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The myriad of biochemical reactions that occur in living systems are nearly all mediated
by a series of proteinaceous, biological catalysts known as enzymes. Enzymes differ
from ordinary chemical catalysts in several important respects namely:
(i) Higher reaction rates (10 6 -1012 times greater)
(ii) Milder reaction conditions � (neutral pH; temperature below 100�C)
(iii) Greater reaction specificity and
(iv) Capacity for regulation

Enzymes were first discovered in 1835 (-amylase) by Jacob Berzelius, but their stereo-
chemical and functional elucidation took some time and it was not until 1965 that the X-
ray crystallography of an enzyme, Lysozyme, was available. Since then however, nearly
some 2000 enzymes have been purified and characterized to at least some extent (Voet
and Voet, 1990).

Enzymes display immense specificity for their respective substrates which is achieved
through geometrically and physically complimentary interactions which permits enzymes
to be absolutely stereo-specific both in binding substrates and catalyzing reactions.

The regulation of enzymatic activity in vivo can occur by allosteric alteration of the active
site, by substrate binding affinity, by substrate or product feedback inhibition or by gene
regulation (Boyer, 1999).
The wide diversity of enzymes and the rapidly growing number of newly discovered
enzymes has led the International Union of Biochemistry to adopt a scheme for the
systematic functional classification and nomenclature of enzymes. This system is used
where possible in conjunction with the informal or trivial name but assumes fundamental



Type of Reaction Catalyzed
1. Oxidoreductases

Oxidation-reduction reactions

2. Transferases

Transfer of functional groups

3. Hydrolases

Hydrolysis reactions

4. Lyase

Elimination to form double bonds

5. Isomerase


6. Ligases

Bond formation with ATP hydrolysis

importance when ambiguity must be minimized e.g. Peptidyl-L-amino acid hydrolase has
classification number EC [where EC signifies "Enzyme Commision"; 3 indicates
the enzymes major class, hydrolase(as in Table 1); 4 denotes the subclass, peptide bonds;
17 designates its subclass, carboxypeptidase and the fourth number 1 is the enzymes
arbitrarily assigned serial number in its subclass].
Enzymes are proteins and as such their conformation is determined by their amino acid
sequence or primary structure which inevitably determines their secondary and tertiary
structure. Enzymic structure plays an important functional role as this will determine the
steric conformation at the active site of the enzyme and hence its effective activity.
Enzyme activity is affected by any or all of the following: Enzyme Concentration,
Substrate Concentration, inhibitors, pH an temperature and as a result the characterization
of an enzyme usually involves an assessment of its optimum performance in relation to
these criteria as well as in terms of their kinetics with regards to models put forward by
Michaelis Menten . According to Michaelis Menten, enzymatic activity can me defined in
terms of the Michaelis constant Km which is equal to the substrate concentration which
gives half the maximum velocity (Datta and Ottoway, 1976). The Km is usually estimated
using the Lineweaver-Burke plot, a reciprocal plot of the substrate concentration on the
initial enzyme velocity where the negative reciprocal of the x- axis intercept yields the
Km of the enzyme (figure 1).


Figure 1.
Line-Weaver Burke Reciprocal Plot of Substrate Concentration

versus Initial Velocity

Km could also be estimated from the graph of initial velocity versus Substrate
concentration or [S] but the hyperbolic nature of the graph (Figure 2) makes it difficult to
estimate the infinite [S] and consequently Vmax (where Km =[S] at � Vmax).

Figure 2. The effect of Substrate Concentration on Initial Enzyme Velocity


The Km value is also of important in determining whether an inhibitor is competitive
(inhibition nullified by increasing substrate concentration) or non-competitive (where
there is always inhibition even at high substrate concentrations). For a non-competitive
inhibitor the Km value for the enzyme in the presence or absence of the inhibitor remains
constant whereas for the competitive Km is less.

Enzymes due to their protein nature can be denatured by extremes of pH and temperature.
The optimum pH and temperature of the enzyme is the point where the velocity or
activity is maximum at a given substrate and enzyme concentration (Figures 3 and 4)..

Figure 3. Effect of pH Change on Enzyme Activity


Figure 4. Effect of Temperature Change on enzyme Activity

With advances in biochemistry and molecular biology it has now also become the norm
for the molecular structure, gene sequence and molecular catalytic mechanism to be
elucidated before the enzyme is deemed fully characterized.
This requires that the enzyme can obtained in pure form (Shi et al, 2001; Ikediobi and
Obasuyi, 1982) and usually involves the following sequence of steps. First the tissue
from which the enzyme is to be isolated from is grounded at very cold temperature in a
stable buffer. Triton X-100 or some other detergent is sometimes added to facilitate the
fragmenting of cellular organelles and membranes to release any bound enzyme. The
resulting milieu is then centrifuged to remove the particulates after which the proteins in
the supernatant are precipitated using ammonium sulfate. The desired enzyme would
constitute just one in a mixture of many proteins in the precipitate, however, they usually
can be separated quite well based on their molecular weights. The ammonium sulfate can
be removed by dialysis and the approximate molecular weight of the enzyme can be
established using SDS-Polyacrylamide gel electrophoresis. Testing of the bands using
substrate to assay enzyme activity should facilitate identification of the desired enzyme.
The molecular weight can then be determined by comparison to known standard. The


use of column chromatography to extract larger volumes of the enzyme from the
supernatant based on the expected molecular weight fraction can be done and once a
sufficient quantity is obtained the required biochemical and kinetic data as well as X-ray
crystallographic and other structural analytical data can be obtained.
This research paper will examine the distribution of o-diphenolase, its structure, modus
operandi and economic importance.


o-Diphenolase and its Role in Enzymic Browning of Foods

o-Diphenolase also referred to as polyphenoloxidase or catechol oxidase which catalyze
the oxidation of catechols or ortho-diphenols to orthoquinones has been established to be
a copper containing protein (Robb

Figure 5. Oxidation of Catechol Catalysed by o-Diphenolase

et al, 1965; Kidron et al, 1977, Anosike and Ayaebene, 1982) strongly related to both
tyrosinase and haemocyanin, which all have a dinuclear copper complex with histidine
ligands at the active site (Siegban, 2004). O-Diphenolases are ubiquitous enzymes
capable of mediating or participating in a number of physiological processes. There is a
general dubiousness surrounding some of the many functions associated with o-
diphenolase, however its role in enzymic browning has been long established. O-
Diphenolase has six histidine residues one of which is covalently linked to a cysteine
molecule. The distance between the copper atoms has been resolved by X-ray studies to
range from about 2.5 to 2.9 Angstr�ms depending on whether it is substrate bound or
free. The role which copper assumes involves the binding of oxygen at the active site
(Mayer and Harel, 1979; Fennol et al, 2004 and Siegban, 2004). While o-diphenolases
from animal issues are relatively specific for tyrosine and dopa (Mason, 1955), the fungal
and higher plant enzymes act on a range of mono and diphenols (Mayer and Harel, 1979;
Siegban, 2004; Fennol et al, 2004).


Figure 6. Crystal Structure of a Plant Diphenolase

Extracted from Klabunde et al, 1998

Marked differences in both the level of o-diphenolase activity and the content of its
substrates have been observed between cultivars of fruits (Matthew and Parplia, 1971),
vegetables (Ben-Shalom et al, 1978) and yams (Ikediobi and Obasuyi, 1982).
o-Diphenolase has been found to be extensively a membrane bound enzyme. Apart from
its location in chloroplasts, diphenolases have been reported to be located in
mitochondria, peroxisomes and microsomes (Mayer and Harel, 1979). The strength of
binding of o-diphenolases to membranes appear to vary depending on the tissues and the
stage of development of the plant (Mayer and Harel,1979). In tobacco, washing with
buffer suffices to release the enzyme from chloroplast lamellae (Hoffer, 1964). In most
cases more drastic conditions are required for the solubilization of membrane bound o-
diphenolases such as the use of detergents e.g Triton X-100 (Hrel, 1964; Walker and
Hulme, 1966) and sodium dodecyl sulfate (Yamaguchi et al, 1969)
In situ solubilization occurs following exposure to certain stress conditions (Volk et. al.,
1977) and also under more natural conditions of ripening of fruits or aging. Thus apple


(Harel et. al, 1966), grape ad banana ( Mayer and Harel, 1979) diphenolases become
increasingly soluble during fruit ripening

The compartmentalization of phenolic substrates of the enzyme, both in special cells
(Mace, 1963) and within cells (Roberts, 1962) have been reported. This results in the
separation between the enzyme and the bulk of its phenolic substrates in situ.
The rise in diphenolase activity which generally accompanies wounding and stress has
been attributed to the de novo synthesis of the enzyme (Hyodo and Uritani, 1966). Other
researchers have attributed the rise to activation of alredy existing enzyme rather than re-
synthesis (Balasbrumani et al, 1971).

Many other roles have been ascribed to diphenolase enzymes due to it's activity response
to various stimuli and also by virtue of it's location in the plant cell. Some of these roles

It has been correlated with fruit formation in certain fungi and bacteria
(Wilson, 1968; Leonard, 1973).
It has been correlated with melanin formation and as such has been deemed to
play a role in cellular resistance (Kuo and Alexander, 1967).
Diphenolases have been suggested to play a role in electron transport
(Kabowitz, 1938).
Its presence in chloroplast membranous structures have implicated a possible
role in photosynthesis (Mayer and Harel, 1979).
Its involvement in affecting the regulation of plant growth has been implied
(Gordon and Paley, 1961; Tomazowski and Thieman, 1966).
It has been implicated in rendering seed coats impermeable to water (Marbach
and Mayer, 1975).

Enzymic browning is one of the most importantt color reactions that affect foods. It is
catalyzed by diphenolase enzymes which facilitate the conversion of phenols to the
brown pigment melanin in an oxidation reaction.


Figure 7. Formation of melanin (Browning) from tyrosine. (From Lerner, 1953).

Ikediobi and Obasuyi (1982) purified the enzyme from yam and found the molecular
weight to be 107,000 � 5400 with temperature and pH optima of 25�C and 6.8
respectively. Activity was illustrated on catechol, chloregenic acid, dopamine and
pyrogallol. The enzyme was found to be inhibited strongly by dithiothreitol,
diethyldithiocarbamate, potassium cyanide, sodium metabisulfite, 2-mercaptoethanol
and L-cysteine. The rate for catechol conversion in sweet potatoes has been
measured to be 2.3 x 103S-1 (Baruah and Swain, 1959) corresponding to a rate-
limiting free energy barrier of around 13 kcal/mol.


Figure 8. Comparison of reactions catalysed by o-Diphenolase and p-

Diphenolase. (From Walker, 1995).

Studies done on ripe banana o-diphenolase show that dopamine is the only significant
substrate in the browning reaction of banana. The first reaction results in the
orthohydroxylation of phenol and the second, oxidation of the diphenol to orthoquinone.
The remaining portion of the reaction sequence involve non-enzymic oxidations and
ultimate polymerization of indole 5,6-quinone to brown pigments (Melanins) � as
schematized in figures 7 and 8.
Table 3. lists a number of phenols found in fruits and vegetables. Relatively few of these
serve as sustrate for diphenolase. The most important are catechin, 3,4
dihydroxyphenylalanine (DOPA) and tyrosine and the substrate specificity varies
depending on the source of the enzyme.

Table 2. Phenolic substrates of Diphenolase in fruits, vegetables, and seafoods.
Phenolic substrates
chlorogenic acid (flesh), catechol, catechin (peel), caffeic acid, 3,4-
dihydroxyphenylalanine (DOPA), 3,4-dihydroxy benzoic acid, p-cresol, 4-methyl
catechol, leucocyanidin, p-coumaric acid, flavonol glycosides


isochlorogenic acid, caffeic acid, 4-methyl catechol, chlorogenic acid, catechin,
epicatechin, pyrogallol, catechol, flavonols, p-coumaric acid derivatives
4-methyl catechol, dopamine, pyrogallol, catechol, chlorogenic acid, caffeic acid,
3,4-dihydroxyphenylethylamine (Dopamine), leucodelphinidin, leucocyanidin
catechins, leucoanthocyanidins, anthocyanins, complex tannins
Coffee beans
chlorogenic acid, caffeic acid
chlorogenic acid, caffeic acid, coumaric acid, cinnamic acid derivatives
catechin, chlorogenic acid, catechol, caffeic acid, DOPA, tannins, flavonols,
protocatechuic acid, resorcinol, hydroquinone, phenol
tyrosine, caffeic acid, chlorogenic acid derivatives
dopamine-HCl, 4-methyl catechol, caffeic acid, catechol, catechin, chlorogenic acid,
tyrosine, DOPA, p-cresol
tyrosine, catechol, DOPA, dopamine, adrenaline, noradrenaline
chlorogenic acid, pyrogallol, 4-methyl catechol, catechol, caffeic acid, gallic acid,
catechin, Dopamine
chlorogenic acid, catechol, catechin, caffeic acid, DOPA, 3,4-dihydroxy benzoic acid,
chlorogenic acid, catechin, caffeic acid, catechol, DOPA
chlorogenic acid, caffeic acid, catechol, DOPA, p-cresol, p-hydroxyphenyl propionic
acid, p-hydroxyphenyl pyruvic acid, m-cresol
Sweet potato
chlorogenic acid, caffeic acid, caffeylamide


flavanols, catechins, tannins, cinnamic acid derivatives
Extracted from Marshall et al, 2002 (Structures of the listed Phenolics can be found in the Appendix)

Browning is desirable in some foods e.g tea and coffee and in most plants it has been
associated with pest and bacterial resistance and wound healing .Some have even
ascribed anticancer and antioxidant properties to the melanins produced during the
browning reaction (Marshall et al, 2000).
Projected increases in the fruit and vegetable market for the future will however not occur
if enzymatic browning is not understood and controlled (Marshall et al, 2000). It is
estimated that over 50 percent losses in fruit occur as a result of enzymatic browning
(Whitaker and Lee, 1995) and this has increased interest in understanding and controlling
diphenolase enzymes in foods. Browning or melanosis has also been observed during the
storage of some high value crustaceans such as shrimp and lobster connoting spoilage
(Otwell et al, 1992) and losses and browning has been shown to adversely affect flavor
and nutritional value of foods (Marshall et al, 2002)..

Figure 9. Examples of enzymatic browning in banana

Extracted from Marshall et al, 2000

Figure 10. Examples of enzymatic browning in potato


Extracted from Marshall et al, 2000

Browning does not occur in intact plant cells due to vacuolar separation of the phenolic
substrates from the enzyme which is present in the cytoplasm. Cutting or damage to the
tissue brings the enzyme and substrate together resulting in the observed brown
pigmentation which impacts both the organoleptic and biochemical characteristics of
fruits and vegetables (Marshall et al, 2000).
The role of browning has been shown to be mediated by several factors, namely:
Tissue Diphenolase level [E]
Tissue Phenolic content [S]


Temperature and
Oxygen Availability

The control of browning consequently can be effected through the manipulation of these
factors (Marshall et al, 2002). Some resulting methods of control include:
The elimination of oxygen by vacuum packing, immersion in liquid, treatment
with reducing agents e.g. ascorbic acid and antioxidants e.g butylated
hydroanisole (BHA).
Inactivation of the enzyme by: (a) Chelating the copper prosthetic group of the
active site using EDTA, Sorbic Acid or (b) denaturing with steam treatment,
blanching, solar drying or freezing or (c) Inhibition with cysteine, honey,
heylresocinol etc.
Removal of the enzyme e.g. from juices by precipitation and ultrafiltration
Reducing Enzyme activity by acidifying or lowering the pH e.g Citric Acid
(Most diphenolases exhibit optimal activity at pH 6.8).

Other non-conventional methods of reducing enzymic browning of foods include the use
of antienzymes or enzymes which destroy some cofactor necessary for the reaction e.g
some cleavage oxygenases (Kelly and Prinkle, 1969); Catechol Transferase (Prinkle and
Nelson, 1963) and Protease (Labuza et al, 1992).


With the advent of recombinant DNA technology, numerous amino acid sequences of
diphenolase isozymes have been deciphered using cDNA sequencing techniques
(Marshall et al, 2000).
The inactivation of genes coding for these enzymes using anti-sense RNA specific for
diphenolase should lower the browning reaction as it effectively reduces diphenolase
gene expression and hence the concentration of the enzyme in situ. Anti-sense RNAs
were recently observed to selectively block the gene expression of other plant enzymes
such as polygalacturonase and peroxidase in tomatoes (Marshall et al, 2002).
Bachem et al (1994) determined that the expression of diphenolase in potatoes was
decreased through the use of anti-sense cDNA.
It is hoped that though the use of this technology that browning resistant varieties maybe
developed to prevent or significantly curtail the production of Diphenolase.



In closing Diphenolases have been shown to be ubiquitous enzymes which can have
significant impact on the shelf life and indeed the quality of fruits, vegetables an certain
shell fish due to their facilitation of enzymic browning. Enzymic browning was shown to
be responsible for about 50% of spoilage of fruits and vegetables and therefore its control
can be of significant economic impact to the global food supply. Several method for its
control during the processing and handling of foods were explored bearing in mind that
treatments administered should not affect product flavor, texture and color.
The use of enzyme inhibitors, reducing agents, anti-oxidizers, heating, refrigerating, anti-
enzyme and anti-sense RNA technology were all examined.
It is hoped that in the very near future the use of anti-sense RNA technology will be able
to see the production of food varieties with a much reduced propensities for enzymic

Study to quantify the economic value of global food losses due to
enzymic browning.
Elucidation of the genes for diphenolase enzymes for all major food
crops impacted by enzymic browning to facilitate the use of anti-sense
RNA technology to reduce browning.
Further explore the potential of melanin in fighting cancer.
Intensify research in food technology to find novel yet economical
ways to reduce enzymic browning in foods accompanied by an
education drive to enlighten the masses and other stakeholders.



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