protein concentration in the coated and
uncoated Seeds of C. mannii
32
3.5 Result of the pH profile on the crude lipase
34
3.6 Result of localization of Cucumeropsis manni lipase
35
3.7 Result of solubilisation of Cucumeropsis mannii lipase
35
3.8 Result of purification of Cucumeropsis mannii lipase
36
3.9 Result of the preliminary study on ammonium sulphate precipitation
37
10
3.10 Result of the preliminary study on cold acetone precipitation
37
3.11 Result of the effect of pH on activity and stability of C. mannii lipase
40
3.12 Result of the effect of temperature on activity and stability of
C. mannii lipase
40
3.13 Result of effect of metal ions on Cucumeropsis mannii lipase
45
3.14 Result of kinetic studies on Cucumeropsis mannii lipase
47
CHAPER FOUR: DISCUSSION AND CONCLUSION.
49
4.0 Discussion 49
4.1 Conclusion 52
References 53
Appendices
63
LIST OF FIGURES
Figure.1: Three- dimensional structure of lipase
4
Figure 2: Chart showing how triacylglycerols (TAG) stored in the lipid bodies
are 7
hydrolyzed to fatty acids (FA) and glycerol by the sequential action
of one or more lipases
Figure.3: Variation of lipase activity with period of germination
31
Figure.4: Variation in protein concentration during germination
31
Figure.5: Comparison of lipase activity in coated and uncoated seeds of
C. mannii during germination.
32
11
Figure.6: Comparison of changes in protein concentration in coated and
Uncoated seeds of C. mannii during germination.
32
Figure.7: pH profile of the crude lipase
34
Figure.8: localization of lipase activity in the 3 fractions obtained
35
after centrifugation.
Figure.9: Result of solubilization of lipase .
35
Figure.10: Result of preliminary study on ammonium sulphate precipitation.
37
Figure.11: Result of preliminary study on cold acetone precipitation.
37
Figure.12: Chromatogram for the first gel filtration
38
Figure.13: Chromatogram for the second gel filtration
38
Figure.14: pH optimum for lipase A
41
Figure.15: pH optimum for lipase B
41
Figure.16: pH stability of lipase A
42
Figure.17: pH stability of lipase B
42
Figure.18: Optimum temperature for lipase A
43
Figure.19: Optimum temperature for lipase B
43
Figure.20: Temperature stability of lipase A
44
Figure.21: Temperature stability of lipase B
44
12
Figure.22: Effect of metal ions on lipase A
46
Figure.23: Effect of metal ions on lipase B
46
Figure.24: Lineweaver-Burk plot for lipase A
48
Figure.25: Lineweaver-Burk plot for lipase B
48
LIST OF TABLES
Table.1: Some of the most studied seed lipases and their main
Features and biochemical properties.
9
Table.2: Some seed lipases and their applications.
13
Table.3: Purification table for lipases isolated from Cucumeropsis mannii.
39
13
LIST OF PLATES
Plate.1: Photograph of the plant Cucumeropsis mannii.
15
Plate.2: Photograph of dry seeds of Cucumeropsis mannii.
15
Plate.3: Photograph of seeds of Cucumeropsis mannii at different
days of germination .
22
Plate.4: Photograph showing the clarity of lipid bodies during
solubilization at different concentrations of detergent.
26
14
PURIFICATION AND
CHARACTERIZATION OF LIPASE (EC.3.1.1.3)
FROM THE SEEDS OF Cucumeropsis mannii
(WHITE MELON).
BY
EZEMA, BENJAMIN ONYEBUCHI.
PG/M.Sc/09/51506.
DEPARTMENT OF BIOCHEMISTRY,
UNIVERSITY OF NIGERIA, NSUKKA.
JULY, 2012.
UNIVERSITY OF NIGERIA, NSUKKA.
DEPARTMENT OF BIOCHEMISTRY.
15
PURIFICATION AND CHARACTERIZATION OF
LIPASE(EC.3.1.1.3) FROM THE SEEDS OF Cucumeropsis mannii
(WHITE MELON).
BY
EZEMA,BENJAMIN ONYEBUCHI
PG/M.Sc/09/51506.
BEING A PROJECT REPORT SUBMITTED TO THE SCHOOL
OF POSTGRADUATE STUDIES IN PARTIAL FULFILMENT OF
THE REQUIREMENTS FOR THE AWARD OF MASTER OF
SCIENCE (M.Sc) DEGREE IN BIOCHEMISTRY,
UNIVERSITY OF NIGERIA,NSUKKA.
SUPERVISOR: DR.S.O.O. EZE.
DEPARTMENT OF BIOCHEMISTRY,
UNIVERSITY OF NIGERIA,NSUKKA.
CERTIFICATION
Mr Ezema,Benjamin.O, Reg. No: PG/M.Sc/09/51506, a Postgraduate student of the
Department of Biochemistry, University of Nigeria, Nsukka, has satisfactorily
completed the requirements of research work for the Award of Degree of Master of
Science (M.Sc) in Biochemistry. The work incorporated in this dissertation is original
and had not been submitted in full or in part, for any other diploma or degree of this
or any other University.
……………………………… ………………………………
Dr.S.O.O. Eze Prof. L.U.S. Ezeanyika
(Supervisor) (Head of Department)
16
…………………………………………
External Examiner
LIST OF APPENDICES
Appendix.1: Protein determination. 63
Appendix.2: Absorbance values for protein standard curve. 64
Appendix.3: Protein standard curve . 65
Appendix.4: Method for preparation of fatty acid standard curve . 66
Appendix.5: Absorbance values for fatty acid standard curve. 67
Appendix.6: Fatty acid standard curve. 68
Appendix.7: Variation of lipase activity and protein concentration
during seed germination.
69
Appendix.8: Result of the preliminary study on ammonium
sulphate and cold acetone precipitation. 70
CHAPTER ONE
GENERAL INTRODUCTION
Lipases (triacylglycerol acylhydrolase EC.3.1.1.3.) are enzymes which
hydrolyze triacylglycerol to release free fatty acids and glycerol (Abdelmonaem et al.,
2011 ). They hydrolyze ester bonds of long chain aliphatic acids (fatty acids) from
glycerol at oil-water interface. Lipases are present in animals, plants and micro-
organisms (Ejedegba et al., 2007). Many biotechnological applications for lipases
have been described in food, detergent, oil and fat and pharmaceutical industries
(Barros et al., 2010). Plant lipases have attracted much interest in recent years as
biocatalyst, for the biotransformation of lipids (Hellyer et al., 1999). The storage
triacylglcycerol of oil rich seeds are hydrolyzed during germination of such seeds by
17
the action of endogenous lipases (Ivan et al., 1995). Cucumeropsis mannii, commonly
known as white melon seed is a member of the cucubitaceae family. The plant is a
species of melon native to tropical West-Africa where its cultivation is usually
associated to banana plant, corn and cassava (Fomekong et al., 2008). It is consumed
largely as thickener of traditional soup called egusi soup in Nigeria, Republic of
Benin and pistachio soup in Coted’Ivore (Koffi et al., 2008; Hanno and Susanne,
2010). The seed constitute about 44% oil (Badifu and Ogunsua, 1990). It therefore
represents a very good source of lipase. Despite its agronomic and cultural (traditional
medicine) importance, the plant lack attention from research and development so that
it is categorized as orphan crop (Loukou et al., 2007). The Limit of proper knowledge
of other possible utilization of the seed apart from consumption as food and in
traditional medicine is a major deterrent to its wider production, which should result
to increased income for the local farmers. Finding its use as a source of industrial
material (source lipase) would encourage its production and therefore improve the
local economy. In our quest for finding a cheap source of lipases we therefore report
the isolation, purification and characterization of lipases from the endosperm of
germinating seeds of Cucumeropsis mannii (White melon)
LITERATURE REVIEW
1.0 Brief History Of Enzyme:
The existence of catalysis in biology was first recognized as early as 1835 by Jons
Jacob Berzelius. He coined the word catalysis when he noted that potatoes contain
substances that catalyzed the breakdown of starch (Zubay et al., 1995).
Between 1850 – 1860, Louis Pasteur demonstrated that fermentation, the anaerobic
breakdown of sugar to CO2 and ethanol occurred in the presence of yeast cells
(Dubos, 1951). In 1877, a German physiologist. Wilhelm Kuhne first used the word
enzyme which he coined from Greek word evsuµov meaning in leaven to describe this
process (Kuhne, 1877). Later the word enzyme was used to refer to non living
substances such as pepsin and the word ferment was used to refer to chemical activity
18
produced by living organisms. In 1926, James B. Sumner showed that the enzyme
urease was a pure protein and crystallized it. John Northrop in 1930 isolated and
characterized a series of digestive enzymes, trypsin, chymotrypsin and pepsin. The
discovery that enzymes could function outside a living cell allowed their structure to
be resolved using x-ray crystallography. The first enzyme to be structurally resolved
was lysozyme, by a group pf scientists lead by David Chilton Philips (Blake et
al.,1965). This high structural resolution of lysozyme marked the beginning of the
field of structural biology and enzymology.
1.1 Meaning of Enzyme: In biology, one of the factors that define living things is
the organism’s ability to carryout chemical reactions that are crucial for its survival.
These reactions are controlled by the activity of enzymes. Enzymes are organic
substances that catalyze the repertoire of chemical reactions found in living things.
Like all catalysts, enzyme work by lowering the activation energy for a reaction, thus
increasing the rate of the reaction. However, enzyme differ from most catalysts by
being much more specific (Zubay et al., 1995).
1.2 Lipases: Lipases are known as triacylgcerol acylhydrolase, with the enzyme
commission number EC 3.1.1.3. Lipases catalyze the hydrolysis of various
forms of fatty acyl esters and needs oil- water interface for optimum activity
(Ejedegba et al., 2007). They catalyze the hydrolysis of ester-carboxylate
bonds releasing fatty acids and organic alcohol (glycerol) (Pereira et al.,
2003). The chemical equation bellow shows how lipase catalyzes the
hydrolysis of triacylglycerol to release free fatty acids and glycerol.
COOHOHHLipaseCCOCR RCII
21
1
1
1
2+−−−−−−
Equation.1: Hydrolysis of Triacylglycerol by Lipase.
C-CH2 – O- RC1
11
−
O
CH2OH R1COOH
CH2-O 11C R
3
O
O CH2-OH R
3COOH
+
Fatty acids
Glycerol
Triacylglycerol
19
However, in a water restricted environment, they catalyst the reverse reaction;
esterification or even transesterificaion and interesterification reactions (Castro, et al;
2000). The term transesterification refers to the exchange of groups between an ester
and an acid (acidolysis), between an ester and alcohol (alcoholysis) or between ester
(interesterification).
Lipases also display broad substrate specificity. Their specificities can be
further divided into three main groups: substrate specificity, regioselectivity and
enantioseletivity (Barros, et al; 2010).
Lipases are produced by animals, plant and microorganisms (Enujiugha,
2009). Many biotechnological applications of lipase have been described in food,
cosmetics, pharmaceutical and detergent industries (Barros, et al; 2010). These
potential applications have been the driving force in lipase research in the last few
years (Eze and Chilaka, 2010). Plant lipases are less studied when compared to
studies on those of micro organism and animal lipases (Bahri, 2000).
1.3 Structure of Lipase: Although lipases belong to many different protein
families, they have the same architecture. Ollis et al (1992) defines this
structure as the α\β – hydrolase fold. Generally lipase activity has been shown
to rely on triad usually formed by serine, histidine and aspartate residues
(Arpigny and Jaeger, 1999). In amino acid sequence of α\β hydrolases, the
three residues follow the order Ser-Asp His. Lipases also have consensus
sequence of Gly –Xaa-Ser-Xaa-Gly where X may be any amino acid residue
(Kanaya et al, 1998). According to Abdelmonaem et al (2011), the modeled
enzyme is a monomer folded into α\β domain consisting of eight central
stranded β-sheet flanked by twenty two α- helices. The number of α- helices
and β-sheets differ from one specie to another. The figure bellow shows the 3-
dimensional structure of lipase. The yellow and red coloured chains represent
the α and β-sheets respectively.
20
Figure.1: Three- dimensional structure of lipase (Adapted from Abdelmonaem
e t al, 2011).
1.4 Classification and Nomenclature: Lipases are varied in nature and may be
classified based on the substrate they catalyze, their optimum pH or their sources.
Based on the substrate they act on, they are classified as follows:
a. Acylglyceride lipase (glycerol ester hydrolase Ec 3.1.1.3) which hydrolyzes
acylglycerids.This work is centered on purification and characterization of this
class of lipase from germinating seeds of Cucumeropsis mannii. Among the
acylglyceride lipases are tri, di and mono – acylglyceride lipases (Gurr and
Harwood, 1991).
b. Lipoprotein lipase is a glycerol ester hydrolase which acts preferably on
triacylglycerides moiety of low density lipoproteins and very low density
lipoprotein (Verger and Abounsallam; 2000).
c. Phospholipases are interfacial enzyme which catalyzes the hydrolysis
of phospholipids. They are classified according to the position of their attacks
on the substrate into four groups namely; phospholipase A, B, C and D.
phospholipase A is divided into two, namely, phospholipase A. (EC 3.1.1.3)
which catalyzes the hydrolysis of the ester bonds in position 1 of the
phospholipids. Phospholipase A2 (EC 3.1.1.4) catalyzes the hydrolysis of ester
bounds in position 2 of the phospholipids forming lysophospholipid. (Gurr and
Harwood, 1991).
Phosphalipase B removes the remaining acyl group of the lysophospholipid
forming the corresponding glyceryl phosphoryl base. Phospholipase C (EC
3.1.4.3) hydrolyszs the ester bond in position 3 of phospholipids yielding 1, 2,
diacylglycerol and phosphoryl base.
Phospholipase D (EC.3.1.1.4.4) catalyzes the hydrolysis of the base
moiety from phospholipids.
21
Based on their optimum pH, lipases are classified as
a. Acid lipases which have their optimum activity in the acidic pH range.
b. Alkaline lipases have their optimum activity in the alkaline pH range.
According to sources, lipases are grouped as; bacterial, fungal, yeast, plant
and animal lipases.
1.5 Sources of Lipase: Lipases can be found in virtually every living thing. In
animals as pancreatic, hepatic, gastric and lipoprotein lipases; microbial as;
bacterial, fungal and yeast lipases and plant lipases. They could be intracellular or
extra cellular lipases (Vankampen et al; 1998). Lipases from different sources vary
in their catalytic properties (Barros, et al., 2010).This work dwelt on plant lipase
particularly seed lipase from the seeds Cucumeropsis mannii (White melon).
1.5.1 Seed Lipases: In recent times, seed lipases have been the focus of much
attention as biocatalysts (Barros et al., 2010). In most cases lipases from oil seeds
present advantages over animal and microbial lipases due to some quite interesting
features such as specificity, low cost, availability and ease of isolation. This makes the
lipases from seeds a great alternative for potential commercial exploitation as
industrial enzyme (Enujiugha et al., 2004, Paques and Macedo, 2006; Hellyer, et al,
1999). Lipase activity has been demonstrated in seeds and nuts of some plants such as
oil palm, coconuts, corn seedlings, conorphor nut, Jatropha. Curcus. (Ejedegba, et al.,
2007; Abigor, et al., 2002). Other plant lipases extensively studied include
glyoxysomal lipase of castor bean (wang and Huang, 1987) and Rice bran (Aizono et
al 1976).
Studies on plant lipases, despite its advantages have advanced slowly as
against microbial and animal lipases. This may be due to their solubility and diversity
(Aizono et al., 1976).Plant lipases may be classified into three major groups
(Abdelmonaem et al, 2011). The first group consists of the triacylglycerol hydrolase
that are primarily present in seeds. This group of plant lipases is of economic
important due to their implication in seed alteration during storage. The second group
of plant lipase is called acylhydrolase, which are present in various plant tissues.
These exhibit little specificity for their substrate and are able to hydrolyze
triglycerides but can catalyze some transesterification reactions (Hills and Mukherjee
1990). The main acylhydrolase are the phospholipases A and B, glycolipase,
22
sulfolipases and monoglyceride lipases. The third group of plant lipases is the
phospholipases C and D.
1.6 Role of seed lipases: In order to study seed lipases, one must understand their
physiological functions as well as their role in agricultural products during storage oil
hydrolysis in germinating seed. Seed germination is usually followed by a phase of
rapid growth as the seeding strives to establish a root system and achieve
photosynthetic competence. This growth is fueled by the mobilization of storage
reserves that were laid down during seed maturation (Bewley and Black, 1994). Oil in
the form of triacylglycerol is one the most common storage food compound is seeds
(Levin, 1974). They are stored in oil bodies surrounded by a phospholipid monolayer
(Huang, 1992; Murphy, 1993).
The initial step in oil break down is catalyzed by lipase (EC 3.1.1.3) which
hydrolyzes triacylglycerol (TAG) at the oil/water interface to yield free fatty acids and
glycerol. The free fatty acid are then transferred to the glyoxysome and activated to
acyl-CoAs for subsequent catabolism by β-oxidation. Most of the acetyl-CoAs
produced are then converted to sugars by the glyoxylate cycle and gluconeogenesis
(Peter, 2006). The fatty acid could also be converted to amino acids such as asparagin,
aspartate, glutamine and glutamate and provide carbon skeleton required for
embryonic growth (Quttier and Eastmond, 2009; Ejedegba, et al., 2007; Borek, et al.,
2006, Huang et al., 1988).
In most seeds, the activities of lipase are only detectable upon germination and
increase with disappearance of TAG. These lipase activities are often membrane
associated and can be found in the oil bodies, glyoxysome or microsomal fractions of
seed extracts ( Mukherjee, 1994). Lipolysis is an important control point in the overall
sequence of fat utilization in seeds especially during germination. This process is
under the control of lipases. The role of lipase during seed germination is summarized
in figure 2. Apart from mobilization of stored triacylglycerol during germination and
deterioration of stored seeds, lipases are also involved in other aspects of plant
metabolism such as rearrangement and degradation of chlorophyll during leaf growth
and senescence as well as in fruit ripening process (Tsuchiya et al, 1999).
Oil body
Triacylglycard Diacylglycard Manoacylglycrol
1 2 3
Glycerol
CYTOSOL
23
Figure 2: Chart showing how triacylglycerols (TAG) stored in the lipid bodies are
hydrolyzed to fatty acids (FA) and glycerol by the sequential action of one or more
lipases (Adapted from Quettier and Eastmond, 2009).
1.7 Specificity of seed lipases: With few exceptions, oilseed lipases are generally
more active with traicylgylycerol containing short chain fatty acids. Commonly used
substrates include commercially produced plant oils with unknown purity and
traicylglycerols with short chain fatty acids such as acetic and butyric acids, saturated
and unsaturated acylglycerols (Enujiugha et al, 2004). According to Hellyer et al
(1999), seed lipases show selectivity for the dominant fatty acids in the seed. For
example castor bean lipase shows selectivity for triricinolein; oil palm lipase for
tricaprion or trilaurein, elm lipase for tricaprion and vermonia sp lipase for
trivernolein. Other seed lipases can quickly hydrolyze a greater variety of fatty acids
such as canola and pinus seed lipases. Corn lipase, presented greater activity with the
triacylglycerol containing oleic and linoleic acid which are the predominant
constituent of corn oil (Lin, et al, 1986; Hammer and Murphy, 1993). With synthetic
substrates, lipases are found to present the same pattern they present with natural
substrates (Lin et al., 1986).
24
1.8 Oilseed lipases: Genuine lipases are those that hydrolyze fatty acids bonded to
their respective traicylglycerols (Barros et al, 2010). In vegetables, they are present in
oleoginous seeds (oil seeds) and other cereals. During the germination of oilseeds, the
lipid reserve is rapidly used up in the production of energy for embryonic growth.
During this period, lipolytic activities are usually very high and depending on the
plant species, the lipase may be located in the membrane of the lipid bodies or in
other cellular compartments such as the microsomes (Eze et al, 2005).Some widely
studied oilseed grains with respect to lipase extraction and characterization includes;
bean seed (Enujiugha et al, 2004); sunflower seed (Sagiroglu and Arabaci, 2005)
linseed (Sammaour, 2005); peanut (Huang and Moreau, 1978) and cotton seed
(Rakhimov et al, 1970).
Summary of the most widely studied seed lipases, their main physical and chemical
features and their application are shown in table 1.
Table 1: Some of the most studied seed lipases and their main features and
biochemical properties (Source: Barros et al, 2010).
Lipase Source Optimum
pH
Optimum
Temperature
Activator Inhibitor Substrate Specific
position
Application
African bean
seed (pentacle
thra
macrephylla
benth
7.0
30oC
Ca 2+
EDTA
Coconut
oil
-
Hydrolysis
Castor bean seed
(phasedus
vulgaris)
4.5 30oC Ca
2+ p-
chlorome
rcunbenz
oic acid
p-
nitrophenyl
butypate
Sn-1
Sn-2
Esterificaton
Rapeseed 7.0 37oC Bi
3+ Fe
2+, Olive oil - Esterifcation
25
(brassicanapus.L
)
Ca 2+
Fe2+
transesterific
ationcation
Barbados nut
(jatropha
curcas L)
7.5 37oC Ca
2+
Mg 2+
Fe2+
Olive oil - Hydrolysis
Lupin seed
(lupinus
lutens L)
5.0 45oC Ca
2+
Mg2+
K+
_ Lupin oil Sn-1
Sn-2
Hydrolysis
Almond seed
(amyadolus
communis L)
8.5 65oC Ca
2+,
fe2+
Co2+
Mn2+
and Ba2+
Mg2+,
Cu2+
and
Ni2+
Soybean
oil
_ Hydrolysis
Rice seed
(oryza sativa)
11.0 80oC - _ Olive oil Sn-2 Hydrolysis
Wheat seed
(Triticum
Oestivum)
8.0 37oC - _ Triolein _ Hydrolysis
esterificatio
n
Coconut seed
(cocos
nicifera linn
8.5 30-40oC _ - Olive oil Sn-1
Sn-3
Hydrolysis
1.9 Factors affecting lipase activity:
1.9.1 Moisture content: Lipase activity is inhibited by very low moisture content.
Lipases act at the water/oil interface and therefore require enough moisture build up
to work. Moisture content affects temperature and hence activity of lipase.
1.9.2 Temperature: Lipases are most active at temperature range of 30-40oC. The
optimum activity of some plant lipases are: Africa bean seed: 30oC, French bean
lipase 35oC, castor oil bean seed lipase 30
oC, rapeseed lipase, 37
oC and coconut lipase
40oC. Some seed lipases have high optimum temperature such as oat seed lipase 65-
75oC, rice seed lipase; 80
oC and almond seed lipase 65
oC (Barros et al, 2010).
26
1.9.3 Effect of pH: Lipases from different sources have different pH optimum for
their activity. Acid lipase has optimum pH within the acidic range while alkaline
lipases have optimum pH within the alkaline region. In general the optimum pH range
for most studied seed lipases is in the range of 4.5 – 9.0 (Barros et al, 2010).
1.9.4 Activators: A lot of factors have been shown to activate or improve lipase
activities in the reaction mixtures. Lipase catalyzed reactions can be increased by
providing optimum pH and optimum temperature. The degree of substrate emulsion is
also important in studying activation of lipase. The emulsifiers help to form oil
micelles thereby providing oil/water interface for lipases to act. Examples of such
emulsifiers are bile salt, cholic acid and deoxycholic acid. (Aizono et al, 1976). Some
metallic ions such as Na+, Ca
2+ K
+, NH
+4 and Ba
2+ have be shown to be strong
activator of lipases at low concentrations of 0.0IM. (Enujiugha, 2009, Barros et al,
2010). According to Tan et al (2003) Mg2+
, Na+ and K
+ are beneficial for synthesis of
lipases. Ca2+
form complexes with ionized fatty acids removing them from the oil-
water interface and thus enhancing lipase activity (Pancholy and Lynd, 1972).
1.9.5 Inhibitors of lipases: Enzyme inhibitors are compounds which combine with
the enzyme and inhibit or present the enzyme from carrying out its catalytic action.
For hydrolytic enzymes (lipases) heavy metals such as Cu2+
, Hg2+
, Zn2+
,Pb2+
, Fe2+
are
inhibitors (Barros et al, 2010).Oxidizing agents such as H2O2, atmospheric oxygen
and alloxan inhibit lipase probably due to their oxidation of the sulfyhydryl (SH)
groups of the amino acid side chain of the enzyme. EDTA may also inhibit lipase by
chelating divalent metal ions such as Mg2+
, Ca 2+
which aids lipase activity. Extremes
of pH and temperature away from the optimum values are all inhibitors of lipase
activity (Gracille, et al, 2000).
1.10 Applications of lipases: Lipases have become more and more prominent on the
enzyme biotechnology scenario due to their versatility for hydrolysis and synthesis
(esterification), their catalytic reaction often being chemo-selective, region-selective
or enantio-selective (Barros et al., 2010). Lipases are used in many sectors such as the
food, pharmaceutical, fine chemical, oil chemical (oleochemical), biodiesel and
detergent industries (Alonso et al., 2005).
27
1.10.1 Application of lipase in food industries: Lipases are employed in food
manufacturing to liberate fatty acids into food products by selective hydrolysis of the
fats and oil present in many kinds of food (Barros et al., 2010). Depending on the
carbon chain length and on the degree of unsaturation, the fatty acid obtained provides
the food with flavours, colours and characteristic smells playing an important role in
the physico-chemical and nutritional properties of many food products (Gandhi,
1997). Wheat, barley, corn and canola seed lipases have been employed in the
production of low molecular weight esters in an organic environment (Liaquat and
Apenten, 2000). These low molecular weight esters such as isopentyl butyrate,
butylcaproate, butyl acetate and ethylacetate confer characteristic aroma to foods. A 1,
3, specific lipase known as novozyme 677 has been greatly used in the bakery
industries to improve dough since 1995.
1.10.2 Application of lipases in detergent industries: Owing to their ability to
hydrolyze fats, lipases find a major use as additive in industrial laundry and house
hold detergent. The uses of lipase as functional compounds in the formulation of
detergents have been responsible for the sale of about 32% of the total lipase sale
every year (Sharma et al., 2001). The major properties of lipases exploited for this
purpose include their stability under washing conditions of pH between 10.0 and 11.0
and temperature between 30oC and 60
oC, resistance to other components of the
formulation such as protease and low substrate specificity (i.e. ability to hydrolyze
fats with various compositions) (Barros et al., 2010). Rice and oat seed lipases
possess suitable features for their use in detergent because they are stable at alkaline
pH and temperature of about 60oC.
1.10.3 Application of lipases in the oil and fat industries: Lipases show a wide
range of application in oleo-chemical industries. Their usage reduces expenses and
minimizes the large amount of heat involved in degrading compounds in comparison
to traditional chemical process (Freire and Castilho, 2008). Castor bean lipase has
been largely used as a biocatalyst in the esterification of fatty acids and glycerol. The
enzyme showed optimal efficiency in the formation of new tri, di and monoglycerols,
presenting great potential for the production of traicylglycerols of interest (Tuter,
1998).
28
1.10.4 Application lipase in fine chemical industries: The pharmaceutical and fine
chemical industries use lipase in their production processes. The enzyme’s regio,
enantio and chemoselectivity features allow their use for the resolution of recemic
mixtures. Lipases show excellent stability in the presence of organic solvent in which
the substrate are soluble (Jaeger and Eggert, 2002; Gotor-Fernadez et al., 2006). Xia
et al., (2009) used wheat germ lipase for the kinetic resolution of secondary alcohols.
1.10.5 Application of lipase in biodiesel production: The use of lipase in biodiesel
production has shown promising results in recent years. Using the enzymatic route,
the by-product glycerol can easily be removed without the requirement of a complex
separation process. In addition, oil free fatty acids, which can be used as the raw
materials are also converted completely into alkyl esters (Fukuda et al., 2001). Every
plant extract present lipolytic activity, which can be applied in the production of fatty
acids from triacylglycerol obtained from a variety of sources, considering that the
fatty material can be esterified by methanol or ethanol for biodiesel production
(Barros et al., 2010). Table 2 presents a summary of seed lipases and their
applications.
Table 2: Some seed lipases and their applications (adapted from Barros et al, 2010).
Source Application
1 Barley seed (Hordeum vulgare L) Production of low molecular weight esters.
2 Maize seed (Zea mays) Production of low molecular weight ester
3 Linseed (Linum usitatissi mum) Production of low molecular weight ester
4 Rape seed (Brassica napus L) Production of low molecular weight ester
and esterification
5 Black cumin seed (Nigella sativa L) Synthesis of structured lipids
6 Castor bean seed (Phaseolus vulgaris) Synthesis of structured lipids
7 Wheat germ Seed (Triticum sp) Esterification
29
8 Vernonia seed (Vernonia gala mensis) Hydrolysis of oil
1.11 CUCUMEROPSIS MANNII (WHITE MELON):
Cucumeropsis manni is a specie of melon native to tropical Africa west of the
Great Rift Valley, where it is grown for food and as a source of oil. Its common
names include ahu-ilu in Igbo, egusi in Yoruba and agushi in Hausa, in English it is
known as mann’s cucumeropsis and white –seed melon (Obute et al., 2008). It
produces climbing vines up to 4 meters long which are covered in stiff hairs. The
leaves which are heart-shaped or roughly palmate are up to 12 centimeters long and
14cm wide. It bears small yellow male and female flowers with petals about a
centimeter long. The fruit is egg-shaped or elongated ovate shape, up to about 19
centimeters long and 8cm wide. Its fruit has creamy colour with green streaks. Both
the fruits and seeds are edible. The plant is grown more often for the seed than the
fruit (Obute et al., 2008). Plates 1and 2 shows the Cucumeropsis mannii plant with
fruit and the processed seeds respectively.
1.11.1 Nutritional composition: Dehulled seeds from Cucumeropsis mannii
mainly consist of fats 44.4%, protein 36.1% and carbohydrate 13.2% (Badifu and
Ogunsua, 1991). Minerals and water amount to 3.7 and 5.9% respectively. From this
composition a caloric value of 2190 KJ/100g is calculated (Mbuli-Lingundi et al,
1983). The major component of the oil is linoleic acid and constitutes 57.9% of its oil
content. Short chain fatty acids are absent. All important macro and micro nutrients
are present it sufficient amount for human nutrition. Consumption of 100g dehulled
seeds covers the daily requirement of essential fatty acids, vitamin E and amino acids
(Mbuli- Lingundi et al; 1983 Abiodun and Adeleke, 2010).
1.11.2 Scientific classification of the plant:
Kingdom: Plantae – plants
Division: Magoliophyta – flowering plant
Class: Magnoliopsida – Dicotyledons
Order: Cucurbitales –
Family: Cucurbitaceue – Cucumber family
30
Genus: Cucumeropsis Naudin – Cucumeropsis
Specie: Cucumeropsis mannii Naudin – Mann’s cucumeropsis
Binomial name: Cucumeropsis mannii Naudin.
(http://zipcode200.com/plant/c/cucumeropsismannii.Retrieved 9/10/2010).
31
Plate 1: Photograph of the plant Cucumerepsis manni with fruits.
Plate 2: Photograph of the seeds of C. Mannii Naudin.
1.11.3 Production and Processing: Cucumeropsis mannii Naudin is regarded as the
original indigenous melon in West and Central Africa and the seed can be found in
most markets in the region. Although its production is declining, white melon is still a
common article in the markets. In West Africa, white melon is usually planted
between March and May and harvested 6-8 months later (September – December).
Fruits are collected when the stems have dried and fruits have changed colour from
green to creamy white or yellow. After collection, fruits are cracked or split open;
they are then placed in a heap or pit and are left for 14-20 days to let the fruit pulp rot.
The seeds are removed and thoroughly washed to remove thick mucilage covering
32
them. The seeds are dried to 10% moisture content before packing (Zoro-Bi et al.,
2003).
1.11.4 Uses of Cucumeropsis manni naudin: Cucumeropsis mannii is mainly grown
for its oily seed. The seeds are sold almost three times more expensive than cocoa and
about seven times more expensive than coffee (Zoro-Bi et al., 2003). The kernels are
milled into a whitish paste which is consumed as thickeners of a traditional soup
called egusi soup in Nigeria Cameroon and Benin and pistachio soup in Cote d’Ivoire
(Loukou et al., 2007 and Zero-Bi et al, 2003; Koffi et al., 2008).The seeds of C.
Mannii are used to prepare dough or a sauce and soup flavour when fermented (ogiri)
(Fomekong et al., 2008) It has an important value in the African traditional society
(Ponka et al., 2005). The flesh of the fruit, though edible is not commonly eaten. In
Ghana the fruit juice mixed with other ingredients is applied to the navel of new born
babies to accelerate the healing process until the cord-relics drops off. Macerated
leaves are used in Gabon for purging constipated suckling babies. In Sierra Leone
cattle boys traditionally use the dried fruit-shell of C. manni as a warning horn.
(Hanno and Susanne, 2010). Despite its agronomic, cultural and nutritional
importance,the plant lack attention from research and development so that it is
categorized under the orphan crops of Africa (Chweya and Ezaguirre, 1999).
1.12 AIM OF RESEARCH:
White melon seeds are consumed widely in West Africa and especially in Nigeria
where they are used in traditional medicine and in making soups when ground. This
research is aimed at isolating lipase from germinating seeds of Cucumeropsis mannii,
purifying the enzyme and studying its basic kinetic properties. Owing to the fact that
lipases have wide range of industrial and biotechnological applications, successful
isolation and characterization of lipase from locally available oil seeds such as white
seed melon might enhance the production of oleochemicals and also help to improve
the economy of the local farmers who are the main producers.
33
1.13 Objective of study: The specific objective of this study includes;
(a) To isolate lipase from germinating seeds of Cucumeropsis mannii (white melon).
(b) To purify the lipase.
(c) To characterize the purified lipase.
34
CHAPTER TWO
MATERIALS AND METHODS
2.0 Materials
2.1 Plant material: Matured and freshly processed seeds of Cucumeropsis mannii
were purchased from the local market in Nsukka, Enugu State, Nigeria.
2.2 Chemicals :
BSA (Bovin serum albumin) and Sephadex G-200 were obtained from BDH,
England. Other reagents and solvents used were of analytical grade. All laboratory
reagents were prepared fresh.
2.3 Equipments:
� Centrifuge model 800.
� Digital electronic weighing balance( mettler Toledo B 204-5).
� Glass column (62x2.5cm).
� Magnetic stirrer; model AM-3250B.
� Pestle and mortar.
� pH meter; model PHS-3C.
� Spectrophotometer ( Jenway 6405 UV/Vis).
� Water bath; model DK.
� Other laboratory materials such as test tubes, pipette, beakers were used where
and when appropriate.
2.4 METHODS.
2.4.1 Reagent Preparation:
1. Extraction buffers.
(a) Tris-HCl buffer: 150mM Tris-HCl buffer (PH 7.5) consisting of 0.4M
sucrose, 2.0mM beta-mercaptoethanol and 0.5mM EDTA was prepared by
dissolving the following calculated quantities of compounds in about 700ml
of distilled water in a 1000ml beaker. Tris-base (18.17g), sucrose (136.92g),
EDTA (0.146g) and 0.14cm3 of beta-mercapthoethanol.
The beaker and its content were placed on a magnetic stirrer and the
pH adjusted to 7.5 using dilute HCl (0.01). The volume was made up to
1000ml with distilled water. It was then stored in the refrigerator.
35
(b) Phosphate buffer:150mM Sodium phosphate buffer (pH 5.9) consisting of
0.4m sucrose, 2.0mM Beta-mercaptoethanol and 0.5mM EDTA was prepared
by dissolving the following calculated quantities of compounds in about
700ml of distilled water in 1000ml beaker. Na2HPO4 (21.297g) conjugate
base, NaH2PO4.H2O (20.702g) conjugate acid, sucrose (136.92g), EDTA
(0.146) and 0.14Cm3 of beta-mercaptoethanol.
The beaker and its content were placed on a magnetic stirrer and the
pH adjusted to 5.9 using dilute HCl (0.01N). The volume was then made up to
1000ml using distilled water. It was then stored in the fridge.
(c) Assay Buffers: The buffers used in assaying for lipase activities in this
experiment (Tris HCl pH 7.5 and phosphate buffer, pH 5.9) were prepared as
described above with the exclusion of sucrose. The assay buffers therefore
contained the respective conjugate acid and conjugate base, 0.5mM EDTA and
2.0mM beta- mercaptoethanol.
2. Stock Solution of standard fatty acid (25mM oleic acid) used for preparation
of fatty acids standard curve was prepared by dissolving 0.75 ml of oleic acid
in 70ml of chloroform and the volume made up to 100ml using chloroform.
From this stock, a concentration range of 0.5 – 5.0mM was prepared in
chloroform by serial dilution and their corresponding absorbancies at 540mm
measured after addition of 0.5ml sodiumdiethyldithiocarbamate. The obtained
values was used to prepare a standard curve. The principle of this reaction
involves the reaction between copper(ii)ion and the fatty acid to form copper
soap. The copper soap then form a yellow complex with
sodiumdiethyldithiocarbamate. The intensity of colour produced is
proportional to the concentration of free fatty acid in solution.
3. Stock solution of standard protein (BSA) (concentration; 0.5mg/ml) was
prepared by dissolving 0.05g of BSA in 100ml of distilled water. From this
stock solution, different protein concentration ranging between 0.05mg/ml-
0.5mg/ml was prepared by serial dilution and their absorbance at 750nm was
measured using spectrophotometer based on Lowry method of protein
estimation. The absorbance plotted against concentration was used to prepare
the protein standard curve.
36
4. 1.0M aqueous triethanolamine was prepared by dissolving 13.5 ml of
triethanolamine in small quantity of distilled water and the volume made up to
100ml using distilled water.
5. 6.45% (w/v) Cu (NO3) 2 3H20 was prepared by dissolving 6.45g of Cu (NO3)2.
3H2O in 100ml of distilled water.
6. 0.1% (w/v) sodium diethyldithiocarbamate was prepared by dissolving 0.1g of
sodiumdiethyldithiocarbamate in butan-1-ol and the volume made up to 100ml
using butan-1-ol.
7. 0.1N NaOH was prepared by dissolving 0.4g of NaOH pellet in 100ml of
distilled water.
8. 2% alkaline Na2CO3 solution was prepared by dissolving 2.0g of Na2CO3
in 100ml of 0.IN NaOH
9. Copper Sulphate- sodium- potassium tatrate solution was prepared by
dissolving 1g of sodium – potassium tatrate and 0.5g of CuS04.5H2O in 100ml
of distilled water.
2.4.2 Seed Germination.
Seeds of Cucumeropsis manii (white melon) were soaked in tap water for 48hr
with intermittent change of water every 6 hr. The end of inhibition period was
designated day zero of germination. Germination was carried out in moist jut bag in a
dark cupboard at room temperature (Eze and chilaka 2010). After 4 days the
endosperms were removed and used for enzyme isolation/extraction. Plate 3 shows
Cucumeropsis mannii seeds at different stages germination (in days)
2.5.0 Germination parameters.
2.5.1 Germination energy.
Fifty seeds of Cucumeropsis mannii were each placed in a petri-dish
containing two filter papers. Four milliliters of distilled water was added into the dish
and the dish covered and left in the dark for 96hours. The number of seeds that
germinated was counted at 24hours intervals and the values expressed as percentage
of the total number of seeds in the dish.
37
2.5.2 Water sensitivity.
Fifty seeds of Cucumeropsis mannii were placed in each of four petri-dishes,
containing two filter papers. To two of the petri-dishes, 4.0ml of water was added and
8.0ml to each of the remaining two. The petri-dishes were then covered and kept in
the dark for 96hours. The number of C.maninii seeds in each petri-dish that
germinated after 96hours was counted and recorded as a percentage of the total
number of seeds in each petr-dish (the germination energy). The difference between
the germination energy found in the 4.0ml and 8.0ml water sample was used as the
water sensitivity value for the seed.
2.5.3 Average root length.
Seeds of Cucumeropsis mannii (50) were germinated for four days and the
length of each root was measured and recorded. The average length was then
calculated as follow.
Average Root Length = Sum of the root lengths ÷the number of seeds.
38
Germination after 1 day
Germination after 2 days
Germination after 3 days Germination after 4 days
Plate.3: Photographs of seeds of Cucumeropsis mannii at different stages of
germination.
39
2.6.0 Lipase Extraction: This was carried out using the method of Eze et al (2005),
with little modification. The endosperm tissues of four day germinated seeds
of Cucumeropsis mannii were carefully removed and washed in distilled
water. The endosperms were ground using mortar and pestle in a cold grinding
medium containing 0.4M sucrose, 0.5mM EDTA, 2.0mM beta-
mercaptoethanol, 1.95% (w/v) Tween 80 in 150mM Tris- HCl buffer pH 7.5
on one hand and 150mM phosphate buffer pH 5.9 on another portion of the
endosperms. The homogenates was filtered through four layers of cheese
cloth. The filtrates were centrifuged at 5000xg for 30min at 40C. The fat layer
(lipid pad) was removed using spatula, the supernatant (water soluble portion)
and the pellet collected into separate containers. Lipase activity was assayed
for in the three fractions.
2.7.0 Preparation of substrate emulsion (oil emulsions):Ten grammes each of
gum Arabic and olive oil were combined in a 400ml beaker. The volume was
brought to 200ml with appropriate buffer (150mM Tris-HCl pH 7.5 or 150mM
phosphate buffer pH 5.9) and homogenized for 5 minutes at low speed in other
to avoid excessive foaming. The emulsion formed was continuously subjected
to low speed magnetic stirring during the entire day of use.
2.8 Lipase Assay: A modified colorimetric method of Duncombe (1963) was
used. The free fatty acids released by lipase were converted to copper soaps and
quantified using sodiumdiethyldithiocarbamate as a colour reagent. The reaction
mixture contained 0.5 ml of assay buffer, 1.0 ml of enzyme solution pre incubated
for 5.0 minutes. Then 5.0ml of substrate already emulsified as described above was
added and the mixture incubated at 370C in a water bath with constant shaking for 30
minutes. At the end of incubation period, 1.0 ml of 6.0N HCl was added to terminate
the reaction and enhance extractability of the fatty acid released by the action of
lipase. The mixture was allowed to stand for 5.0 minutes, then 5.0 ml of chloroform
was added and the test tube was shaken vigorously. 2.5 ml of copper reagent
(consisting of 9 volume of I.0 M triethanolamine, 1 volume of I.0N acetic acid and
10 volume of 6.45% (w/v) Cu(NO3)2.3H20) was added and the mixture shaken
vigorously for 2 minutes. The test tube containing the above mixture was centrifuged
at 5000хg for five minutes to separate the phases clearly. 3.0 ml of the lower
40
chloroform layer was transferred into a clean dry test tube with cap and 0.5 ml of
sodium-diethyldithiocarbamate (0.1% w/v in butanol) was added. The absorbance of
the corresponding yellow colour developed was measure at 440 nm against a blank.
The concentration of the free fatty acid (FFA) released was determined from a
standard curve of fatty acid.
Lipase activity = Fxtimeincubation
umlibratedacidfarryofionconcentrat
(min)
)(
Where F = dilution factor, where necessary.
2.8.1 Measurement of lipase activity during the period of seed germination:
The seeds of Cucumeropsis mannii (white melon) were washed in distilled
water containing 0.01% HgCl to prevent fungal growth. The seeds were then soaked
in tap water for 48 hours. The end of inbibition period was designated day zero of
germination. Germination was carried out in a moist jute bag in a dark cupboard at
room temperature for a period of 7 days. At the end of every 24 hours, starting from
day one to day seven, 50 seeds were harvested and the endosperm tissues carefully
removed. The endosperms were used for enzyme isolation according to the method of
Eze et al, (2005). The endosperm tissues were washed in distilled water and then
ground with mortar and pestle in 50ml of grinding medium containing 0.4 M sucrose,
0.5 mM EDTA, and 2.0 mM beta-mercaptoethanol in 150 mM Tris-HCl buffer pH
7.5. The homogenate was filtered through four layers of cheese cloth and washed with
about 5 ml of the grinding medium to a total volume of 55 ml. The filtrate was
centrifuged at 5000 x g for 30min at 4oC. The top lipid body was removed using
spatula. The supernatant and pellet were collected in separate containers. The
supernatant was regarded and used as the crude lipase. Lipase activity was assayed in
each crude extract isolate at the end of every 24 hours, throughout the period of seven
days. Protein was also determined by the method of Lowry et al (1951).
2.8.2 Localization of Cucumeropsis mannii Lipase: The endosperm tissues of
four days germinated Cucumeropsis mannii seeds were ground with mortar and pestle
in grinding medium as described in section 2.6.0. The homogenate was filtered
through four layers of cheese cloth. The filtrate was centrifuged at 5000 x g for 30min
at 4oC to yield three fractions: the fat pad (lipid layer), water soluble fraction
(supernatant) and sediment (pellet).
41
The fatty layer was resuspendend in buffer and designated the lipid bodies. The
sediment earlier obtained was resuspended in a small volume of the grinding medium
and centrifuged again. The resulting pellet was designated the particulate fraction
(pellet) while the water soluble fraction was pooled and combined with the
supernatant obtained above. The pellet was redissovled in buffer and used as the pellet
fraction. Lipase activity was carried out in the three fractions: lipid bodies,
supernatant and pellet using the Duncombe (1963) assay method as described section
2.8. Protein was also determined using lowry method.
2.8.3 Solubilization of membrane bound Cucumeropsis manni lipase: Some four
days germinated seeds of Cucumeropsis manni were selected and the endosperms
carefully remove. The endosperms were washed in distilled water and lipase isolated
from them using the method described in section 2.8.2. The lipid body fraction was
resuspended in 150mM Tris-HCl buffer pH 7.5. Two detergents were selected: tween-
80 and triton X-100. To equal volume of the lipid body fractions these detergents
were added in separate test tube from concentration range of 0.65 to 3.9% (w/v).
Higher concentration of 5 and 10% were also prepared. The mixture was stirred at
4oC for 30 min and then centrifuged at 5000g for 30min to remove all unsolubilized
materials. Lipase activity was assayed for in the supernatant using Doncumbe’s
(1963) method as described 2.8. Plate 4 shows the visual clarity of tween-80 and
triton X-100 at different concentrations.
42
Plate.4: The clarity of lipid body during solublization at different detergent
concentration.
Different concentration
of Triton X-100, (0,5
and 10%w/v Different
concentrations of
Tween80 (0, 5 and
10%w/v)
43
2.9 Purification of lipase from Cucumeropsis manni Seed:
The procedure adopted for purification of cucumeropsis lipase consisted of
the following steps.
2,9.1 Isolation of lipase: Cucumeropsis mannii lipase was isolated using the method
of Eze et al (2005) with modifications as described in section 2.6.0 . The supernatant
obtained after centrifugation was used as the crude lipase.
2.9.2 Solubilisation of Cucumeropsis lipase: In other to solubilize the membrane bound
lipase, 1.95% (w/v) concentration of tween-80 was incorporated into the homogenization
buffer. The homogenate was stirred for 30min and then filtered through four layers of
cheese cloth. The filtrate was centrifuged at 5000xg for 30min at 40C and the
unsolubilized lipid bodied was removed with spatula and the supernatant collected and
used as the crude enzyme. Lipase activity was assayed using the method of Duncombe
(1963).
2.9.3 Precipitation: The crude lipase fraction obtained from the isolation and
solubilization steps was divided into two equal parts (volume). One portion was
subjected to ammonium sulphate precipitation while the second portion was
subjected to cold acetone precipitation. The precipitates obtained from both (ie
ammonium sulphate saturation and cold acetone precipitation) were combined and
subjected to dialysis.
2.9.4 Ammonium Sulphate (NH4)2SO4) Precipitation: The crude lipase was
subjected to 70% ammonium sulphate saturation by adding solid ammonium
sulphate to the solution of enzyme in a 1000 ml beaker slowly over a period (one
hour). the solution was continuously being stirred slowly to help in dissolution of the
(NH4)2SO4) while the beaker was immersed in an ice bath. The solution was kept
undisturbed in a fridge at about 4oC for 36 hours. The fatty upper part was carefully
removed and the precipitates formed in the clear down layer were collected after
centrifugation at 5000xg for 20min. The precipitates obtained was redissolved in
buffer to a total volume of 90 ml. Enzyme activity was determined using Doncumbe
(1963) method as earlier described 2.8. Protein concentration was also determined
using Lowry method (1951).
44
2.9.5 Acetone Precipitation: The second portion of the crude lipase was gradually
made up to 50% (v/v) acetone saturation by adding cold acetone slowly to the enzyme
solution in a 1000ml beaker immersed in an ice bath, to prevent excessive heat which
may denature the enzyme. The solution (mixture) was allowed to stand undisturbed in
a refrigerator for 24hrs. At the end of the 24 hours, the upper acetone layer which
contained the precipitates was collected and centrifuge at 5000xg for 15min. The
precipitate were air dried to evaporate the acetone and then the acetone free
precipitate redisolved in Tris HCL buffer (pH:7.5) to a total volume of 90 ml. Lipase
activity and protein concentration were determined.
2.9.6 Dialysis: The lipase fractions obtained from the two separate precipitations
(ammonium sulphate and acetone) were combined and the resultant solution dialysed
using a dialysis bag in 0.15M phosphate buffer pH 5.9 for 24 hours with changes of
buffer every 6 hours.
2.9.7 Gel-filtration column chromatography: 7.5 ml of the dialysate obtained from
the dialysis above was loaded unto a gel filtration column (2.5x62 cm) of sephadex G-
200. The column was continuously packed with the gel to a height of
62 cm. It was washed with distilled water and then equilibrated with 0.15M
phosphate buffer pH 5.9. The enzyme was eluted with the same buffer. The eluent
was fractionally collected in test tubes at a flow rate of 2.2 ml per hour.
2.10.0 Characterization of Cucumeropsis mannii lipase: The purified lipase was
characterized with respect to pH, pH stability, temperature, heat stability, effect of
some selected metal ions and effect of substrate concentration.
2.10.1 Determination of effect of pH on the activity and stability of C. mannii
lipase: The activity of lipase was determined within the pH range of 3.0 - 9.0 using
the following buffer systems: Sodiumacetate buffer (0.15M, pH 3.0 – 4.5); sodium
phosphate buffer (0.15M, pH 5.0 – 6.5); Tris – HCl buffer (0.15M, pH 7.0 – 9.0), with
olive oil as substrate. The effect of pH on lipase stability was determined by
incubating 0.5ml of the lipase fraction in various buffer solutions (as above) ranging
from (3.0 – 9.0) for one hour at 37oC in a water bath. The residual activity was then
determined using the standard Duncombe’s (1963) method.
45
2.10.2 Determination of effect of temperature on C. manni lipase activity and
stability: The temperature dependence of lipase activity was measured by incubating
fractions of the enzyme at temperatures 30oC – 70
o for 30 minutes and activity
determined. For temperature stability, enzyme solution was incubated in a water bath
at temperatures 30 – 800C for one hour. At the end of the incubation period, the
enzyme solution was rapidly cooled at 0oC for another 1 hour. The residual activity
was determined using the Duncombe’s (1963) method.
2.10.3 Determination of effect of metal ions and EDTA on the activity of
C. mannii lipase: To determine the effect of metal ions on lipase activity, 0.5ml of
the enzyme were incubated with equal volumes of the various solutions containing
metal ions of concentration 1.0 mM at 370C for 30 min in a water bath. A control
containing equal volume of distilled water in place of metal ion was also incubated.
The relative activity of lipase after incubation period was determined using the
method of Duncombe (1963). The relative activity was expressed as the percentage of
the activity obtained in the control.
46
CHAPTER THREE
RESULTS
3.0. RESULTS
3.1 Germination parameters.
Studies on the germination properties of coated and decoated seeds of C-manii
showed that the coated seeds have germination energy, water sensitivity and average
root length of 82, 18 and 4.28 cm respectively compared to 68, 20 and 3.46cm for
decoated seeds. This suggests that the coated seed of C.mannii performed better in
terms of endosperm modification; hence it was used for subsequent experiment.
3.2 Variation of lipase activity during germination. Lipase activity increased
during germination from day zero to a peak on the 4th
day of germination after which
it decreased progressively to the 7th
day. Figure 3 shows the pattern of changes in
lipase activity during germination. A different pattern was obtained for the changes in
the protein concentration during germination. The protein content of the seeds
increased gradually from day zero to a maximum on the 3rd
day and decreased until
the 7th
day of germination. Figure 4 shows the changes in protein concentration during
germination of Cucumeropsis mannii seeds. The changes in lipase activity and protein
concentration in coated and uncoated seeds of C. mannii were compared. The result
obtained showed that the coated seeds had more lipase and protein concentration on
the peak days when compared to the uncoated seeds. The results are presented in
figures 4and 5.
0
2
4
6
8
10
12
0 1
Sp
eci
fic
act
ivit
y (
µm
ol/
min
/mg
pro
tein
)
Figure 3: Variation of lipase activity during seed
0
2
4
6
8
10
12
14
0 1
Pr0
tein
co
nce
ntr
ati
on
(mg
/ml)
Duration of germination (Days)
Figure.4:Changes in protein concentration during
1 2 3 4 5 6 7
duration of germination (Days)
Figure 3: Variation of lipase activity during seed
germination
1 2 3 4 5 6
Duration of germination (Days)
Figure.4:Changes in protein concentration during
germination
47
7
0
20
40
60
80
100
120
140
0 1
Act
icv
ity
(µ
mo
l/m
in)
Figure.5:Compariso of thevariations in lipase activity during
seed germination in coated and uncoated seeds
0
2
4
6
8
10
12
14
16
18
0 1
pro
tein
co
nce
ntr
ati
on
(m
g/m
l)
Figure.6: comparison of the changes in protein concentration
during germination in coated and uncoated seeds
2 3 4 5 6 7
Days
Figure.5:Compariso of thevariations in lipase activity during
seed germination in coated and uncoated seeds
Decoated
coated
2 3 4 5 6 7
Days
Figure.6: comparison of the changes in protein concentration
during germination in coated and uncoated seeds
Decoated
coated
48
Figure.5:Compariso of thevariations in lipase activity during
Figure.6: comparison of the changes in protein concentration
49
3.3 Localization of Cucumeropsis manni Lipase. When the endosperm of the 4 days
germinated seeds of Cucumeropsis mannii was ground, filtered and the filtrate
separated into 3 different fractions:Lipid bodies (fat pad), water soluble fraction
(supernatant) and pellets (particular fraction) through centrifugation, lipase
activity was found in the three fractions. The activity being highest in the lipid
bodies, followed by the water soluble fraction and least in the pellet (Figure 5).
3.4 Solubilization of Cucumeropsis mannii Lipase. The high activity of lipase found
in the lipid body fraction after separation (figure 6) shows that Cucumeropsis
mannii lipase is a lipid membrane bound enzyme. In order to liberate the
membrane bound lipase the lipid body fraction was solubilized using two selected
detergents; Tween 80 and Triton X-100 as described in section 2.8.3.Result
obtained showed that Tween 80 is a better solubilizing agent than Triton X-100.
The best solubilization was achieved at 1.95% concentration of Tween 80. This
percentage concentration (1.95%) (w/v) of Tween 80 was incorporated into the
extraction buffer used during the mass extraction and purification of the lipase in
order to solublize the lipid bodies. Figure 6 shows the result of solubilization of
the lipid bodies using varying concentrations of Tween 80 and Triton X-100.
pH profile of the crude enzyme showed that the enzyme had two pH optima, one
on the acidic side (pH 5.9) and the other on the alkaline pH 7.5 Result shown in
figure 6.
50
0
20
40
60
80
100
120
3.7 4.7 5.7 6.7 7.7 8.7 9.7
Act
ivit
y (
µm
ol/
min
)
pH
Figure.7: pH profile of crude lipase from germinated white melon
seeds (Cucumeropsis mannii)
0
20
40
60
80
100
120
140
160
180
Lipid body
Lip
ase
act
ivit
y (
µm
ol/
min
)
Figure.8:Localization of lipase activity in the 3 fractions
0
100
200
300
400
500
600
0 2
Lip
ase
act
icv
ity
(µ
mo
l/m
in)
Figure.9:Result of Solubulization of lipase using two
Lipid body supernatant Pellet
Fractions
Figure.8:Localization of lipase activity in the 3 fractions
4 6 8 10 12Detergent concentration (%)
Figure.9:Result of Solubulization of lipase using two
detergents
Tween-80
TitonX-100
51
52
3.5 Purification of lipase from Cucumeropsis mannii:
Water soluble fraction of the homogenates from the 4 day germinated seeds of
Cucumeropsis mannii were extracted in two separate buffers containing 1.95%
(w/v) of Tween-80, Tris-HCl, pH 7.5 and phosphate buffer, pH, 5.9 respectively
as described in section 2.5.3. Maximum enzyme activity was obtained in the
precipitates formed at 70% (NH4)2SO4 saturation and 50% cold acetone saturation
on the other hand. Combining the precipitates from 70% (NH4)2SO4 and 50% cold
acetone for each buffer system yielded enzyme solution with more activity than
the one obtained from individual precipitation procedures. The combined fractions
were then dialysed overnight resulting to a slight increase in the lipolytic activities
of the two fractions.
When equal volumes of the dialyzed fractions were combined and run
on sephadexG-200 gel column chromatography, no clear peaks were
obtained.Thus peaks showing relatively high lipase activity were pooled
and subjected to Sephadex G-200 gel filtration column chromatography
again.Two peaks showing lipase activity were obtained. The first peak
was designated A and the second peak B .Fractions from the two peaks
were pooled into separate containers designated lipase A and Lipase B and
were characterized separately as such, pH profile of the fractions showed
that lipase A has optimum pH of 7.5 while lipase B has optimum pH 5.9.
The result of purification is summarized in table 3 which shows increase in
the specific activity of enzyme as purification progresses. This shows that
the enzyme is becoming purer as the purification proceeds.
53
0
10
20
30
40
50
60
70
20 30 40 50 60 70 80
Act
ivit
y (
µm
ol/
min
/)
% Ammonium sulphate
Figure.10: preliminary study for ammonium sulphate
precipitation
0
10
20
30
40
50
60
70
20 30 40 50 60 70 80
Act
ivit
y (
µm
ol/
/min
)
Cold acetone (%)
Figure.11:preliminary study for acetone precipitation
54
0
0.2
0.4
0.6
0.8
1
1.2
1.4
1.6
0 10 20 30 40 50
Ab
sorb
an
ce (
nm
)
Tube Numbers (0-50)
Figure.12: Chromatogram for protein absorbance at 280nm and
lipase activity against tube numbers for the first gel filtration
Absorbance at
280
Absorbance at
440
-0.1
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
0 10 20 30 40 50
Ab
sorb
an
ce(2
80
nm
)
Tube Numbers (0-50)
Figure.13: Chromatogram for protein absorbance at 280nm and
lipase activity against tube numbers for the second gel filtration
ProtienAbsorbance
at 280
Lipase ActivityPeak A
Peak B
55
Table 3: Purification table.
Purification
Step
Volume
(ml)
Protein
(mg/ml)
Total
protein
(mg/ml)
Enzyme
activity
(µm/min)
Total
activity
(µm/min)
Specific
activity
(µm/min/mg
prot)
Yield
(%)
Purifi-
cation
(fold)
Crude Extract 400
88.8 35,520 67.37 26,948 0.76 100 1.0
Combined
(NH4)2SO4 and
Acetone
precipitation
30 81.5 2,445 71.67 2150.1 0.79 8.0 1.04
Dialysed
37 67.4 2493.8 131.1 4850.7 2.0 18.0 3.0
Ch
rom
ato
gra
ph
y
in
sep
ha
dex
G-2
00
Lip
ase
A
7.2
6.4
46.15
33.40
240.5
5.2
0.7
5.4
Lip
ase
B
9.4
6.3
59.22
50.60
475.6
8.03
1.76
11.0
56
3.6 Effect of pH on the activity and stability of lipase from C. mannii:
Two pH optima were obtained for the different fractions. Lipase A showed
optimum pH of 7.5 while lipase B showed optimum pH of 5.9. This could indicate
that the two fractions contained different forms of lipases; lipase A is alkaline
lipase while lipase B is acid lipase. It was observed that lipase A was stable
between pH 6.5 and 8.0 while lipase B was stable between pH 4.5 and 6.0.
3.7 Effect of temperature on the activity and stability of Cucumeropsis mannii
lipase:
The two lipase fractions (lipase A and B) had optimum temperature of 370C and
were found to be stable at 450C for 1 hour. Their stability decreased progressively
from 500C. Activity was detectable at higher temperature of 80
0C showing that the
enzyme is heat stable.
57
0
10
20
30
40
50
60
70
80
0 2 4 6 8 10
Sp
eci
fic
act
ivit
y(U
/mg
pro
tein
)
pH Values
Figure .15:pH Optimum for lipase B
0
10
20
30
40
50
60
70
80
0 2 4 6 8 10
Sp
eci
fic
act
ivit
y(U
/mg
pro
tein
)
pH Values
Fig.14: pH Optimum for lipase A
58
0
5
10
15
20
25
30
35
40
0 2 4 6 8 10
Sp
eci
fic
act
ivit
y(U
/mg
pro
tein
)
pH Values
Figure.16:pH Stability of lipase A
0
5
10
15
20
25
30
35
40
45
50
0 2 4 6 8 10
Sp
eci
fic
act
ivit
y(U
/mg
pto
tein
)
pH Values
Figure.17:pH Stability for lipase B
59
0
5
10
15
20
25
30
35
40
45
50
1 11 21 31 41 51 61 71 81
Sp
eci
fic
act
ivit
y(u
/mg
pro
tein
)
Temperature
Figure.18:Temperature optimum for lipase A
0
10
20
30
40
50
60
0 20 40 60 80
Sp
eci
fic
act
ivit
y (
U/m
g p
rote
in)
Temperature (oC)
Figure.19: Temperature optimum for lipase B
60
0
5
10
15
20
25
30
35
40
45
50
30 40 50 60 70 80 90
Sp
eci
fic
Act
ivit
y(U
/mg
pro
tein
)
Temperature
Figure.20:Temperature Stability of lipase A
0
5
10
15
20
25
30
35
40
45
50
30 40 50 60 70 80 90
Sp
eci
fic
Act
ivit
y (
U/m
g p
rote
in)
Temperature
Figure.21: Temperature Stability of lipase B
61
3.8 Effect of Metal Ion: In this study, the residual activity of the two enzymes
observed in the different metal ions followed a close pattern. Ca2+
proved to be the
best activator of the two lipases, increasing lipase activity by 150.7% followed by
Zn2+
(111.4%). Mg2+
(68%) and Na+ (77%) caused slight deactivation while Pb
2+
strongly inhibited activity in the two enzyme fractions by about 80%. EDTA, a
chelating agent showed no significant effect on activity of these enzymes. This could
suggest that these lipases may not be metalloenzymes and that the slight decrease in
activity observed with the EDTA may be due to the interference with absorption on
the substrate (oil) water interface. Figure 17 and 18 shows the effect of metal ions on
the activity of lipases from Cucumeropsis mannii.
62
.
0
20
40
60
80
100
120
140
160
Control Ca2+ Zn2+ Na+ Mg2+ AL3+ Pb2+ EDTA
Re
sid
ua
l a
ctit
vit
y (
%)
Metal ions
Figure 22: Effect of metal ions on lipase A
0
20
40
60
80
100
120
140
160
Control Ca2+ Zn2+ Na+ Mg2+ AL3+ Pb2+ EDTA
Re
sid
ua
l a
ctiv
ity
(%
)
Metal ions
Figure 23: Effect of metal ions on lipase B
63
3.9 Kinetic studies: Figure 19 and 20 shows the double reciprocal plot (Lineweaver –
Burk Plot) of the effect of substrate (olive oil) concentration on lipase A (alkaline
lipase) and lipase B (acid lipase) from Cucumeropsis mannii. From the graphs, the
Vmax and Km values of the two lipases were calculated. Lipase A was found to have
Vmax value of 142.85unit and Km value of 13.28g/L (0.01328mg/ml), while lipase B
was found to have Vmax value of 166.67unit and Km value of 15.67g/l (0.01567mg/ml)
respectively.
64
y = 0.095x + 0.006
R² = 0.986
-0.04
-0.02
0
0.02
0.04
0.06
0.08
0.1
0.12
-0.4 -0.2 0 0.2 0.4 0.6 0.8 1 1.2
I/V
(U
/mg
pro
tein
)-1
1/[S]
Figure 24: LineWeaver- Burk plot for lipase A
y = 0.094x + 0.006
R² = 0.992
-0.02
0
0.02
0.04
0.06
0.08
0.1
0.12
-0.2 0 0.2 0.4 0.6 0.8 1 1.2
I/V
(U
/mg
pro
tein
)-1
1/[S]
Figure 25: LineWeaver- Burk plot for lipase B
65
CHAPTER FOUR
DISCUSSION AND CONCLUSION
DISCUSSION.
This study has highlighted the potential for exploitation of lipases from a
common and inexpensive plant source for industrial purpose. Studies on the
Cucumeropsis mannii showed that for germination properties, the coated seed
(C.mannii seed with the coat intact) appeared to be more appropriate or better in terms
of endosperm modification. A germination energy, water sensitivity and average root
length of 82, 18 and 4.28cm respectively compared to 68, 20 and 3.96cm for the
decoated shows that the results were within the acceptable values of 65, 15 and 3.8
minimum values respectively for viable seeds (Meerssche et al., 1983). Lipase was
found to be present in the germinated seeds of Cucumeropsis mannii. The activity
increased with seedling growth to a peak on the 4th
day of germination and decreased
gradually until the seventh day. This agrees with what was earlier obtained by Ivan et
al (1995) who reported optimum lipolytic activity on day 4 of germination of Rape
seed. Beevers and Hills (1987) also showed that Gossipium hirsatum has the highest
lipase activity on the 4th
day of germination, Helianthus annus on the 7th
day, glycine
max on the 5th
day and Lycoperison esculentum on the 3rd
day. The increase in lipase
activity during germination may be associated such to high metabolic activity taking
place in the endosperm of seeds during germination in which there was increased
lipolysis and the stored triacylglycerol were converted to sucrose through the
glyoxylate pathway and gluconeogenesis to provide energy for the seedlings growth
(Peter, 2006). An entirely different pattern was obtained in the changes in the protein
concentration during germination. The protein content had its peak on the 3rd
day.
This suggests that there are other proteins present in the seed which are mobilized
alongside lipase during germination.
The results obtained from the localization of lipase activity showed that white
melon seed lipase is a membrane bound enzyme as lipase activity was highest in the
lipid body followed by the supernatant and least in the pellet. Lipase activity in the
supernatant may be attributed to fragments of lipid body remaining after hydrolysis of
the stored triacylglycerols. This is in accordance with earlier reports by Huang et al.,
(1983) for rape seed and mustard seeds. Huang and Lin (1984) also observed that
lipid bodies from corn scutella contained lipases. Lipase activity in the pellet may be
as a result of debris or particles of the seeds that settled at the bottom of the test tube.
66
The lipase in the lipid bodies of Cucumeropsis mannii were solubilized by
incorporating 1.95% (w/v) of Tween 80 in the extraction buffer. Bahri (2000)
achieved solubilization of oil bodies from sunflower with 1.3% (w/v) of Tween 80. In
this study, even though Triton X-100 seemed to give better solublization in terms of
visual assessment (clearity of solution, figure 2), when compared to Tween 80, Tween
80 gave better result in terms of lipase activity. This suggests that Triton X-100 under
the condition of study inhibited lipase activity. The inhibition by Triton X-100 was
concentration dependent. Detergents do not only solubilize membrane bound lipase
but also interact with triacylglylcerol emulsions tending to accumulate at the oil-water
interface.(Verger, 1997; Cannan et al., 1998).Since the oil-water interface is the site
where lipases act; this explains the inhibitory effect of Triton X-100.
Purification of the Cucumeropsis mannii lipase using combined ammonium
sulphate and cold acetone saturation followed by dialysis achieved 3.0 fold increase in
lipase activity and 18.0% yield or recovery of the enzyme. After the final stage of
purification by Sephadex G-200 gel filtration column chromatography two peaks A
and B were obtained. The purification fold for the two lipases (A and B) increased to
5.4 for lipase A and 11.0 for lipase B while their yield (recovery) decreased to 1.0%
for lipase A and 2.0% for lipase B respectively. The relative low recovery observed
after the final stage of purification may be due to emphasis on purity rather than
recovery. Lin and Huang (1984) reported that their emphasis on purification reduced
the recovery of lipase from scutella of corn seedlings. Two pH optima of 7.5 and 5.9
were obtained for the two lipases, lipase A and lipase B respectively. This suggests
that the white melon seed contained two forms of lipases. The lipase A is an alkaline
lipase and lipase B is an acid lipase.
Researchers in the past reported that oil seeds contain acid lipase and/ or
alkaline lipase. This study demonstrates that Cucumeropsis mannii belong to the last
class of seed which contain both acid and alkaline lipases. Muto and Beevers (1974)
reported the isolation of both alkaline and acid lipase from castor bean (pH 9.0 and
5.0). Huang (1982) recorded a pH optimum of 9.0 with glycine – HCl buffer and 6.5
with imidazole – HCl buffer for soybean lipase. Abigor et al., (2002) reported pH 7.5
for lipase from Jatropha Curcas, Sana et al., (2004) recorded pH 7.0 for lipase from
germinating Brassica napus. A study by Hills and Murphy (1988) suggested that
microsomal and lipid body lipases from the same source have different pH optima and
substrate concentrations required to saturate them. The presence of acid and alkaline
67
lipase activity in Cucumeropsis mannii will enable the lipase isolated from this seed
to be applied in a wider range of industrial processes.
Temperature optimum of 370C was obtained for the two lipases. This result
agrees with earlier findings on lipases form Sesanum indicum (Mukherjee and
Murphy 1966), oil palm, Cucumeropsis edulis (Opute, 1975) and Brassica napus
(Sana et al., 2004). The two lipases were observed to be stable at temperatures up to
450C for 1 hour, but activity was still detectable up to temperature of 80
0C. This
shows that the lipases from white melon seeds are thermo stable. The result obtained
made in this work, correlates the evidence that Ca2+
is a very good activator of lipases
(Enijiugha et al., 2004; Isbilir et al., 2008). Although Kermasha and Van de Voort
(2004) observed that Ca2+
inhibited lipase activity from French bean. EDTA, a
chelating agent did not show significant effect on the activity of lipase from
Cucumeropsis mannii seeds. It is therefore, suggested that the enzyme may not be a
metalloenzyme. A slight decrease in activity observed in the presence of EDTA may
be as a result of interference with the enzyme adsorption on the substrate water
interface. Lin et al. (1986) and Enijiugha et al, (2004) reported that EDTA inhibited
lipase activity in African bean seeds. Lin and Huang (1984) and Beevers and Hills
(1987) had observed that EDTA inhibited lipase activity in castor bean seeds.
Studies on the effect of substrate concentration showed that the activity of the
lipases increased with increase substrate concentration until a saturation point of
0.06mg/ml, corresponding to 6% (w/v) of the substrate. The decline after this
concentration may be due to the effect of enzyme substrate concentration ratio or
enzyme inhibited by substrate concentration or change of physicochemical
characteristics (Khan et al., 1991). Ejedegba et al., (2007), observed a saturation point
of about 8mM for lipase isolated from coconut. Vajanti et al., (2001), recorded
optimum substrate concentration of 10% with Caesalpinia bonducella. L seed lipase.
The result obtained in this study could be very useful for manipulation of industrial
bio-catalytic reactions involving the use of lipase from Cucumeropisi mannii seed and
other plant lipases.
68
CONCLUSION
In summary, the results obtained demonstrate that the seeds of
Cucummeropsis mannii, which is widely grown in tropical West Africa and consumed
largely as food (soup thickener) contains two types of lipases (acid and alkaline
lipases). Results obtained showed that C.mannii lipases are membrane bound as more
activity was obtained in the lipid bodies after centrifugation of the homogenate.
Successful purification of the membrane bound lipase depends greatly on choosing a
good solubilizing agent. Though Tween 80 was used to achieve solubilization of the
enzyme, further purification was achieved through combined ammonium sulphate and
acetone precipitation, dialysis and Sephadex G-200 gel filtration column
chromatography. The investigation shows further that the two lipases have optimum
temperature of 370C and are stable at temperature up to 45
0C for 1 hour. Ca
2+
activated the lipases while Pb2+
inhibited their activity strongly. Cucumeropsis mannii
seeds could be good sources of lipase for industrial bio-catalytic processes especially
when it involves changing pH and required low cooling cost.
69
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79
APPENDIX 1: PROTEIN DETERMINATION
The concentration of protein in the extract was determined using the method
of Lowry et al., (1951), using Bovine Serum Albumin (BSA) as standard.
REAGENTS.
Reagent A - Consist of 2% Na2CO3 + 0.1N NaOH
Reagent B - Consist of 0.5% CuSO4 + 1% Na-K tartrate
Reagent C - Consist of 50ml of reagent A + 1ml of reagent B
Concentration of BSA used = 50mg/100ml = 0.5mg/ml and was obtained by
dissolving 0.05g of BSA in 100ml of water.
PREPARATION OF STANDARD CURVE: Twenty test tubes each containing 1ml
of varying concentrations (0 – 0.5mg/ml) of BSA was set up in duplicates. To the test
tubes 5ml of reagent C was added and allowed to stand for 10min. Then 0.5ml of
diluted folin ciocalteau (1:1) solution was also added to all the tubes, mixed and
allowed to stand for 30min. The absorbance of the resulting blue colour was read at
750nm. The values obtained were used to plot a standard graph from where the
concentration of proteins in the extracts were estimated.
80
APPENDIX 2: ABSORBANCE VALUES FOR PROTEIN STANDARD CURVE
Tube Number Average absorbance (750nm) Protein concentration (mg/ml)
1 0.00 0.00
2 0.11 0.05
3 0.15 0.10
4 0.27 0.15
5 0.34 0.20
6 0.41 0.25
7 0.45 0.30
8 0.52 0.35
9 0.56 0.40
10 0.64 0.45
81
APPENDIX 3: PROTEIN STANDARD CURVE
y = 1.487x
R² = 0.976
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0 0.1 0.2 0.3 0.4 0.5
Ab
sorb
an
ce@
75
0n
m
Proteinn concentration(mg/ml)
82
APPENDIX 4: METHO FOR PREPARATION OF FATTY ACID STANDARD
CURVE: Concentration of free fatty acid released by the activity of lipase was
determined using the method of Duncombe (1963). A standard curve was prepared by
the principle of Duncombe (1963) using oleic acid as the standard fatty acid.
REAGENT: Chloroform, 6.4% (w/v) Cu(NO3)2 .3H2O, 0.1%
sodiumdiethyldithiocarbamate in butanol. 1.0N aqueous triethanolamine
Concentration of oleic acid used = 25mM prepared by dissolving 0.75ml of oleic acid
in chloroform and making up the volume to 100ml.
PREPARATION OF FATTY ACID STANDARD CURVE: Twenty test tubes
each containing 1ml of varying concentration of oleic acid (0 – 25mM) was set up in
duplicates. To the test tubes, 0.5ml of sodiumdiethyldithiocarbamate was added and
allowed to stand for 20 min. the absorbance of the corresponding yellow colour
developed was read at 440nm. The absorbance values obtained were used to plot a
standard curve from where the concentration of free fatty acid released by the lipase
was estimated.
83
APPENDIX 5: VALUES FOR FATTY ACID STANDARD CURVE
Tube Number Average absorbance (750nm) Protein concentration (mg/ml)
0 0.00 0.00
1 0.44 2.50
2 0.51 5.00
3 0.60 7.50
4 1.06 10.00
5 1.13 12.50
6 2.01 15.00
7 2.21 17.50
8 2.30 20.00
9 2.34 22.50
10 2.59 25.00
84
APPENDIX 6: FATTY ACID STANDARD CURVE
y = 0.110x
R² = 0.953
0
0.5
1
1.5
2
2.5
3
0 5 10 15 20 25 30
Ab
sorb
an
ce@
44
0n
m
Concentration of fatty acid (mM)
85
APPENDIX 7: VARIATION OF SPECIFIC ACTIVITY AND PROTEIN
CONCENTRATION DURING SEED GERMINATION
Days of
germination
Specific activity (µm/min/mg
Protein
Protein (mg/ml)
0 1.70 8.92
1 2.97 11.80
2 7.41 12.90
3 7.28. 12.70
4 9.97 12.70
5 8.08 12.20
6 2.40 11.20
7 2.35 10.90
86
APPENDIX 8: RESULTS OF PRELIMINARY STUDY ON AMMONIUM
SULPHATE AND COLD ACETONE PRECIPITATION.
Conc (%)
AMMONIUM SULPHATE
ACETONE
Supernatant Precipitate Supernatant Precipitate
10 46.30 22.53 45.54 15.38
20 22.55 9.17 40.37 1416
30 42.19 16.83 50.18 23.24
40 39.28 32.14 38.13 22.39
50 36.59 39.10 22.57 64.95
60 26.06 5.18 27.78 41.95
70 20.47 59.5 46.38 36.19
80 34.29 11.08 26.15 30.75
90 20.01 36.23 30.02 49.08