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Chapter – 2
REVIEW OF LITERATURE
Natural colours are generally defined as materials extracted, isolated or otherwise
derived from plants, animals or microorganisms that are capable of imparting a
distinguishing colour when added.
2.1. Pigments Classification
2.1.1. Based on Origin
Pigments can be classified based on their origin as natural, synthetic or inorganic.
Natural pigments are produced by living organisms such as plants, animals and
microorganisms. Natural and synthetic pigments are organic compounds. Inorganic
pigments are found in nature or reproduced by synthesis (Bauernfeind, 1981).
2.1.2. Based on chemical structure of the chromophore
Pigments can be classified according to the chemical structure of the
chromophore as
Chromophores with conjugated systems (carotenoid, anthocyanin,
betalain,caramel and synthetic pigment) and
Metal-coordinated porphyrins (myoglobin, chlorophyll and their derivatives)
(Wong, 1989)
2.1.3. Based on application as food additives
Based on the application of the pigments as food additives, the Food and Drug
Administration (FDA) has classified pigments into two types
Certifiable (synthetic pigments and lakes) and
Exempt from certification (pigments derived from natural sources such as
vegetables, minerals or animals and synthetic counterparts of natural derivatives)
(Food and Nutrition Board, 1971; Frick and Meggos, 1988 and Wong, 1989)
2.1.4. Based on the structural characteristics of the natural pigments
Natural pigments are also classified based on their structural characteristics as
Tetrapyrrole derivatives (chlorophylls and heme colours),
Isoprenoid derivatives (carotenoids and iridoids),
N-heterocyclic compounds different from tetrapyrroles (purines, pterins,
flavins, phenazines, phenoxazines and betalains),
Benzopyran derivatives (anthocyanins and other flavonoid pigments) and
Quinones (benzoquinone, naphthoquinone, anthraquinone and melanins)
(Bauernfeind, 1981 and Hari, et al., 1994)
2.2. Characteristics of Major Pigments
2.2.1. Quinone
Quinones include large number of structural variants and have a great number of
colouring compounds. Their basic structure consists of a desaturated cyclic ketone that is
derived from an aromatic moncyclic or polycyclic compound. Coenzyme Q, represented
by the ubiquinone has the quinine structure. This group is widely present in animals,
plants and microorganisms and plays an important role in electron transport system in
their cells.
Quinones can be classified based on their structure as benzoquinones,
naphthoquinones, anthraquinones, dibenzoquinones, dianthraquinones and
dinaphthaquinones. Large number of quinines is found due to their variability in the kind
and structure of substituents. Quinones are found in plants: plastoquinones are found in
chloroplasts of higher plants and algae; ubiquinones are ubiquitous in living organisms;
menaquinones are found in bacteria; naphthoquinones in animals and anthroquinones are
found in fungi, lichens, flowering plants and insects. Many types of quinine are
byproducts of the metabolic pathways and a few organisms (fungi) produce large
quantities of quinines. In general, quinines produce yellow, red or brown colourations,
while quinine salts show purple, blue or green colours (Hari, et al., 1994 and Thomson,
1962b).
Structure of Quinone
2.3. Microbial Pigments
Pigments from natural sources have been obtained since long time ago, and their
interest has increased due to the toxicity problems caused by those of synthetic origin. In
this way the pigments from microbial sources are a good alternative. Some of more
important natural pigments are the carotenoids, flavonoids (anthocyanins) and some
tetrapirroles (chloropyls, phycobilliproteins). Another group less important is the
betalains and quinones. The carotenoids are molecules formed by isoprenoids units and
the most important used as colorant are the alpha and beta carotene which are precursors
of vitamin A, and some xantophylls as astaxanthin.
Beta-carotene is a pigment used in the industry, which is obtained from some
microalgae and cyanobacteria. Astaxanthin, an important carotenoid (red pigment) of
great commercial value, and it is used in the pharmaceutical feed and aquaculture
industries. This pigment is mainly obtained from Phaffia rhodozyma, Haematococcus
pluvialis and other organisms. The phycobilliproteins obtained from cyanobacteria and
some group of algae, have recently been increased on the food industries. Recently these
pigments are used as fluorescent marker in biochemical assays. Research group has
carried out studies about the factors that improve the production of these pigments
obtained from different microbial species as well as the methods for their extraction and
application (Canizares, et al., 1998).
Among the natural sources of colourants, microorganisms offer great scope and
hope. The ease of cultivation, extraction, the genetic diversity in microbes and
sophistication of technology has made their choice more feasible (Juailova, et al., 1997).
Among the different living organisms, bacteria, yeast, algae, fungi and actinomycetes
appear more efficient and attractive sources of biocolourants.
Nature is rich in colours (minerals, plants etc.) and pigment producing
microorganisms (fungi, yeasts and bacteria). Microbial pigments such as carotenoids,
melanins, flavins and quinines are used in many food industries (Dufosse, 2006). The
ability to synthesis pigments varies with the organism and its environment. Fluorescent
Pseudomonads are ubiquitous bacteria that are common inhabitants of the water, soil,
plants and animals and they are the most studied group (Jayalakshmi, 2008).
There are a number of microorganisms, which have the ability to produce
pigments in high yields, including species of Monascus (Hajjaj, et al., 2000) and Serratia
(Williams, et al., 1971a). The red pigments, produced in solid-state culture by several
species of the genus Monascus, have been traditionally used in many Asian countries for
colouring and securing a number of fermented foods (Francis, 1987).
Due to the global increase in the manufacture of processed foods and the
continued consumer demands for natural food ingredients, the market for natural
colorants for food use is estimated to grow (Downham and Collins, 2000). Currently, the
vast majority of the natural food colorants permitted in the European Union and the
United States are derived by extraction of the pigments from raw materials obtained from
the flowering-plants of the kingdom Plantae (Mortensen, 2006).
The production of many existing natural colorants of plant origin has a
disadvantage of dependence on the supply of raw materials, which are influenced by
agro-climatic conditions – in addition, their chemical profile may vary from batch-to-
batch. Moreover, many of the pigments derived from the contemporary sources are
sensitive to heat, light, and oxygen and some may even change their colour in response to
pH changes as in case of anthocyanins (Mapari, et al., 2005). Many ascomycetous fungi
naturally synthesize and secrete pigments and may thus provide a more reliable source
for natural, "organic" food colorants with improved functionalities (Duran, et al., 2002).
The diversity of fungal pigments is not only found in their chemical structures but also in
the colour range of these pigments that may add new or additional hues to the colour
palette of the existing colorants derived from contemporary sources (Mapari, et al.,
2006).
The use of additives in food plays a significant role to enhance its quality, hence,
acceptability (Corsetti and Settanni, 2007). In this context colors contribute a lot. Of 29
approved food colors, 16 are synthesized from petrochemicals and their long-term intake
is a matter of great concern (Evans and Wang, 1984). Thus, the urge for alternative
combines with the exploration of microorganisms. Several classes of fungi are known to
produce a wide range of water soluble pigments, but low productivity becomes a
significant bottleneck for commercialization (Hejaji and Wijffels, 2004). Since,
basidiomycetous fungi are usually difficult to grow in industrial scale, scientists
emphasized on ascomycetous fungi. The red mold, Monascus purpureus, traditionally
used for preparation of red rice, is a promising source of pigment (Mapari, et al., 2005
and Sandipan Chatterjee, et al., 2009).
The microorganisms such as Monascus, Rhodotorula, Bacillus, Achromobacter,
Yarrowia and Phaffia produce a large number of pigments. Commonly found microbial
pigments are carotenoids and astaxanthin. Carotenoids are yellow, orange and red
pigments, which are widely distributed in nature. They are utilized as food or feed
supplements and as antioxidants in pharmaceutical formulations (Miura, et al., 1998).
Several microorganisms have been shown to produce astaxanthin. They include
Agrobacterium aurantiacum (Misawa, et al., 1995), Phaffia rhodozyma and
Haematococcus pluvalis (Johnson and An, 1991). Among them Phaffia rhodozyma is a
potential candidate for commercial production due to its high astaxanthin content
(Andrews, et al., 1972). Astaxanthin added to poultry feed can improve the colour of
both egg yolks and flesh (Johnson, et al., 1980). Astaxanthin possesses an unusual
antioxidant activity, which has caused a surge in the nutraceutical market for the
encapsulated product.
Astaxanthin has potent antioxidant activity and may have a role in decaying or
preventing degenerative diseases in humans and animals (Schroeder and Johnson, 1993).
Research on the health benefits of astaxanthin is very recent and has mostly been
performed in vitro or at the pre-clinical level with humans (Higuera-ciapara, et al., 2006).
The medical literature suggests that β-carotene may exhibit anticancer activities and aid
in reducing the incidence of cardiovascular diseases (Sies and Krinsi, 1995). Further,
health benefits such as cardiovascular disease prevention, immune system boosting,
bioactivity against Helicobacter pylori, and cataract prevention, have been associated
with astaxanthin consumption (Ciapara, et al., 2006).
Wong and Koehler, 1981 reported that the species of Monascus produces several
natural pigments, including primary colour, red and the secondary colour, orange during
the solid state fermentation. These colours are widely used in Asia as food colourants.
Monascus are traditionally used in oriental countries, originally in China and Thailand, to
prepare fermented rice with strong red colour, which finds several applications ranging
from conferring colour to products such as wine, cheese and meat, to medicinal uses and
as a meat preservative.
List of some of the pigment producing microbes
Organisms Pigment Colour Reference
Janthinobacterium
lividum
Violacein Purple Matz, et al., 2004
Xanthomonas oryzae pv.
Oryzae
Xanthomonadin Yellow Rajagopal, et al., 1997
Staphylococcus aureus Zeaxanthin Yellow Hammond and White,
1970
Pseudomonas aeruginosa Pyocyanin Blue-
green
Baron and Rowe, 1981
Phaffia rhodozyma Astaxanthin Red Florencio, et al., 1998
Haematococcus pluvialis Astaxanthin Red Kobayashi, et al.,2001
Serratia rubidaea Prodigiosin like
pigment
Red Moss, 2002
Vibrio gazogenes Prodigiosin like
pigment
Red Moss, 2002
Alteromonas rubra Prodigiosin like
pigment
Red Moss, 2002
Rugamonas rubra Prodigiosin like
pigment
Red Gerber, 1975
Streptoverticillium
rubrireticuli
Prodigiosin like
pigment
Red Gerber, 1975
Bradyrhizobium Canthaxanthin Orange Lorquin, et al., 1997
Corynebacterium
insidiosum
Indigoidine Blue Starr, et al., 1966
Micrococcus roseus Canthaxanthin Orange-
pink
Cooney, et al., 1966
Dunaliella salina β-carotene Orange Jacobson and Wasileski,
1994
A maroon dye was extracted from the rhizome of Arnebia nobilis (Indrayan, et
al., 2004). Phycoerythrin pigment was isolated from the cyanobacterium, Nostoc
muscorum (Ranjitha and Kaushik, 2005). The nutritional requirements for pyoverdine
pigments from Pseudomonas aeruginosa was reported by Barbhaiya and Rao (1985b).
The pigment Xanthomonadin produced by the genus Xanthomonas has a role against
photo damage (Rajagopal, et al., 1997).
2.4. Pseudomonads
Pseudomonads are ubiquitous bacteria in nature. They possess variable metabolic
abilities that enable them to utilize a wide range of organic compounds, and occupy an
important ecological position in the carbon cycle. An essential pre - requisite for a
detailed investigation of the roles and evolution of Pseudomonads is an accurate system
of classification and identification. However, the classification of Pseudomonas strains is
not fully established due to the lack of an accurate taxonomic system (Satoshi
Yamamoto, et al., 2000).
Bacteria of the genus Pseudomonas are characterized as straight or slightly bent
Gram-negative rods with one or more polar flagellae, not forming spores. Their
metabolism is strictly aerobic and chemo-organotrophic (Palleroni, 1984; Staley, 1984;
Woese, 1992 and Palleroni, 1992). On the basis of rRNA / DNA-homology studies the
genus Pseudomonas has been classified and divided into five groups. However, recent
polyphasic and genomic taxonomic studies indicated that Pseudomonas is a multigeneric
entity and that the five rRNA groups are distantly related to each other. The most of the
rRNA groups are now assigned to new genera such as Comamonas, Hydrogenophaga,
Burkholderia, Ralstonia (Kersters, et al., 1996).
Therefore the genus Pseudomonas sensu stricto is now reserved to the RNA
homology group I of Palleroni, 1984 which presently contains more than 100 fluorescent
and non-fluorescent species, some of them was described only during the last four years
(Doudoroff, 1973; Pecknold and Grogan, 1973 and Palleroni, 1993). The nomenclature
“fluorescent” and “non-fluorescent” which is followed in Table 3.2 is based on the
observation that some of these bacteria, the fluorescent Pseudomonads, produce yellow-
green fluorescent water-soluble pigments (Palleroni, 1984), called pyoverdines
(occasionally also named pseudobactins) (Elliot, 1958).
Some Main Species and Newly Assigned Species Relevant for Pseudomonas Sensu
Stricto (RNA Homology Group I) (Palleroni, 1993 and Meyer, et al., 1997)
Pseudomonas Species
Fluorescent P. aeruginosa
P. putida
P. syringae
P. cichorii
P. agarici
P. monteilii
P. jessenii
P. mandelii
P. libanensis
P. orientalis
P. migulae
P. brassicacearum
P. fluorescens
P. chlororaphis
P. tolaasii
P. viridiflava
P. asplenii
P. rhodesiae
P. veronii
P. mosselii
P. cedrella
P. gessardii
P. thivervalensis
Non-fluorescent P. fragi
P. mendocina
P. pseudoalcaligenes
P. corrugate
P. stutzeri
P. alcaligenes
P. graminis
P. plecoglossicida
Pseudomonas fluorescens has multiple flagella. It has an extremely versatile
metabolism, and can be found in the soil and in water. It is an obligate aerobe but certain
strains are capable of using nitrate instead of oxygen as a final electron acceptor during
cellular respiration. Optimal temperatures for growth of P. fluorescens are 25-30C. It
tests positive for the oxidase test. The word Pseudomonas means 'false unit', being
derived from the Greek words pseudo (Greek: ψευδο 'false') and monas (Latin: monas, fr.
Greek: μονάς/μονάδα 'a single unit'). The word was used early in the history of
microbiology to refer to germs. The name 'fluorescens' is because secretes a soluble
fluorescent pigment called pyoverdine or pyoverdin (formerly called fluorescein), which
is a type of siderophore.
P. fluorescens is also a nonsaccharolytic bacterium. Heat stable lipases and
proteases are produced by Pseudomonas fluorescens and other similar Pseudomonads.
The lipases produced by Pseudomonas sp have become an integral part of the modern
food industry and are used in the preparation of a variety of products including fruit
juices, baked food, vegetable fermentation and dairy enrichment. Lipases have got wide
application in pesticides, detergents, cosmetics and perfume industry. They are also used
in leather industry for processing hides and skins (bating) and for treatment of activated
sludge and other aerobic waste products where they remove the thin layer of the fats and
facilitate oxygen transport. Lipases are of wide spread occurrence throughout the earth's
flora and fauna. They are found abundantly in microbial flora comprising bacteria, fungi
and yeast (Rekha and Ahmed John, 2010).
2.4.1. Importance of Pseudomonads
The strong interest in Pseudomonads is based on their different properties.
Included in this genus are human or phyto-pathogenic and plant growth promoting
bacteria. Other Pseudomonads influence the growth of plants. Rhizobacteria belonging to
the P. fluorescens and P. putida species are able to promote plant growth (O'Sullivan and
O' Gara, 1992; Jacques, et al., 1993 and Yoshikawa, et al., 1993). It has been shown that
these bacteria rapidly colonize plant roots and cause at the plant level statistically
significant yield increases. The presence of Pseudomonas leads to the suppression of
plant deleterious microorganisms both by the production of antibiotically active
substances (Budzikiewicz, 1993) and by the excretion of effective siderophores, the
pyoverdines, with high iron binding constants that cannot be utilized by pathogenic fungi
or bacteria (Kloepper, et al., 1980). Another mechanism is the production of organic
acids like succinic and lactic acid that increase plant growth directly (Yoshikawa, et al.,
1993).
Kloepper and Schroth, 1981 reported that the genus Pseudomonas is one of the
most diverse Gram negative bacterial genera (isolated from sources ranging from plants
to soils and water) which are straight or slightly curved rods, motile by means of polar
flagella. Pseudomonas is characterized by their ability to grow in simple media at the
expense of a great variety of simple organic compounds, without needing organic growth
factors. King’s B media is an optimal for most species of Pseudomonas isolation,
developed the first selective media (King’s A and King’s B) for the isolation of
fluorescent Pseudomonads (King, et al., 1954). In recent years, there has been much
success in obtaining biological control of plant pathogens using bacterization techniques.
Bacteria used as inoculants are mostly Pseudomonas fluorescens and
Pseudomonas putida types obtained from soils and plant surfaces. Some of these bacteria
produce metabolites, which chelate the environmental iron thus making it unavailable to
pathogens. To date a number of these diseases suppressive antibiotic compounds have
been characterized chemically and include N-containing heterocycles such as phenazines,
pyrrole type antibiotics, pyo-compounds and indole derivatives. A small number of
antibiotics like compounds that do not contain nitrogen have also been isolate from
fluorescent Pseudomonads, one of these metabolites 2, 4- diacetylphloroglucinol
(DAPG), is a major factor control of a range of plant pathogens. The antibiotic DAPG is
produced by Pseudomonads of worldwide origin, and its biosynthetic locus is conserved
in Pseudomonads obtained from diverse geographic locations. Bacteria that produce
DAPG play a key role in agricultural environments, and their potential for use in
sustainable agriculture is promising (Prasanna Reddy and Rao, 2009).
2.5. Pyoverdine
2.5.1. Occurrence and Structure
Pyoverdines are chromopeptides that consist of three structural parts, a quinoline
chromophore, a peptide chain and a side chain. Pyoverdines are yellow - green, water
soluble, chromopeptides, fluorescent pigment (Meyer, 2000) and the typical siderophore
of the fluorescent members of the bacterial genus Pseudomonas sp in the rRNA
homology group I of the family pseudomonadaceae (Insa Barelmann,et al., 2002). They
consist of three distinct structural parts, viz. a dihydroxyquinoline chromophore
responsible for their fluorescence, a peptide chain comprising 6 to 12 amino acids bound
to its carboxyl group, and a small dicarboxylic acid (or its monoamide) connected
amidically to the NH2- group (Diana Uria Fernandez, et al., 2003).
The chromophore 5 – amino -2, 3- dihydro- 8, 9- dihydroxy- 1 H- pyrimido - [1,2-
a] quinoline -1- carboxylic acid is identical for all pyoverdines and its catechol unit is one
of the binding sites for iron (III) (Regine Fuchs, et al.,2001). Several pyoverdine
isoforms, which structurally differ one from another by the nature of the dicarboxylic
acid side chains attached to the chromophore (Suc, Mal or their amides, Glu or Kgl),
varies from one strain to another.
Usually several pyoverdins co-occur having the same peptide chain, but differing
in the nature of the dicarboxylic acid. The peptide chains have a twofold function. They
provide two of the ligand sites for Fe3+
, and they are responsible for the recognition of
their Fe3+
complexes at the surface of the producing cell (Budzikiewicz, 1997a). The
members of the fluorescent Pseudomonas group are the best-studied plant growth-
promoting rhizobacteria. Some Pseudomonas species have been used as seed inoculants
on crops to promote plant growth (Kloepper, et al., 1980; Schroch and Hancock, 1982;
Leong, 1986 and Cook and Leong, 1987). Enhanced plant growth caused by these strains
is often accompanied by reductions in populations of deleterious microorganisms, the so
called minor pathogens (Burr and Caesor, 1984; Leong, 1986; Schippers, et al., 1987;
Kloepper, et al., 1980; Loper and Buyer, 1991 and Soeng, et al., 1991). It has been
suggested that Pseudomonads exert their beneficial effects in part by producing yellow-
green fluorescent siderophores, called pyoverdines (or pyoverdins or pseudobactins),
under iron-deficient conditions (Kloepper, et al., 1980; Schippers, et al.,1987 and Seong,
et al.,1991). Pyoverdines are thought to chelate iron efficiently, making it less available
to phytopathogens and thus promoting plant growth.
According to the structural features of the peptide chain, pyoverdines can be
divided into four groups (Regine Fuchs, et al., 2001)
Group 1: The most common structural form of a pyoverdine has a linear peptide
chain and cyclo-Nhydroxy-Orn as the C-terminal amino acid. A tetrahydro-
pyrimidine ring can be observed in some pyoverdines. It results from
condensation of Dab with the preceding amino acid of the chain.
Group 2: The second largest group is characterized by a C-terminal cyclic part
consisting of three to four amino acids, formed by an amide bond between the
carboxyl group of the C-terminal amino acid and the e-amino group of an in-chain
Lys.
Group 3: A small number of pyoverdines has peptide chains with a C-terminal
cyclodepsipeptidic substructure formed by an ester bond between the carboxyl
group of the C-terminal amino acid and an in-chain Ser or Thr.
Group 4: Several pyoverdines have a C-terminal free carboxyl group. Actually,
some or all of them may be hydrolysis products of the rather labile cyclic esters
(Group 3) which are readily hydrolysed at pH values of nine or higher.
2.5.2. Extraction of Pyoverdine
For pigment analysis, cells were grown on King’s B agar and Nutrient agar, and
testing for pigment production and spectral characteristics was performed by extraction
with methanol, according to Hildebrand, et al., 1994, using a visible – UV Kontron
Uvikon 860 spectrophotometer (Alvaro Peix, et al., 2005).
The culture supernatant was concentrated under vacuum. Then, FeC13.6H20 (1
g/liter of original medium) was added, and it was extracted twice with chloroform-phenol
(ml/g); to the pooled organic phase was added about three volumes of ether, and that
solution was extracted several times with small volumes of water. After repeated rinsing
with chloroform to remove the phenol, the pooled aqueous phase was evaporated to
dryness under vacuum; this yielded the crude extract. In some cases, the concentrated
supernatant was first saturated with sodium chloride or ammonium sulfate; the resulting
crude extract was then dissolved in water and the whole procedure was repeated before
further purification. Crude extract, 2-10 g, was dissolved in 0.05 M pyridine-acetate, pH
5, applied to a Carboxymethylethyl (CM)-Sephadex column (1.5 x 30 cm) and eluted
with the same buffer. Three fractions were collected: the first and second are brown and
are labeled A and B, respectively, A being dominant. The third fraction appeared as a
red-orange band that remained at the top of the column; it was eluted with a gradient of
increasing pyridine-acetate concentration (0.05 to 2 M, pH 5) which separates ferribactin
from the rest of the crude extract.
Fraction A, 0.1-0.2 g, was applied to a 50-cm CM-Sephadex column and eluted
with 0.05 M pyridine-acetate buffer. It was resolved into five fractions, numbered I
through V. All are brown except III, which is purple. Fraction IV, the main component,
was further fractionated by chromatography on Sulfopropyl (SP)-Sephadex with 5% (v/v)
acetic acid brought to pH 3.1 with pyridine (0.2%, v/v). This yielded brown ferric
pyoverdine and a minor yellow compound we call ferric pyoverdine A whose presence is
erratic and is likely to represent a modified pyoverdine. Fraction V was not
chromatographically distinguishable from fraction B.
The ferribactin fraction was purified by passage through a CM- cellulose column
in 0.05 M pyridine-acetate, pH 5; this was repeated as necessary. The criterion for purity
was that the optical spectrum showed a single peak in the visible region (460 nm); for the
other compounds, the criterion was that they move as a single spot on thin layer sheets
(Stephen B. Philson and Miguel Llinas, 1982).
2.5.3. Purification and Characterization of Pyoverdine
Bacteria were grown for 40 h at 25C in standard Succinate medium, the pH
being maintained in the range of 7.0 to 7.3 by periodic addition of HCl. At the time of
harvesting, batches were pooled to a volume of 20 1itre; FeCl, (200 mg Fe3+
1-l) was then
added to the culture, which was centrifuged, the pelleted cells being discarded. The
supernatant was concentrated to approximately 1 litre under reduced pressure, saturated
with NaCl, and extracted with 0.5 vol. CHCl3 / phenol (1:1, v/w). The aqueous phase was
discarded, and the organic phase was treated with 2 vol. diethyl ether and 100 ml distilled
water (Meyer and Abdallah, 1978).
The aqueous phase, containing the pigment-iron complex, was re-extracted three
times with ether in order to remove phenol completely, and then evaporated under
reduced pressure. This solution (20 ml) was applied to a column of CM - Sephadex C-25
(2.5 x 90 cm) eluted with 0.1 M pyridine/acetic acid buffer (pH 6.5). A front-running
minor fraction, generally representing less than 10 % of the total, was shown by
electrophoresis to contain a mixture of pigmented degradation products. The major
fraction consisted of the native Fe (III)-pigment complex. Repetition of the CM-
Sephadex C-25 step showed this fraction to be 99% pure, in agreement with the results of
electrophoretic analysis. The yield (after lyophilization) was about 130 mg per litre of the
initial culture medium (Meyer and Abdallah, 1978).
All purification procedures were carried out at room temperature. The cell-free
supernatant was buffered with 1 M HEPES (N-2-hydroxyethylpiperazine – N 9- 2 -
ethanesulfonic acid) buffer (pH 7.0) and then applied to a Chelating Sepharose Fast Flow
column (1.5 by 25 cm) that had been presaturated with CuSO4. The column was
equilibrated with 20 mM HEPES buffer (pH 7.0) containing 100 mM NaCl at a flow rate
of 100 ml/h. The column was then washed with 200 ml of 20 mM HEPES buffer and
eluted with 20 mM acetate buffer (pH 5.0) containing 100 mM NaCl. The A400 of each 10
ml fraction collected was determined.
The pyoverdine-containing fractions (peaks FI, FII, and FIII) were pooled
separately and lyophilized. The dried material from each of the fractions was dissolved in
1 ml of distilled water containing 10 mM EDTA and then applied to a Sephadex G15
column (1.5 by 100 cm) that had been pre-equilibrated with deionized water. Elution was
carried out with distilled water at a flow rate of 20 ml/h. Fractions (3 ml) were collected,
and the A400 of each fraction was determined. In addition, the fluorescence of each
fraction was determined at emission and excitation wavelengths of 460 and 400 nm,
respectively.
The pyoverdine-containing fractions were pooled, lyophilized, and stored at 48C
until they were needed. On the basis of the fraction fluorescence profiles after gel
filtration, peaks FI, FII, and FIII each yielded one major fluorescent compound; since
pyoverdines are the only fluorescent siderophores produced by P. fluorescens 2-79, the
three compounds were designated Pf-A, Pf-B, and Pf-C, respectively. The concentrations
of pyoverdines Pf-A, Pf-B, and Pf-C were estimated by using an extinction coefficient of
20,000 M-1
cm-1
(Rong Xioa and Kisaalita, 1995).
Pyoverdine absorption spectra were determined with a Beckman model DU 650
spectrophotometer. The effects of different pH values on the spectra of iron-free
pyoverdines Pf-A, Pf-B, and Pf-C were determined as follows: pyoverdines were
dissolved in 0.1 M citric acid phosphate buffers (pH 3.0, 5.0, 7.0, and 8.0) and 0.1 M
Tris-HCl buffer (pH 9.0), and then the absorption spectra from 300 to 500 nm were
determined and compared. To study the interaction of Cu2+
with pyoverdine Pf-B, 10.0
mM CuSO4 or 10.0 mM FeCl3 was added to 100 mM acetate buffer (pH 5.0) containing
3.0 mM pyoverdine Pf-B. The reaction mixture was kept at room temperature for 30 min,
and then the absorption spectra between 340 and 500 nm for free pyoverdine Pf-B, Cu2+
pyoverdine Pf-B complexes, and Fe3+ pyoverdine. Pf-B complexes were determined and
compared (Rong Xioa and Kisaalita, 1995).
Pyoverdin was purified by a modification of the methods of Meyer and Abdallah,
1978 and Stephen B. Philson and Llinas, 1982 for pyoverdin siderophores. Strain B301D
was grown from an initial concentration of 5 x 106 CFU/ml in 1-liter flasks containing
250 ml of deferrated NM liquid medium. Cells were grown at 25°C with rotary shaking
(250 rpm); after 24 h, 4 ml of a FeCl3 stock (1 M) was added per liter of medium. After
the mixture was stirred for 20 min, cells and precipitate were removed by centrifugation
at 10,000 X g for 20 min. The culture supernatant was concentrated 50-fold by flash
evaporation at 55°C, and then contaminating proteins were precipitated by saturation
with NaCl. After removal of the precipitate by centrifugation, the concentrated culture
supernatant was extracted with an equal volume of chloroform- phenol (1:1[v/w]) (Cody
and Gross, 1987).
The organic phase was mixed with 3 volumes of diethyl ether, and the pigment
then was partitioned into deionized water (20 ml) until color could no longer be
extracted from the organic phase. The resulting aqueous extract was washed three times
with diethyl ether to remove phenol and reduced to dryness by lyophilization. The crude
siderophore extract, 50 to 150 mg, was further purified through a carboxymethyl (CM)-
Sephadex (Pharmacia, Piscataway, N.J) cation-exchange column (1.5 by 90 cm)
equilibrated with 50 mM pyridine-acetate buffer, pH 5.0. The column was eluted with the
same buffer. Absorbance was monitored at 405 nm and all peaks were collected. The
predominant reddish-brown peak was non-fluorescent and consisted of ferric
pyoverdinpss. The ferric pyoverdinpss was lyophilized, suspended in pyridine-acetate
buffer (1 ml), and again passed through the CM- Sephadex column to increase its
homogeneity.
The purity of the pyoverdine was determined by thin-layer chromatography, using
Silica Gel G (Merck and Co., Inc., Rahway, N.J.) Uniplates (Analtech, Newark, Del.)
developed with a 70:30 (v/v) ethanol-water solvent system. A homogeneous preparation
of pyoverdin was noted by the presence of only one sharp reddish-brown spot; no
fluorescent spots were observed when plates were illuminated with UV light (366 nm)
(Cody and Gross, 1987).
The siderochromes were monitored during various procedures by thin layer
chromatography, using silica gel-impregnated glass fiber sheets (ITLC SA, Gelman) and
70:30 (v/v) ethanol-water as solvent. The colored ferric compounds were observed
directly. Since all the iron-free pigments but deferriferribactin are fluorescent, very small
quantities could be seen under long wavelength UV irradiation, after spraying with an
aqueous EDTA solution (Stephen B. Philson and Miguel Llinas, 1982).
2.5.4. Microbial Production of Pyoverdine
The synthesis under certain growth conditions of yellow-green, fluorescent,
water-soluble pigments is a characteristic property of some Pseudomonas sp. (Stanier, et
al., 1966). These species are all members of the same intra-generic genetic homology
group (Palleroni, et al., 1973). They include P. aeruginosa, P. putida, P. fluorescens and
phytopathogens of the P. syringae type (Palleroni and Doudoroff, 1974). Many different
environmental factors affect the synthesis of these pigments, notably the chemical nature
of the organic carbon and energy source (Lepierre, 1895; Sullivan, 1905; Blancheti Rae,
1920; Giral, 1936; Gouda and Chodat, 1963 and Gouda and Greppin, 1965), the degree of
aeration of the culture medium (Elliot, 1958 and Lenhoff, 1963), pH and light (Greppin
and Gouda, 1965) and the cations Mg2+
(Georgia and Poe, 1931), Zn2+
(Baghdiantz, 1952
and Chakrabarty and Roy, 1964a) and Fe3+
(King, et al., 1948; Totter and Moseley, 1953;
Lenhoff, 1963; Palumbo, 1972 and Lluch, et al., 1973). No clear-cut physiological role
has so far been assigned to this class of pigments.
Various structures have been proposed for the pigments including pteridines
(Giral, 1936 and Chakrabarty and Roy, 1964b), flavines (Birkhoffer and Birkhoffer,
1948) and pyrroles (Lenhoff, 1963 and Greppin and Gouda, 1965), but none is supported
by unequivocal chemical evidence. The factors that affect pigment synthesis, the
purification of the pigment and its physicochemical properties were described by Meyer
and Abdallah (1978). Turfreijer (1942) proposed the term ' pyoverdine ' for the yellow-
green, fluorescent, water-soluble pigment of P. fluorescens. He chose this name by
analogy with that of the phenazine pigment, pyocyanine, produced by P. aeruginosa.
This designation is preferable to those of ' bacterial fluorescein' or ' fluorescin' which
have also sometimes been used in the literature (King, et al., 1948 and Lenhoff, 1963).
Other authors (Elliot, 1958 and Hulcher, 1968) have extended the name of
pyoverdine to include all pigments produced by fluorescent Pseudomonads. However,
since preliminary unpublished studies have shown that there are minor structural
differences between the pigments of the different species of fluorescent Pseudomonads, it
seems desirable to designate the pigments of this class by a suffix indicating the species
responsible for their production, e.g., pyoverdine, for the pyoverdine produced by P.
fluorescens (Meyer and Abdallah, 1978).
P. fluorescens 2-79 was grown on a synthetic succinate medium and the pH was
adjusted to 7.0 by adding a 1 N NaOH solution prior to sterilization. The medium was
dispensed into 500 ml Erlenmeyer flasks, each of which contained 100 ml of medium.
The flasks were then inoculated, at a seed rate of 1.0%, with exponential-phase cells
grown in the same medium. The cultures were incubated at 25C and 200 rpm in a New
Brunswick model Innova 4000 shaker-incubator for 48 h. Batches were pooled and
centrifuged at 10,000 X g for 10 min at 4C. The resulting supernatant was membrane
filtered (pore size, 0.25 mm; Amicon) to yield a cell-free solution of crude pyoverdines.
The amount of pyoverdine produced was estimated by determining the A400 of the
supernatant (Rong Xioa and Kisaalita, 1995).
P. fluorescens SS101 was grown on Pseudomonas agar F (Difco) plates or in
liquid King’s medium B (KB) at 25°C (De Bruijn, et al., 2008). P. fluorescens were
grown in a succinate minimal medium (Budzikiewicz, 1997). For the work-up of the
culture and isolation of the ferri-pyoverdins by chromatography on XAD-4 and Biogel P-
2 (Georgias, et al., 1999). From both strains two fractions were obtained which were
further purified by chromatography on CM - Sephadex C-25 with a pyridinium acetate
buffer (pH 5.0, gradient 0.02 to 0.2 m); final purification by RP-HPLC on Nucleosil-100
with 50 mm acetic acid/methanol (gradient 3 to 30% acetic acid). Decomplexation was
achieved by adsorption of the ferri-pyoverdins on a Sep-Pak cartridge and washing with a
6.5% Sodium oxalate solution (pH 4.3). After removing all salt residues with water the
free pyoverdins were eluted with methanol/ water 1:1 (v/v). The solutions were brought
to dryness and the samples were stored at -25C (Insa Barelmann, et al., 2002).
For qualitative and quantitative analysis of the amino acids by total hydrolysis,
determination of their configuration by GC/MS of their TAP derivatives on a chiral
column and dansyl derivatization of the free amino groups (Briskot, et al., 1986 and
Mohn, et al.,1990). Partial hydrolysis was affected with 6 N HCl at 110C for 15 min.
The peptide fragments were separated by chromatography on Bio-Gel P-2 with 0.1 M
acetic acid. Subsequently they were subjected to total hydrolysis, TAP derivatization and
GC-analysis as above. The 5-hydroxy chromophore from 1 and 2 for CD analysis was
obtained as described by Michels, et al., 1991.
Pseudomonas aeruginosa has the ability to produce a variety of water-soluble
pigments, including pyocyanin, pyoverdin, fluorescein, pyomelanin, and pyorubin.
Although the incidence of strains producing the latter two pigments is low, these
pigments can mask the production of pyocyanin. Two pigment-enhancing media, based
on research using different peptone ingredients in the agar was developed by King, et al.,
1954. Pseudomonas P (Tech) Agar or Pigment Production Agar - A incorporates
pancreatic digest of gelatin, which enhances the production of pyocyanin and inhibits the
pigment fluorescein. Pyocyanin production is also stimulated by the incorporation of low
concentrations of magnesium chloride and potassium sulfate. Once the pigment is
produced, it diffuses into the media giving a characteristic green color.
Pseudomonas F (Flo) Agar or Pigment Production Agar - B incorporates a
combination of casein and animal tissues, along with K2HPO4, which promotes the
production of fluorescein. Pyocyanin production is inhibited. Detection of this pigment,
which produces a yellow-green color, may be aided by the use of ultraviolet light (Gildo
Almeida da silva and Erik, 2006). When these media are used in combination with one
another, they are useful in the identification of P. aeruginosa and P. fluorescens
(Lennette, 1985). A bright yellow-green pigment diffused into the agar is indicative of
fluorescein production. The use of an ultraviolet light may aid in the detection of this
pigment. If fluorescein is present, it will fluoresce with a yellow-green colour. No
pigment production of fluorescence is indicative of a negative test (Washington, 1981).
King Agar B enhances the elaboration of fluorescein and inhibits the pyocyanin
formation. Mixed peptone provides the essential nitrogenous nutrients, carbon, sulphur
and trace elements. Glycerol serves as a C-source and Dipotassium hydrogen phosphate
buffers the medium. Magnesium sulfate is necessary for the activation of fluorescein
production (Mac Faddin, 1985). P. aeruginosa build colonies surrounded by a yellow to
greenish-yellow zone due to fluorescein production which fluoresces under UV light. If
pyocyanin is also synthesized, a bright green colour is produced. For the confirmation of
pyocyanin the coloured pigments can be extracted with chloroform. Most pyocyanin-
producing Pseudomonas strain synthesizes also fluorescein and others produce just one
pigment. The temperature can be a determining factor as most fluorescent strains will not
grow at 35°C. Rather, they grow 25-35°C. Atypical pyocyanin-negative, fluorescein-
positive, P. aeruginosa strains can also be differentiated from P. fluorescens and P.
putida (Blazevic, et al., 1973).
2.5.5. Molecular biology and Genetics
Sequencing a cloned PCR fragment spanning the junction of PA2400 and PA2401
in the P. aeruginosa genome showed that a GC base-pair was missing from the genome
sequence at position 2669175 and when this was included PA2400 and PA2401 form a
single reading frame PA2400/1 (pvdJ). Southern blotting was carried out by standard
methods (Sambrook, et al., 2000), using as probes radiolabelled PCR fragments or cloned
restriction fragments corresponding to individual genes, except that pvdN and pvdO were
part of the same PCR fragment. Membranes were washed at 65°C in 0.1% SDS / 0.1 %
SSC prior to autoradiography.
DNA sequences were obtained from the P. aeruginosa genome project
(http://www.pseudomonas.com) and the P. aeruginosa genome database
(http://pseudomonas.bit.uq.edu.au). Sequences were manipulated using Seqed (Devereux,
et al., 1984) and analysed using NLDNA and Codon use as described previously
(Merriman, et al., 1995). Database searches and analysis of the genomes of other
fluorescent Pseudomonads were carried out at the National Center for Biotechnology
Information (http: // www. ncbi.nlm.nih.gov) with BLAST algorithms (Lamont and
Martin, 2003).
For 16S rRNA gene sequencing and comparative analysis, DNA extraction,
amplification and sequencing of 16S rRNA genes were performed as already reported
(Rivas, et al., 2003). Nearly complete 16S rRNA gene sequences for isolates CH01T and
PA01 were obtained (1532 and 1530 nucleotides, respectively). They were compared
with sequences retrieved from GenBank by using the BLAST program (Altschul, et al.,
1990), and aligned by using CLUSTAL X software (Thompson, et al., 1997 and Alvaro
Peix, et al., 2005).
P. fluorescens is a heterogeneous species that can be subdivided by various
taxonomic criteria into several biotypes.The main aim was: (i) to develop a rapid
polymerase chain reaction (PCR) - based species-specific protocol to identify
P. fluorescens and (ii) to distinguish the different biotypes of this species by combining
molecular techniques with traditional biochemical methods (Mauro Scarpellini, et al.,
2004). The genotyping was performed by PCR-RAPD analysis. The purpose of the study
was to apply the RAPD technique for characterization of P. fluorescens and evaluate the
ability of this technique to differentiate between them (Prasanna Reddy and Rao, 2009).
To confirm strains as fluorescent Pseudomonas, 16S–23SrRNA intervening sequence
specific primers ITS1F (AAGTCGTAACAAGGTAG) and ITS2R
(GACCATATATAACCCCAAG) were used to get an amplicon size of 560 bp
(Gunasekaran, et al., 2002).
Chen and Kuo, 1993 developed a simple and rapid method for the preparation of
Gram negative bacterial genomic DNA. For DNA extraction, the selected strains were
cultivated in LB medium (5 g of yeast extract, 5 g of sodium chloride and 10 g of
peptone/L) during 24 h at 28°C and 40°C. Subsequently, the culture was centrifuged for
10 min at 10000 rpm, and DNA was extracted following the methodology described by
Bramwel, et al., 1995. Finally, 100 µl of buffer was hydrated and frozen at -20°C to be
used. The amplification of subunits 16S rRNA of the B. cepacia and P. fluorescens
strains was performed by PCR. For B. cepacia, the following primers were used: 5'TGG
TAG TCC ACG CCC TAAACG3' (forward) and 5'AGG GCC ATGA GGA CTT GAC
G3' (reverse). For P. fluorescens, the following primers were used: 5'CGT GTG
TGAAGAAGG TCT CGG3' (forward) and 5'TTA CGG CGT GGA CTA CCA GG3'
(reverse).
Amplification was performed in sterile PCR tubes (500 µL) at a volume of 50 µL
of buffer 10X, 1U of polymerase Taq, 100 µM of each d NTP (dATP, dCTP, dTTP and
dGTP), 2 µM of each primer, and 1 µL of purified DNA extract. The first PCR cycle
lasted 5 min, at 94°C for DNA denaturation, 6 min for hybridation (53°C), and extension
(72°C). The other 29 cycles were programmed for 1 min of denaturation at 95°C, and 3
min for hybridation and extension. Final extension was done for 10 min at 72°C.
Amplifications were performed in automatic equipment 2400 (Perkin Elmer) (Rychlik,
1995). As negative control, PCR was done using DNA of A. brasilense Sp7 with the
previous primers. The PCR products were analysed by electrophoresis 1/10 of final
volume of agarose gel reaction at 1% in TBE 1X buffer supplemented with 1 µg/mL of
ethyl bromide, according to the procedure described by Sambrook, et al., 1989.
2.5.6. Application
2.5.6.1. Antagonistic activity of Pseudomonas fluorescens
Nature has been a source of medicinal agents for thousands of years. An
impressive number of modern drugs have been isolated from microorganisms, mainly
based on their use in traditional medicine. In the past century, however, an increasing role
has been played by microorganisms in the production of antibiotics and other drugs
(Fenical, 1993 and Ahmed, et al., 2008).
Microorganisms have been the study of importance in recent years because of
production of novel metabolites, which exhibits antimicrobial, antiviral, antitumor as well
as anticoagulant properties. These secondary metabolites serve as model systems in
discovery of new drugs (Marderosian, 1969; Austin, 1989; Mlinski, 1993; Fenical, 1993
and Bernan, et al., 1997). Most of the current antimicrobial drugs are the derivatives of
the earlier generation and microbial resistance against them further intensify the need for
new drug discovery. Acceptable options available are the metabolites of plants or animal
origin, which are biocompatible, biodegradable and non-toxic in nature. These
metabolites are widely studied and are produced by various groups of microorganisms.
Pseudomonas (Elander, et al., 1968; Flood, et al., 1972; Castric, 1975; Chain and
Mellows, 1977 and Gross and De Vay, 1977) and Streptomyces (Shanshoury, et al., 1966;
Tahvonen, 1982; Yuan and Crawford, 1995 and Trejo Estrada, et al., 1998) are the
groups of microorganisms, studied for their secondary metabolites (Madhava Charyulu,
et al., 2009).
Fungal diseases in agriculture crops pose a great challenge. The currently used
fungicides are either less effective or of great environmental concern. Hence, it is
necessary to develop environmentally safer and potent fungicides of natural origin.
Microbial metabolites have been expected to minimize the deleterious side effects caused
by synthetic fungicides. In this regard, actinomycetes deserve to be studied, as they are
known to produce potent antibiotics. Compared to terrestrial species, marine
actinomycetes are important sources of novel antibiotics (Okami, 1986 and Newman, et
al., 1989). The actinomycetes found in the marine and coastal ecosystems may be viewed
as a rich gene pool possibly containing isolates capable of producing useful metabolites.
However, only few attempts have been made out to isolate marine actinomycetes
especially for their antifungal activity". In the present study, the isolated actinomycetes
from marine samples have been tested against four fungal pathogens that cause
economically important diseases in crops, like rice and sugarcane, to identify potent
antagonistic strains (Kathiresan, et al., 2005).
Strains of Pseudomonas fluorescence showed known biological control activity
against certain soil-borne phytopathogenic fungi and has the potential to produce known
secondary metabolites such as siderophore, HCN and protease that showed antagonistic
activity against Macrophomina phaseolina, Rhizoctonia solani, Phytophthoranicotianae
var. parasitica, Pythium sp. and Fusarium sp. Siderophore production in Pseudomonas
strains inhibited the growth of Staphylococcus sp, Escherichia coli and Aeromonas
hydrophila to 96.7% (Ahmadzadeh, et al.,2006).
The genus Pseudomonas has been heterogenous since Migula first named it in
1984 (Migula, 1984). He designated and described the species associated with the genus
in 1985 (Migula, 1985). Pseudomonas is gram-negative, strictly aerobic, polarly
flagellated rods. They are aggressive colonizers of the rhizosphere of various crop plants,
and have a broad spectrum antagonistic activity plant pathogens, such as antibiosis (the
production of inhibitory compounds) (Cartwright, et al., 1995 and Rosales, et al., 1995),
siderophores production (iron-sequestering compounds) and nutrition or site competition
(Winkelmann and Drechsel, 1997).
Some species of Pseudomonas can also produce levels of HCN that are toxic to
certain pathogenic fungi (David and O’Gara, 1994). These characteristics make
Pseudomonas species good candidates for used as seed inoculant and root dips for
biological control of soil-borne plant pathogen against different plant pathogenic fungi
viz. Alternaria cajani, Curvularia lunata, Fusarium sp., Bipolaris sp. and
Helminthosporium sp. in vitro (Srivastava and Shalini, 2008).
Plant protection is an important area which needs attention since most of the
hazardous inputs added into the agricultural system are in the form of plant protection
chemicals. Pseudomonas fluorescens possess a variety of promising properties which
make it a better biocontrol agent. The objectives of the present study were to isolate
P. fluorescens from soil, to check its antagonistic activity and effect of its secondary
metabolites on three fungal plant pathogens by in vitro techniques. P. fluorescens was
isolated from rhizosphere soil on King’s B medium and its antagonistic effect on three
fungal plant pathogens was studied in vitro. Its antagonistic activity was checked by co-
inoculation with the fungal isolates.
In pour plate method, P. fluorescens on coinoculation with fugal pathogens
decreased their growth rate. Maximum inhibition was observed in Pythium ultimim (80%)
followed by Macrophomina phaseolina (70%) and Pyricularia oryzae (50%). Effect of
the separated secondary metabolites on the fungal growth by broth dilution technique and
antifungal activity by agar well diffusion technique was studied. P. fluorescens produces
a broad-spectrum antifungal compound, which inhibits a variety of plant pathogenic fungi
and inhibits Pythium ultimum more when compared to other plant pathogens in the
present study. Further investigations on the type of antifungal components and in vivo
experiments will make P. fluorescens as one of the most suitable biocontrol agent in
suppressing the phytopathogenic fungi and replace chemical fungicides (Goud and
Muralikrishnan, 2009).
Bacteria - bacterial pathogen interactions are having an increased interest in
antibacterial interactions involved Pseudomonas sp. The genus Pseudomonas actively
solubilizes phosphate in vitro and produces several antibiotics with high specificity
against several microorganisms like Salmonella typhi, Streptococcus mutans, Bacillus
subtilis, Shigella sonnei. The Pseudomonas fluorescens strain seems to be highly
potential in controlling bacterial pathogens of health significance and result in
development of a potential biocide (Rekha and Ahmed John, 2010).
The plant growth promoting rhizobacteria (PGPR) improve plant growth either
directly via production of plant growth regulators such as auxine and cytokinines and by
increasing the plant uptake of some micro and macro elements in the rhizosphere
(Lugtenberg, et al., 1991 and Glick, 1995) or indirectly, through biological control of
pathogens or induction of host defense mechanisms (O’Sullivan and O’Gara, 1992;
Thomashow and Weller, 1996; Van Loon, et al., 1998 and Pieterse, et al., 2000).
Production of antifungal secondary metabolites, such as 2, 4 - diacetylphloroglucinol (2,
4-DAPG), pyoluteorin (PLT), pyrrolnitrin (PRN), phenazines, hydrogen cyanide (HCN),
siderophore and lytic enzymes (protease), is a prominent feature of many biocontrol
fluorescent Pseudomonads (Fenton, et al., 1992; Lemanceau, et al., 1992; Dowling and
O’Gara, 1994 and Thomashow and Weller, 1996).
Fluorescent Pseudomonads that produce the polyketide antibiotic 2, 4-DAPG are
an important group of PGPR that are effective against a broad spectrum of soil-borne
plant pathogenic fungi (Shanahan, et al., 1992; Harrison, et al., 1993; Nowak Thompson,
et al., 1994; Duffy and Defago, 1997 and Sharifi Tehrani, et al., 1998).
A proteobacterium isolated from coastal region produced appreciable secondary
metabolite and partial purification of the obtained secondary metabolite demonstrated
antimicrobial activity against both Gram positive and negative organisms such as
Klebsiella pneumonia, Staphylococcus aureus, Shigella flexneri, Pseudomonas
aureginosa, and Bacillus subtilis including MRSA (methicillin resistant Staphylococcus
aureus). Identification of the isolate using biochemical tests, 16S rDNA sequence
analysis, G+C content and electron microscopy studies revealed that the isolate belonged
to Pseudomonas genera. Extraction, purification, characterization and antimicrobial
activity of secondary metabolite carried out using various standard instrumental
techniques suggested that the active fraction was of 272 m/z with a stable fragment of
244 m/z and also displayed stable free radical activity assessed using EPR analysis. This
stable free radical activity of secondary metabolite mediated its antimicrobial activity
(Madhava Charyulu, et al., 2009).
Antibacterial activity of isolated bacterial strains was checked by cross streak and
agar well diffusion method. Crude extract were prepared from antibacterial metabolite
producing strains using organic solvents. A total of six strains showed antibacterial
activity against Aeromonas punctata, Kokuria marina, Rothia marina, Vibrio cholerae,
Staphylococcus aureus, Staphylococcus epidermidis, Escherichia coli, Staphylococcus
aureus, methicillin resistant Staphylococcus aureus and Proteus vulgaris. Production of
antibacterial metabolites varied within 24 to 72 h, in marine broth 2216. A total of 6
crude ethyl acetate extracts of antibacterial substance were prepared from the
antibacterial metabolite producing strains grown on marine agar 2216 and screened for
their antibacterial activity against test bacterial strains. Out of 6 extracts, 3 extracts
showed activity against marine and clinical isolates (Ahmed, et al., 2008).
Antibacterial metabolite producing strains were identified by 16S rRNA gene
sequence analysis. Active bacterial strains belonged mainly to genera of Pseudomonas,
Bacillus and Bravebacterium. The recovery of strains with antimicrobial activity suggests
that marine environment may represent an ecological niche which harbors a largely
uncharacterized microbial diversity and a yet unexploited potential in the search for new
secondary metabolites (Ahmed, et al., 2008).
The antimicrobial activities of these strains were studied against Salmonella
typhimurium, Staphylococcus aureus, Fusarium culmorum, F. oxysporum and Aspergillus
niger. Despite equal siderophore activities with various Pseudomonas sp. as measured by
the chrome azurol S assay, the study shows how siderophore activity does not correlate
with the antibacterial activity against food pathogens or with the antimould activity
against pathogenic moulds. Furthermore, the results illustrate how siderophores are able
to act both as growth inhibitors and stimulators (Laine, et al., 1996).
2.5.6.2. Pigment in Textile and Paper industry
At present, fabrics are dyed mainly with synthetic pigments. However, natural
pigments are still valuable because of their natural colour tones. Janthinobacterium
lividum was isolated from wet silk thread whose color became bluish-purple. This
bacterium produced large amounts of bluish-purple pigment on some media containing
amino acids, such as Wakimoto medium. The pigment was extracted with methanol and
was identified as a mixture of violacein and deoxyviolacein. This pigment could be used
to dye not only natural fibers like silk, cotton and wool, but also synthetic synthetic fibers
like nylon and vinylon, and generally gave a good color tone. The shade depended on the
material. Silk, cotton and wool showed a bluish-purple color, nylon a dark blue color, and
acetate a purple color (Shirata, et al., 2000).
Dyeing could be performed by a simple procedure consisting of either dipping in
the pigment extract or boiling with the bacterial cells. By changing the dipping time and
the temperature of the dye bath, shades ranging from light purple to deep bluish-purple
could be selected. The color fastness of the dyed material was about the same as that
materials dyed with vegetable dyes, but the color faded easily when the material was
exposed to sunlight. However, since the pigment can be mass-produced by culturing, if
these shortcomings could be overcome, the dye may become promising. The pigment
displayed an antimicrobial activity against phytopathogenic fungi like Rosellinia necatrix
which causes white root rot of mulberry. It could also be used as a bio-fungicide (Shirata,
et al., 2000).
2.5.6.3. Pigment in food industry
Use of microbial pigments in processed food is an area of promise with large
economic potential. However, microbial pigments offer challenges due to high cost,
lower stability and variation in shades due to changes in pH. At present, none of the
microbial pigments can replace synthetic pigment. The microorganisms such as
Monascus, Rhodotorula, Bacillus, Achromobacter, Yarrowia and Phaffia produce a large
number of pigments. An ideal pigment-producing microorganism should be capable of
using a wide range of C and N sources, have tolerance to pH, temperature and minerals,
and give reasonable colour yield.
Non-toxic and nonpathogenic nature of pigment-producing microorganisms
coupled with easy separation from the cell biomass is stressed. The various advantages of
producing pigments from microorganisms include independence from weather
conditions, easy and fast growth, colours of different shades and growth on cheap
substances. Studies revealed unstable, largely degradable and sensitive to heat, light,
acidity and water activity as characteristics of microbial colour. Improvement in stability,
safety and solubility can certainly make widespread use of microbial pigments in the food
industry (Joshi, et al., 2003).
In the recent years, colouring of food with pigments produced from natural
sources is of worldwide interest and is gaining importance. These pigments are looked
upon for their safe use as a natural food dye in replacement of synthetic ones because of
undesirable toxic effects including mutagenicity and potential carcinogenicity. Though
many natural colors are available, microbial colorants play a significant role as food
coloring agent, because of its flexibility in production and easy down streaming process.
Among the various pigment-producing microorganisms, Monascus is reported to produce
non-toxic pigments, agent it enhances the flavour of the food and acts a food preservative
(Vidyalakshmi, 2009).
Natural raw materials and by products of industry have wide use as culture media
in fermentation processes because of their low cost since the medium components can
represent from 38 to 73% of the total production cost. Rice is one of the major
agricultural products of economic importance. This primary product could serve as the
sustainable raw material for secondary value-added products through fermentation of
Monascus molds. Rice broken can be utilized for the production of these useful
microbial metabolites at an inexpensive manner and can be applied to varying food
products. In case of red pigment production by Monascus sp., the cost of the culture
medium cost is certainly a significant fraction of the total cost, because of the low cost of
the product and the lack of purification. The raw fermentation medium of semisolid
cultures of Monascus sp. in rice, for example, is used directly as the red pigment in foods
(Schmidell, 2001).
Nature is rich in colours (minerals, plants etc.) and pigment producing
microorganisms (fungi, yeasts and bacteria). Microbial pigments such as carotenoids,
melanins, flavins and quinines are used in many food industries (Dufosse, 2006). The
ability to synthesis pigments varies with the organism and its environment. Fluorescent
Pseudomonads are ubiquitous bacteria that are common inhabitants of the water, soil,
plants and animals and they are the most studied group.
Isolation and identification of Pseudomonas aeruginosa from marine environment
for phenazine pigment production. Pigment production results revealed that, the strain
could able to produce two different pigments namely, pyocyanin and pyorubrin.
Maximum biomass was observed at 66 h of incubation, but the pigment production seems
to be continuous throughout the culture period (72 h). Antibacterial activity of the
pigments was evaluated against pathogenic bacteria, maximum growth inhibitory activity
was observed with pyocyanin and pyorubrin at 20 µl concentration (1.7 and 1.3 cm,
respectively) against Citrobacter sp. Hemolytic activity of the pigments inferred that,
hemolysis was observed with both pigments at 15, 20 and 25 µl, concentration and no
hemolysis was found at 5, 10 and 15 µl. The pigments were evaluated for their potential
as food colourants with agar. Pleasant colouration was observed with pyocyanin and
pyorubrin at 25 mg mL -1
concentration. Pigments were used as food colourants and this
test was made to ensure the reliability of the pigments and their colouring ability at
boiling temperature in food materials (Jayalakshmi, 2008).
The controversial topic of synthetic dyes in food has been discussed for many
years. The scrutiny and negative assessment of synthetic food dyes by the modern
consumer have raised a strong interest in natural colouring alternatives. Nature is rich in
colours (minerals, plants, microalgae, etc.), and pigment-producing microorganisms
(fungi, yeasts, bacteria) are quite common. Among the molecules produced by
microorganisms are carotenoids, melanins, flavins, quinones, and more specifically
monascins, violacein or indigo. The success of any pigment produced by fermentation
depends upon its acceptability on the market, regulatory approval, and the size of the
capital investment required bringing the product to market (Dufosse, 2006).
A few years ago, some expressed doubts about the successful commercialization
of fermentation-derived food grade pigments because of the high capital investment
requirements for fermentation facilities and the extensive and lengthy toxicity studies
required by regulatory agencies. Public perception of biotechnology-derived products
also had to be taken into account. Nowadays some fermentative food grade pigments are
on the market: Monascus pigments, astaxanthin from Xanthophyllomyces dendrorhous,
Arpink Red from Penicillium oxalicum, riboflavin from Ashbya gossypii, β-carotene from
Blakeslea trispora. The successful marketing of pigments derived from algae or extracted
from plants, both as a food colour and a nutritional supplement, reflects the presence and
importance of niche markets in which consumers are willing to pay a premium for all
natural ingredients (Dufosse, 2006).
Recent increasing concern regarding the use of edible coloring agents has banned
various synthetic coloring agents, which have a potential of carcinogenicity and
terratogenicity. This circumstance has inevitably increased the demands for safe and
naturally occurring natural (edible) coloring agents, one of which is pigment from the
fungus Monascus purpureus. It has long been known that the microorganisms of the
genus Monascus produce red pigments, which can be used for coloring the foods.
Monascus pigments are a group of fungal secondary metabolites, called azaphilones,
which have similar molecular structures as well as similar chemical properties. The
pigments can easily react with the amino group containing the compounds in the medium
to form water-soluble pigments. Due to the high cost of the currently used technology for
the microbial pigment production on an industrial scale, there is a need for developing
low cost process for the production of the pigments that could replace the synthetic ones.
Utilizing cheaply available agro-industrial residues as substrate through solid-state
fermentation can attain such an objective (Sumathy Babitha, 2009).
2.5.7. Indian Scenario
In general, the scenario on research on microbial pigments at national level is
limited to very few reports over the years. The role of pigment Xanthomonasdin
produced by the genus Xanthomonas against photo damage was reported (Rajagopal, et
al., 1997). Barbhaiya and Rao, (1985a and 1985b) optimized the nutritional requirements
for pyoverdine production by Pseudomonas aeruginosa. Studies on a new generation
biocolours from Monascus sp. and related fungi for use as a colourant to the foods was
reported (Tamilselvan, 2001). Cellulose acetate membrane was used for the purification
and concentration of natural pigments (Chaudhuri, et al., 2004). A maroon dye was
extracted from the rhizome of Arnebia nobilis (Indrayan, et al., 2004). Effect of hexoses
and di-hexoses on the growth, morphology and pigment synthesis in the transformed root
cultures of red beet was studied (Ravishankar, 2004). A novel medium for enhanced cell
growth and production of prodigiosin from Serratia marcescens, isolated from soil was
optimized (Giri, et al., 2004).