J Biochem Tech (2012) 3(5): S203-S211
ISSN: 0974-2328
Optimization of lipase production from an indigenously isolated marine
Aspergillus sydowii of Bay of Bengal
Bindiya P*, Ramana T
Received: 21 August 2012 / Received in revised form: 5 November 2012, Accepted: 5 November 2012, Published online: 23 June 2013
© Sevas Educational Society 2008-2013
Abstract
A total of 44 marine sediments from 8 locations along the South
East Coast of Bay of Bengal were screened for lipolytic fungal
isolates by tributyrin agar clearing method and submerged
fermentation. Marine fungus BTSS 1005, isolated from Divipoint
location was identified as Aspergillus sydowii. Further studies
confirmed that BTSS105 produces higher extracellular lipase when
compared with other isolates. Efforts to increase the yield were
achieved by optimizing the production medium and growth
conditions, which would fully exploit the potential of the
microorganism. The optimum medium composition at 32°C, 8.0 pH,
80 rpm, 40% volume ratio, 96hr incubation time and 10% (v/v)
inoculum, under submerged fermentation was sucrose-2% (w/v);
ammonium chloride-3.5% (w/v); olive oil-3% (v/v) and tween 800.2% (v/v).
Keywords: Lipase production, lipolytic fungi, media optimization,
Aspergillus, marine sediment
Introduction
Lipases, (triacylglycerol acyl hydrolases, EC 3.1.1.3), are natural
catalysts of the hydrolysis of triacylglycerols into di- and
monoacylglycerols, fatty acids and glycerol at an oil–water
interface, a phenomenon known as interfacial activation (Schmidt
1998). However, under certain conditions, they are also able to
catalyze synthetic reactions (Carvalho 2006). The most reported of
the reactions carried out by these enzymes are hydrolysis,
acidolysis, alcoholysis, aminolysis, esterification and interesterification (Saxena 2003). Currently, lipases are a popular choice
as a biocatalyst because they can be applied to chemo-, regio- and
enantioselective hydrolyses and also in the syntheses of a broad
range of compounds (Jaeger 2002). These enzymes are considered
to have great potential as biocatalysts in numerous industrial
processes, such as the synthesis of food ingredients (Macedo 2003),
their use as additives to detergents (Liu 2009) and to obtain
enantiopure drugs and other refined products (Wang 2009). In the
Bindiya P*, Ramana T
Department of Biotechnology, C.S.T, Andhra University,
Visakhapatnam – 530003, Andhra Pradesh, India
*Email: [email protected]
chemical industry, they are used for the production of surfactants
and detergents, to resolve the racemic mixtures and for the treatment
of residues that are rich in oils and fats. In the health sector they are
used in medicines, diagnostics, cosmetics and antibiotics (Hasan
2006). In the food industry, lipases are used to synthesize
emulsifiers such as mono-and diglycerides (Kittikun 2008) and for
the production of lipids with high levels of polyunsaturated fatty
acids (Reshma 2008). They are also used for the development of
flavors (Salah 2007), the maturation of cheese (Dupuis 1993) and
sausage meat, among others. Furthermore, lipases have an important
application in the field of bioenergy, particularly for the production
of biodiesel (Park 2006), which is an expanding sector, given the
worldwide concern with the use of renewable energy.
Lipases occur in animals (Gangadhara 2009; Shan 2009), plants
(Paques 2008; 2006) and micro organisms (Melo 2005). Microbial
lipases show a broad spectrum of industrial application due to their
greater stability, substrate specificity and lower production costs
when compared to other sources. In addition, the immense
biodiversity of microorganisms improves their biotechnological
importance and justifies the search for new lipases. Following
proteases and carbohydrases, lipases are considered to be the third
largest group based on total sales volume. Numerous species of
bacteria, yeasts and molds were found to produce lipases. Marine
fungi are a promising source of novel bioactive compounds. Fungi
are one of the most important lipase sources for industrial
application because fungal enzymes are usually excreted
extracellularly, facilitating extraction from the fermentation media.
Filamentous fungi are recognized as the best lipase producers and
are currently the preferred sources (Carvalho 2005) A large number
of filamentous fungi have been studied for lipase production (Maia
2001;Mahadik 2002;Elibol 2001). Enhancing lipase production
during the cultivation process is also an important step in industrial
application of this enzyme. Different environmental factors have
been extensively studied to increase lipase productivity, such as
carbon sources nitrogen sources, oils as lipid sources, pH, and
temperature, and others (Maia 2001; Mahadik 2002; Lin 1996;
Dalmau 2000; Guerzoni 2002). The lipases obtained from the genus
Aspergillus present remarkable importance in biotechnological
applications. Moreover, many Aspergillus sp. lipases present several
properties of immense industrial importance, such as their pH and
temperature stability and excellent enantioselectivity.
S204
J Biochem Tech (2012) 3(5): S203-S211
Approximately 90% of all industrial biocatalysts are produced by
submerged fermentation, frequently using specifically optimized
media and genetically manipulated microorganisms. Many studies
have been undertaken to define the optimal culture and nutritional
requirements for lipase production. These requirements are
influenced by the type and concentration of the carbon and nitrogen
sources, culture pH and growth temperature, etc. (Elibol 2001).
clearance (Musantra 1992). After incubation, isolates were
distinguished from other microbial colonies by their morphological
features. Single separated colonies were selected and the sub
cultures were maintained on potato dextrose agar slants at 4ºC until
further use.
The ocean covers more than70% of Earths surface and is considered
as a great reservoir of natural resources. However, the extent of
marine biodiversity, especially of microorganisms, is barely known.
One of the least studied habitats of fungi is the deep sea that largely
remained neglected. The marine sediments as a source of bioactive
fungi was less exploited. One of the first reports of fungi in deep-sea
sediments was provided by (Raghukumar et al 1992) who isolated
fungi from calcareous sediments of the Bay of Bengal at a depth of
965m and demonstrated germination of spores of Aspergillus ustus
under simulated deep-sea conditions. Subsequently, cultivation of
marine yeasts (Lorenz and Molitoris 1992) and filamentous fungi
and germination of fungal spores (Zaunstock and Molitoris 1995)
under simulated deep-sea conditions of low temperature and
elevated hydrostatic pressure were reported. (Takami et al 1997)
showed the presence of fungi and yeasts in sediment samples
obtained from the Mariana Trench at a depth of 10,500m in the
Pacific Ocean. These were later identified to be Penicillium lagena
and Rhodotorula mucilaginosa, respectively (Takami 1999).
However, these have been sporadic reports and not comprehensive
enough to prove the existence of fungi in deep-sea sediments. The
aim of this study is to isolate a marine fungal isolate and analyze
different cultural parameters to obtain an enhanced lipase
production.
Detection of lipolytic fungi is done by tributyrin agar diffusion
method (Jani et al 1998). 20ml of tributyrin agar medium was
inoculated with a loopful of isolate and incubated at 28ºC for five
days. The composition of tributyrin agar medium is (g/l): (NH4)2SO4
5; Na2 HPO4 6; KH2 PO4 2; MgSO4 3; CaCl2 3; agar 20 and tributyrin
10ml with pH 6.0. Lipolytic zone of the isolates was measured and
these isolates were subjected to secondary screening.
Primary screening for lipolytic isolates
Secondary screening for lipase production
The selected isolates were cultivated in a synthetic medium
containing olive oil (source of natural triglyceride, triolein) as the
sole carbon source under submerged fermentation conditions and
assayed for the lipolytic activity of the culture filtrates. 45ml of
production medium is taken in 250ml Erlenmeyer flask and
inoculated with a loopful culture of each isolate. The flasks were
incubated at 28ºC for 4 days on a rotary shaker (120 rpm). The
culture broth was filtered and the clear filtrate was used as the
source of crude enzyme. The composition of the production medium
is (g/l): Olive oil 10; (NH4)2 SO4 5; Na2HPO4 6; KH2 PO4 2; MgSO4
3; CaCl2 3 with pH 6.0.
Lipase activity determination
All chemicals were analytical grade and all experiments were
carried out in triplicate.
The culture broth was filtered and the lipase activity in the culture
filtrate was determined by titrimetry (olive oil substrate emulsion
method) (Musantra 1992). One unit of enzyme activity is defined as
the amount of enzyme required to liberate 1µmole equivalent fatty
acid/ml/min at 30ºC under the standard assay conditions. All the
experiments were carried out in triplicate and the mean of the three
values was presented.
Sampling site and collection of sediments
Identification of the most promising isolate
A total of 44 marine sediment samples were collected from South
East Coast of Bay of Bengal with a gravity corer (66 cm length and
7 cm diameter) from 8 field stations on board Cruise No. 271 FORV
SAGAR SAMPADA scheduled from Kochi to Tuticorin. The
overlying water was siphoned out and the cores were cut at 2 cm
intervals down to 8 cm and extruded into alcohol sterilized clean
plastic containers and all samples were transported to the laboratory
for the isolation of lipolytic fungi. The maximum depth of collection
was 265m. The locations and depths of these sampling stations are
summarized at the Appendix.
The best lipolytic fungal isolate, BTSS-1005 was grown on various
types of media (Kornerup and Wanscher 1978) (Table: 4a) for
morphological studies. The media was adjusted to pH 6.0 and
sterilized by autoclave at 1200C for 15 min. The colonies were
observed after 7 days of cultivation at 28 ºC. The color names used
in this study were taken from the Methuen Handbook of Colour
(Kornerup and Wanscher 1978). Taxonomic characterization was
done following different guides (Alexopoulos 1979; Arx and Von
1974). The micromorphology of the isolate was studied by viewing
lacto-phenol cotton blue wet mount preparations. Scanning Electron
Micrograph was taken at Advanced Analytical Laboratory, Andhra
University, Visakhapatnam, India. Based on the data, the isolate
was assigned to the genus Aspergillus. Confirmation of the assigned
taxon was carried out by 18S rRNA gene sequence analysis at
Microbial Type Culture Collection, IMTECH, Chandigarh, India.
Partial 18S rRNA gene sequence thus obtained was submitted to
GenBank database at NCBI (GeneBank Accession No:JQ755254).
The identified strain Aspergillus sydowii was deposited in the
culture collection at Microbial Type Culture Collection, IMTECH,
Chandigarh, India (MTCC Ref No.34416).
Materials and Methods
Chemicals and Analysis
Isolation of lipolytic fungi from marine sediments
A portion of the sediment from the middle of each sub-section was
removed with a flame-sterilized spatula and placed in sterile vials
for isolation of fungi (Raghukumar et al. 2004). They were prepared
in seawater and fortified with Rifampicin 5µg/ml (Himedia,
Mumbai) to inhibit bacterial growth. All the media were used at 1/5
strength to simulate the low nutrient condition in the deep sea. The
soil sample (1 g) was suspended in 9 ml of sterile distilled water and
serial dilutions were made. Aliquots (0.1 ml) of appropriate
dilutions were surface plated on tributyrin agar plates and incubated
at 28ºC examining periodically upto 5 days. The fungal strains were
identified as lipase producers on tributyrin agar through zone of
J Biochem Tech (2012) 3(5): S203-S211
S205
Optimization of media and culture conditions
Secondary screening for lipase production
Lipase production in different media by submerged fermentation
All isolates were subjected to submerged fermentation conditions
and assayed for lipolytic activity quantitatively. Since tributyrin is
not a substrate for lipases alone, all positive isolates are confirmed
for their lipolytic activity by the hydrolysis of natural triglyceride
(triolein) under submerged fermentation conditions. As shown in
Table: 1, the isolate BTSS-1005 of DIV 3 sediment of Divipoint
location showed maximum lipolytic activity of 0.166U when
compared with other isolates. Optimization of nutritional and
physical parameters was done further to determine the potency of
BTSS-1005 at an industrial scale.
To study the biosynthesis of lipase by Aspergillus sydowii in
submerged fermentation, different production media reported by
various researchers were investigated (Table: 5a). Fermentation
experiments were run with 45ml each of the above described media
in 250ml Erlenmeyer flasks. The microorganism was grown and
maintained on potato dextrose agar medium for 7 days at 28ºC.
Spore suspensions for inoculation were prepared by adding 3 ml of
sterilized distilled water to fungal slant and vigorously shaking the
culture for 1 min. The number of spores was determined with a
Neubauer counting chamber and the inoculum was adjusted to 2.6 X
107 spores / ml. A 10.0% level of inoculum was used to initiate
growth and the inoculated flasks were kept on rotary shaker
(100rpm) at 28ºC for 5 days and 5 ml of each sample were
withdrawn at every 24h interval from 48h onwards from the flasks
for lipase assay.
Biomass estimation
The culture broth was filtered through preweighed Whatman No.1
filter paper. The filter paper containing the biomass was dried at
60ºC for 24h and its dry weight was estimated.
Optimization of lipase production
The optimization experiments were carried out in 250 ml
Erlenmeyer flasks containing 45 ml of culture medium. Shake flask
were seeded with inocula, at initial concentration of 10% , having
2.6 X 107 spores/ml, initial pH 6.0 which were incubated for 96 h at
28ºC under 120 rev/min on the initial cultivation medium cited
above. The following parameters were investigated sequentially to
optimize the production of lipase according to the experimental
design and the optimized parameters in each step were employed in
subsequent experiments. In order to further increase the yield of
enzyme, the effect of different pure compounds as carbon sources,
nitrogen source, different oils, and different surfactants on the
growth and enzyme production is studied. The effect of varying
temperatures, pH, agitation, incubation time, volume ratio and
inoculum level on the production of lipase was studied at different
ranges. Lipase production was investigated by employing all the
optimized parameters.
Results and Discussions
Isolation of fungal colonies from marine sediments
Using the selective media, 66 lipolytic fungal strains were isolated
(Table:1 (Column 2)).The samples of 7 locations are found suitable
for the isolation of lipolytic fungi. High or low number of active
strains found depends on many factors like the medium and methods
of screening. Moreover, there are so many factors which affect
fungal growth and enzyme production, including the chemical and
biological environment.
Primary screening for lipolytic fungi
There were no reports of lipases from marine fungi by previous
researchers although marine fungi have excellent lipolytic activity.
As indicated in Table 1, all isolates showed lipolytic activity after
primary screening. Screening with the help of tributyrin is a
convenient and presumptive test for the detection of lipolytic
organisms; hence all the isolates were screened using tributyrin agar
clearing method.
TABLE:1: Screening of lipolytic fungi
Sediment No.
Isolate
Lipolytic
No.
zone (R/r)
BTSS
KAK2
1001
1.80
1002
1.83
KAK3
1003
2.00
1004
1.83
KAK4
1005
2.72
1006
1.71
1007
1.45
1008
2.66
1009
1.66
1010
1.6
1011
1.66
1012
1.50
1013
1.80
DIV1
1001
2.72
1002
2.72
DIV2
1003
1.77
1004
1.83
DIV3
1005
1.77
1006
1.83
DIV4
1007
1.83
1008
1.83
1009
1.66
SIN2
1001
2.72
1002
1.90
1003
1.71
S1N4
1004
1.66
CHE3
1001
1.71
CHE4
1002
1.6
CUD1
1001
1.77
CUD3
1002
1.80
NAG2
1001
1.66
1002
1.57
NAG3
1003
1.55
NAG4
1004
1.80
VIS1
1001
1.80
1002
1.83
VIS2
1003
2.00
1004
1.66
VIS3
1005
2.23
1006
2.23
VIS4
1007
2.60
1008
1.63
1009
2.10
1010
1.36
1011
1.70
VIS5
1012
1.545
1013
1.390
1014
1.330
VIS6
1015
1.6
1016
2.33
1017
1.80
VIS8
1018
2.27
Lipase
activity (U/ml)
0.036800
0.036800
0.011000
0.036800
0.076300
0.076315
0.050000
0.016000
0.050000
0.116000
0.142100
0.150000
0.050000
0.07630
0.07630
0.12894
0.03680
0.16600
0.02368
0.03680
0.03680
0.05000
0.07630
0.12894
0.10260
0.05000
0.10260
0.11600
0.12894
0.05000
0.05000
0.01100
0.12894
0.06600
0.133000
0.036800
0.011000
0.050000
0.010000
0.010000
0.01850
0.03142
0.02140
0.03340
0.03142
0.02914
0.02620
0.04050
0.03850
0.02857
0.03142
0.02700
J Biochem Tech (2012) 3(5): S203-S211
S206
1019
2.63
0.03000
1020
2.13
0.01714
1021
1.88
0.05140
1022
2.428
0.03285
1023
1.82
0.04000
1024
4.0
0.02000
VIS9
1026
2.1
0.03142
1027
1.9
0.03285
VIS10
1028
1.625
0.03420
VIS13
1029
1.416
0.02770
VIS14
1030
1.857
0.04710
VIS15
1031
2.0
0.01428
1032
1.58
0.03050
Note: R: Hydrolyzed zone diameter; r: Growth zone diameter
Optimization of the medium composition for enhanced lipolytic
enzyme production using BTSS-1005 of DIV 3 isolate
The lipolytic fungal isolate was subjected to characterization. The
cultural and morphological properties of the isolate are shown in
Table: 2b.A. sydowi can be recognized by the characteristic blueTable 2a: Composition of media used for taxonomic investigation
S.
No.
1
2
3
Medium
Composition (g/l)
Potato dextrose agar
(M1)
Czapek dox agar (M2)
Potatoes infusion 200; Dextrose
20; Agar 15
Sucrose 30;Sodium nitrate 2;
Dipotassium phosphate 1;
Magnesium sulfate 0.5; Potassium
chloride 0.5; Ferrous sulfate 0.01;
Agar 15
Peptone 10; Dextrose 40; Agar,15
4
Sabouraud dextrose agar
(M3)
YEME Agar (M4)
5
Oat meal agar (M5)
6
Inorganic salts starch
agar (M6)
7
Glycerol-aspargine agar
(M7)
8
10
Peptone agar medium
(M8)
Tryptone yeast glucose
agar (M9)
Nutrient agar (M10)
*
Trace salts solution
9
Yeast extract 4; Malt extract 10;
Dextrose 4; Agar 20
Oat meal 20; Agar 20; Trace salt
solution 1ml
Starch 10; K2HPO4 1; MgSO4.
7H2O 1; NaCl 1; (NH4)2SO2 2;
CaCO3 2; Trace salts solution 1ml
L-asparagine 1; Glycerol 10;
K2HPO4 1; Agar 20; Trace salt
solution 1ml
Peptone 1; Agar 20; Sterile
Skimmed milk 10%
Yeast extract 4; Tryptone 10;
Dextrose 4; Agar 20
Peptone 5; Meat extract 3; NaCl 5;
Agar 20
FeSO4.7H2O 1; MnCl2.4H2O 1;
ZnSO4.7H2O 1
green color of its conidial heads on malt agar, the red-brown colors
in the substratum (especially on Czapek's agar), and the more
conspicuously echinulate and slightly larger conidia. The size and
arrangement of the conidial heads as well as the colour of the spores
they bear are important identifying characteristics. Species
identification depends upon pure cultures grown on known media.
Hence, its morphological properties must serve as the primary basis
of characterization. The results indicate that BTSS 1005 of DIV3
was characterized by its bluish green colonies, moderate growth,
reverse in shades of red, conidial heads radiate to nearly globose,
conidiophores colourless, smooth, vesicles globose, sterigmata in
two series, conidia globose to subglobose spores. Identification on
Potato dextrose agar (PDA) was Blue-green color, often reddish
exudate, reverse reddish, extremely rough conidia Colonies (CzA)
spreading, blue-green, with straw-coloured to reddish-brown shades,
often with abundant exudate; reverse usually reddish. Conidial
heads radiate. Vesicles spherical to subspherical, fertile over almost
the entire surface. Conidiogenous cells biseriate. Conidia echinulate,
green in mass, spherical to subspherical, exudate often abundant,
straw color to reddish brown shades; reverse usually in shades of
red, from coral red to maroon to almost black. Colonies on malt
extract agar growing more rapidly 4 to 5 cm. in 2 weeks, essentially
plane, with crowded conidial structures arising from the sub-merged
mycelium but characterized in varying degree by a loose network of
aerial hyphae overlaying the primary sporulating surface. Conidial
heads typical of the species are produced in greater abundance and
are characteristically more blue-green than on Czapek's agar;
exudate lacking; reverse uncolored to pale reddish maroon or with
the green color of the conidial heads apparent through the substrate.
The morphological and cultural features in all the media used for
study coincided with the features of A. sydowii reported earlier by
many researchers. On the basis of morphological and taxonomical
characteristics,(Table:2b, Fig.1.) the isolate BTSS-1005 of Divipoint
DIV3 sediment was identified as Aspergillus sydowii. This report
was confirmed after observing the scanning electron micrograph and
also sequencing studies on the isolate. While a few species of
Aspergillus, including A. sydowii, have been isolated from the ocean
before (Roth et al 1964; Sweeney et al 1976; Kendrick et al 1982;
Abrell et al 1996; Belofsky et al 1998; Raghukumar and
Raghukumar 1998; Toske et al 1998), they are not considered
normal inhabitants of the marine environment. This suggests that the
natural substrates such as marine sediments are also good sources
for isolation of lipolytic Aspergillus sydowii.
Table 2b: Morphological and cultural characteristics
Medium
Growth
Vegetative
mycelium
Aerial
myceliu
m
Yellow
Spore
Colour
Soluble
pigment
M1
Abundant
White
Greenish
Brown
Brown
Brown
Brown
Greenish
Black
Good
White
White
Reddish
brown
White
No
pigment
Brown
Purple
Brown
M2
M3
M4
Abundant
Abundant
Abundant
M5
Brown
M6
Good
White
Brown
Greenish
black
Black
M7
Good
Brown
Black
M8
Abundant
Reddish
brown
White
M9
Good
White
White
brown
White
M10
Moderate
White
White
Dark
brown
Dark
brown
Brown
No
pigment
No
pigment
Brown
Orange
red
No
pigment
No
pigment
Figure 1: Scanning Electron Microscope photograph of BTSS 105, DIV3,
isolated from Divipoint location.
J Biochem Tech (2012) 3(5): S203-S211
(Sharma et al 2001).The results indicated that sucrose at 2% (w/v)
concentration gave highest enzyme activity and biomass (Fig.2b).
Effect of Carbon sources
Carbon is the major component of the cell and the rate at which a
carbon source is metabolized can often influence the formation of
biomass or production of metabolites (Stanbury et al 1997) . Among
different carbon sources studied, the highest biomass and lipase
production was obtained with sucrose (Table: 6a, Fig.2a.) .Different
carbon sources were reported as ideal for maximum lipase
production using different bacteria and fungi. (Mahadik et al 2004;
Coastas et al 2004; Benjamin pandey 1996; Tan etal 2004; Kaimi et
al 1998; Corzo and Revah 1999; Dalmau 2000).The type and
concentration of carbon source may affect lipase biosynthesis
45
40
35
30
25
20
15
10
5
Glycerol
Mannitol
Starch
Lactose
Sucrose
Fructose
Xylose
Galactose
0
Glucose
Lipase activity and Dried biomass
50
Carbon sources (1 % w/v)
Effect of Nitrogen sources
Inorganic nitrogen sources can be used quickly, while organic
nitrogen sources can supply many cell growth factors and amino
acids which are needed for cell metabolism and enzyme synthesis.
(Tan et al. 2004).
Lipase activity U/ml
Dried biomass g/l
25
24.5
48
24
46
23.5
44
23
22.5
42
22
40
Dried biomass(g/l)
Lipase activity (U/ml)
50
21.5
38
21
0.50%
1%
1.50%
2%
2.50%
3%
Concentration of Sucrose(% w/v)
Figure 2b: Effect of Sucrose on enzyme activity and growth
Among various organic and inorganic nitrogen sources tested as
additives for lipase production ammonium chloride is found to be
the best source (Fig 3a) and the optimum concentration was found to
be at 3.5% (w/v) (Fig.3b.).
Lipase activity U/ml
Dried biomass g/l
60
50
40
30
20
10
P eptone
Tryptone
S oyabean
m eal
M alt extract
Y east extract
B eef extract
U rea
A m m onium
acetate
A m m onium
sulfate
A m m onium
chloride
A m m onium
nitrate
0
S odium
nitrate
TABLE:3b:Lipase activity (U/ml) in different reported media
S.No.
Medium
48hr
72hr
96hr
120hr
1
I
9.324
21.312
28.638
26.64
2
II
12.654
15.984
18.648
17.982
3
III
23.31
17.982
16.65
13.32
4
IV
5.328
10.656
14.652
13.986
5
V
6.66
9.324
8.658
7.992
6
VI
7.992
11.322
13.32
11.988
7
VII
4.662
5.994
9.99
6.66
8
VIII
1.998
2.664
3.33
1.998
9
IX
2.664
3.33
5.328
3.996
10
X
15.318
21.978
25.308
22.644
11
XI
17.316
25.308
26.64
23.31
Dried biomass g/l
Figure 2a: Effect of different carbon sources on enzyme activity and growth
Lipase activity and D ried biom ass
Table 3a: Optimization of lipase production in reported medium
S.
Medium
Composition (g/100ml)
No.
1
I
Dextrose 0.1; Peptone 0.2;Yeast extract 0.5;
(NH4)2SO4 0.5; MgSO4.7H20 0.02; FeSO4.7H2O
0.001; NaCl 0.5; Olive oil, 1.
2
II
Glucose 1; Yeast extract 0.3; Malt extract 0.3;
Peptone 0.5; Olive oil 2
3
III
Peptone 0.5; Malt extract 0.3; Yeast extract 0.3;
Glucose 2.0; Olive oil 1.
4
IV
Peptone 0.5; Yeast extract 0.1; NaNO3 0.05;
MgSO4.7H20 0.05; KCl 0.05; KH2PO4 0.2; Olive oil
1.
5
V
Glucose 2; Peptone 1; KH2PO4 0.6; K2HPO4 0.2;
KCl 0.1; MgSO4.7H2O 0.5; Tween 80 1; Olive oil 1
6
VI
Peptone 0.5; Glucose 1.0; KH2PO4 0.25; KCl 0.05;
MgSO4.7H20 0.05; Sunflower oil 1.5
7
VII
Yeast extract 0.1; CaCl2.2H20 0.01; MgSO4.7H20
0.05; Olive oil 1
8
VIII
Glycerol 1; Urea 0.4; KH2PO4 0.6; MgSO4.7H20 0.1;
FeCl3.6H2O 0.001; inositol 0.0000004; biotin
0.0000008; thiamine 0.00002; Olive oil 1.
9
IX
Glucose 0.4; NH4Cl 0.1; Na2HPO4.2H2O 0.7;
KH2PO4 0.3; NaCl 0.05; MgSO4.7H2O 0.025;
CaCl2.2H2O 0.002; Olive oil 1
10
X
Peptone 5.0; KH2PO4 1.4; K2HPO4 0.24;
MgSO4.7H2O 0.04; Olive oil 1
11
XI
Peptone 2.0; Glucose 1.0; KH2PO4 1.4; K2HPO4
0.24; MgSO4.7H2O 0.04; Olive oil 1
Lipase activity U/ml
Maltose
The fermentation experiments were conducted in different reported
media (Table: 3a) to select the best medium that induces maximum
enzyme production by A. sydowii. The results indicated Media-I
exhibited maximum lipase production at 96hrs. (Table: 3b.) Loss of
enzyme activity after peak production in each case was observed in
all media. This may be due to the cessation of enzyme synthesis
after the completion of growth followed by the deactivation and
autolysis of the enzyme. Some media could not support adequate
growth and lipase synthesis indicating that the organism might
require some growth factors and inducers.
P otassium
nitrate
Lipase production in different media by submerged fermentation
S207
Nitrogen sources (1%w/v)
Figure 3a:
growth
Effect of different nitrogen sources on enzyme activity and
Effect of different oils
The incorporation of lipids has been shown to increase lipase yields
in many cases (Silva et al 2005; Coastas et al 2004; Mahadik et al
2004; Sharma et al 2001). In this investigation, results indicated that
all lipidic substrates supported lipase activity and growth especially
Olive oil (Fig.4a.).Olive oil was also used successfully to increase
lipase production by several investigators using different
microorganisms. (Silva et al 2005;Thomas et al 2003; Cihangir and
J Biochem Tech (2012) 3(5): S203-S211
S208
Sarikaya 2004;Tan et al 2004;Sugihara et al 1990). Lipase
production in microorganisms is enhanced by varying not only the
lipid source but also its concentration (Silva et al 2005; Maia et al
26
25
40
24
30
23
20
22
50
40
30
20
10
0
Tween 20
Lipase activity U/ml
Gum arabic
Dried biomass g/l
63
28.5
28.45
Lipase activity (U/ml)
28.4
62
28.35
28.3
61.5
28.25
61
28.2
28.15
60.5
28.1
60
p
28.05
59.5
Dried biomass g/l
28
0.10%
60
0.20%
0.30%
0.40%
Concentration of Tween 80(%v/v)
50
Figure 5b: Effect of Tween 80 on enzyme activity and growth
40
Effect of incubation temperature
30
20
Incubation temperature was shown to effect lipase production. It
was another critical parameter that has to be controlled which varies
from organism to organism. In our investigation 32ºC was found to
the optimum (Fig.6.).Different optimal temperatures were reported
by many researchers for different fungi (Cihagir and Sarikaya 2004;
Gulati et al 1999; Thomas et al 2003; Essamri et al 1998).
10
Vippa oil
Tributyrin
Sunflower oil
Mustard oil
Sesame oil
Castor oil
Groundnut oil
Olive oil
Neem oil
Palm oil
Coconut oil
0
Oils(1%v/v)
Lipase activity U/ml
Figure 4a: Effect of different oils on enzyme activity and growth
29
60
58
28
27
56
54
26
52
50
25
24
48
46
23
44
Lipase activity (U/ml)
Dried biomass g/l
Dried biomass (g/l)
Lipase activity U/ml
62
28.8
65
64
63
62
61
60
59
58
57
28.6
28.4
28.2
28
27.8
27.6
20ºC
22ºC
1%
1.50%
2%
2.50%
3%
3.50%
4%
4.50%
5%
Concentration of Olive oil (%v/v)
Figure 4b: Effect of Olive oil on enzyme activity and growth
Effect of surfactants
The addition of surfactants to the culture medium has been shown to
increase the secretion of lipolytic enzymes in a number of
microorganisms, maybe due to the alteration of cell permeability
leading to increase in protein secretion or to surfactant effects on
cell bound enzyme (Coastas et al. 2004). A wide variety of
surfactants like Tweens, Triton, SDS, PEG and Gum Arabic have
been studied by different investigators and at different
concentrations (Coastas et al. 2004;Abdelfattah 2002;Muralidhar et
al 2001; Ferrer and Sola 1992; Odibo et al 1995; Lin et al 1995)
.Tween 80 was found to be the best surfactant in our study (Fig.5a.)
at a concentration of 0.2%(v/v)(Fig.5b.).
24ºC
26ºC
28ºC
30ºC
32ºC
34ºC
Temperature ( in celsius)
22
0.50%
Dried biomass g/l
Dried biomass (g/l)
Lipase activity and Dried biomass
PEG
62.5
2001; Lin 1996; Corzo and Revah 1999).Olive oil concentration was
found to influence lipase activity as reported by many researchers
(Mahadik et al 2004; Odibo et al 1995; Gulati et al 1999; Silva et al
2005).In our investigation lipase yield was found to be maximal at
3 %( v/v) concentration (Fig.4b.).
Lipase activity (U/ml)
SDS
Figure 5a: Effect of different surfactants on enzyme activity and growth
Figure 3b: Effect of Ammonium chloride on enzyme activity and growth
Lipase activity U/ml
Triton X
Surfactant (0.1%v/v)
Concentration of Ammonium chloride (%w/v)
p
Tween 80
5%
4%
4.50%
3%
3.50%
2%
20
2.50%
21
0
1%
10
60
Dried biomass (g/l)
50
1.50%
L ip as e activ ity a n d D ried b io m a s s
27
Dried biomass (g/l)
Dried biomass g/l
60
0.50%
Lipase activity (U/ml)
Lipase activityU/ml
Lipase activity U/ml Dried biomass g/l
70
Figure 6: Effect of temperature on enzyme activity and growth
Effect of initial pH
The change in pH influences the growth of the organism, effects
product stability and also induce morphological change in organism
and enzyme secretion (Gupta et al. 2003).The initial pH of the
medium influences lipase secretion in different microorganisms
(Cihangir and Sarikaya 2004; Tan et al 2004; Thomas et al 2003)
.pH 8 is found to be best in the present study (Fig.7.).
Effect of agitation
The agitation supplies greater aeration to the culture and also creates
conditions for greater availability of nutrients to the cells. The
decrease in lipase activity was observed at both lower and higher
J Biochem Tech (2012) 3(5): S203-S211
S209
29
28.5
28
27.5
27
26.5
26
25.5
pH 6.5
pH 7
pH 7.5
pH 8
pH 8.5
62
60
58
56
54
20%
25%
30%
35%
pH 9
optimum agitation was found to be at 80 rpm in our study (Fig.8.)
Dried biomass g/l
56
54
60rpm
70rpm
80rpm
90rpm 100rpm 110rpm 120rpm
Lipase activity (U/ml)
60
58
Dried biomass (g/l)
Lipase activity (U/ml)
62
29
28.5
28
27.5
27
26.5
26
25.5
25
2%
Figure 8: Effect of agitation on enzyme activity and growth
6%
8%
10% 12% 14% 16% 18% 20%
Figure 11: Effect of inoculum level on enzyme activity and growth
Lipase activity U/ml
Dried biomass g/l
28.8
28.6
28.4
28.2
28
27.8
Dried biomass (g/l)
65
64
63
62
61
60
59
58
57
27.6
96hrs
120hrs
144hrs
168hrs
Incubation period (hrs)
Figure 9: Effect of incubation period on enzyme activity and growth
Production of lipase under optimal conditions
Lipase production was investigated by employing all the optimized
parameters .It is evident from the result the lipase productivity
employing optimized fermentation conditions was 223.25% higher
than the lipase yield by using initial fermentation conditions.
Overall data indicates that a significant increase of lipase yield was
achieved by newly formulated production medium and optimized
cultural conditions.
Lipase activity U/ml
Lipase activity and Dried biom ass
The incubation period is an important factor in the rate of
production of extracellular lipases by the microorganism (Shirazi et
al. 1998). Lipolytic activity in the filtrate was detectable very early
in the incubation period (Fig. 9). The maximum lipase activity was
obtained after 96h of incubation in our study (Fig.9.) .
Lipase activity (U/ml)
4%
Level of inoculum (%v/v)
Effect of incubation period
72hrs
Dried biomass g/l
66
64
62
60
58
56
54
52
50
Agitation speed (rpm)
48hrs
50%
fermentation process since a lower inoculum density may give
insufficient biomass causing reduced product formation, while a
higher inoculum may produce too much biomass leading to the poor
product formation (Mudgetti 1986). The optimum level in this study
was found to be at 10 %( v/v) (Fig.11.).
Lipase activity U/ml
29
28.5
28
27.5
27
26.5
26
25.5
64
45%
Figure 10: Effect of Volume ratio on enzyme activity and growth
Figure 7: Effect of pH on enzyme activity and growth
66
40%
Volume of medium in 250ml flsk (%v/v)
Initial pH
Lipase activity U/ml
29
28.5
28
27.5
27
26.5
26
25.5
64
Dried biomass (g/l)
pH 6
Lipase activity (U/ml)
66
64
62
60
58
56
54
52
Dried biomass g/l
66
Dried biomass (g/l)
Dried biomass (g/l)
Lipase activity (U/ml)
Lipase activity (U/ml)
Lipase activity U/ml
Dried biomass (g/l)
agitation speeds in agreement with other reports (Thomas et al
2003;Cihangir and Sarikaya 2004;Nahas 1988;Rao et al 1993).The
70
60
50
40
30
20
10
0
Effect of volume ratio
Effect of production media volume on lipase production was studied
in 250ml flask from 20 to 50% ratio. The high productivity was
attained at volume of 100 ml in 250 ml flasks. (Fig.10). Lipase
activity gradually decreases by increasing volume. It was evident
from the results that aeration influenced lipase production (Cihangir
and Sarikaya 2004).
Effect of inoculum level
It is important to provide an optimum
inoculum
level in
Dried biomass g/l
Optimized
fermentation
conditions
Initial fermentation
condition
Fig :12: Lipase production employing optimized parameters
Conclusion
Based on the screening results it had been shown that sediments of
Bay of Bengal possess lipolytic fungi and maybe tapped as one of
the potential source for lipase production. The results in general
reflect on the lipase production potentiality among the relatively less
explored group of marine fungi. It is suggested that frequent and
systematic screening for fungi in the Bay of Bengal could provide
novel species as well as promising lipolytic fungi. Among scientists
S210
working on Aspergillus, there is a continuing fascination with their
biotechnological potential. In addition to producing numerous useful
extracellular enzymes and organic acids, these moulds also produce
secondary metabolites of importance in biotechnology. Marine
microorganisms, with their unique nature differ very much in many
aspects from their terrestrial counterparts and are known to produce
diverse spectra of novel and useful substances. Among the different
genera of fungi, Aspergillus is known as the dominating group for
industrial production of enzymes, particularly lipase. It is concluded
that A. sydowii BTSS1005 DIV3 isolated from marine sediment has
potential for use in industries for the production of extracellular
lipase. The strain can be ordered through MTCC, Chandigarh, India
(Strain name: Aspergillus sydowii MTCC 34416).
Reference
Abdel-fattah YR (2002) Optimization of thermo stable lipase
production from a thermophilic Geobacillus sp. using BoxBehnken experimental design Biotechnol Lett 24:1217—1222
Abrell LM, Borgeson B, Crews P (1996) Chloro polyketides from
the cultured fungus (Aspergillus) separated from a marine
sponge Tet Let 37: 2331–2333
Alexopolus CJ,Mims CW (1979) Introductory Mycology. 3rd edn.
Wiley Eastern Ltd.publishers,New Delhi,India
Arx JA, Von (1981). The Genera of Fungi Sporulating in Pure
Cultures, 3rd edn, J.Cramer, Germany.
Belofsky G, Jensen PR, Renner MK,Fenical W (1998). New
cytotoxic sesquiterpenoid nitrobenzoyl esters from a marine
strain of the fungus Aspergillus versicolor Tetrahedron 54:
1715–1724
Benjamin S, Pandey A (1996) Optimization of liquid media for
lipase production by Candida rugosa Bioresource Technol
55:167-70.
Cardenas F ,Alvarez E, deCastro Alvarez MS ,Sanchez Montero JM,
Valmaseda M, Elson SW, Sinisterra JV (2001) Screening and
catalytic activity in organic synthesis of novel fungal and yeast
lipases J Mol Cat .B Enzymatic 14:111-123
Carvalho PO, Contesini, FJ, Ikegaki M (2006) Braz J Microbiol 37:
329–337
Carvalho PO, Contesini FJ, Bizaco R,Macedo GA (2005) Food
Biotechnol 19: 183–192.
Cihangir N,Sarikaya E(2004) Investigation of lipase production by a
new isolate of Aspergillus sp World J Microbiol Bioechnol
20:193-197
Corzo G,Revah S(1999) Production and characteristics of the lipase
from Yarrowia lipolytica 681 Bioresour Technol 70:173-80
Costas M, Francisco JD,Maria AL(2004) Lipolytic activity in
submerged cultures of Issatchenkia orientalis Processs
Biochem 39:2109-2114
Dalmau E, Montesinos JL, Lotti M, Casas C(2000) Effect of
different carbon sources on lipase production by Candida
rugosa. Enzyme Microb Technol 26:657-663.
Dupuis C, Corre C, Boyaval P(1993) Lipase and esterase activities
of Propionibactenium freudenreichii subsp. freudenreichii.
Appl Environ Microbiol 59 (12): 4004–4009
Elibol M, Ozer D (2002) Response surface analysis of lipase
production by freely suspended Rhizopus arrhizus Process
Biochem 38:367–72
Elibol M, Ozer D (2001) Process Biochem. 36:325–329
Essamri M, Deyris V, Comcau L (1998) Optimization of lipase
production by Rhizopus oryzae and study on the stability of
lipase activity in organic solvents J Biotechnol 60:97-103
Ferrer P,Sola C (1992) Lipase production by immobilized Candida
rugosa cells Appl Microbiol Biotechnol 37:737-741
Gangadhara P, Ramesh Kumar V, Prakash J. (2009) Am. Oil Chem
Soc 86:773–781
J Biochem Tech (2012) 3(5): S203-S211
Guerzoni ME, Lanciotti R, Vannini L, Galgano F, Favati F, Gardini
F et al (2002) Variability of the lipolytic activity in Yarrowia
lipolytica and its dependence on environmental conditions Int J
Food Microbiol 69:79–89.
Gulati R, Saxena RK, Gupta R, Yadav RP, Davidson WS (1999)
Parametric optimization of Aspergillus terreus lipase
production and its potential in ester synthesis. Process Biochem
35: 459-464
Gupta R, Gigras P, Mohapatra P, Goswami VK, Chuhan B
(2003).Microbial amylases : a biotechnological perspective
Process Biochem 38:1599-1616
Hasan F, Shah AA, Hameed A (2006) Industrial applications of
microbial lipases Enzyme Microb Technol. 39 (2): 235–251
Jaeger KE, Eggert T (2002) Curr. Opin. Biotechnol. 13: 390–397
Kaimi NR, Mala JGS, Puvanakrishnan R(1998) Lipase production
from Aspergillus niger by solid-state fermentation using
gingerelly oil cake Process Biochem 33: 505-511
Jani TR, Patei RB, Sharma G, Shah DA, Dave SR (1998).Screening
of lipase using oil seed industry wastes J.Sci Ind.Res.57: 785789
Kendrick B, Risk M, Michaelides J,Bergman K (1982) Amphibious
microborers: bioeroding fungi isolated from live corals. Bull.
Mar. Sci. 32: 862–867
Kittikun AH, Kaewthong W, Cheirsilp B (2008) Continuous
production of monoacylglycerols from palm olein in packedbed reactor with immobilized lipase PS Biochem Eng J 40:
116–120
Kornerup A, Wanscher JH (1978) Methuen Handbook of colour. 3rd
edn, Methuen, London.
Lin SF (1996) Production and stabilization of a solvent-tolerant
alkaline lipase from Pseudomonas pseudoalcaligenes F-111 J
Ferment Bioeng 82: 448-451
Lin SF, Chiou CM, Tsai YC (1995)Effect of Tritonx-100 on
alkaline lipase production by Pseudomonas pseudoalcaligenes
F-111 Biotechnol Lett17: 959-62
Liu R, Jiang X, Mou H, Guan H, Wang H, Li X (2009) Biochem
Eng J 46: 265–270.
Lorenz R, Molitoris HP (1992) High-pressure cultivation of marine
fungi: cultivation experiments. In: Balny C, Hayashi R,
Masson, P. (Eds.) High Pressure and Biotechnology John
Libbey & Co London pp. 315–319
Macedo GA,. Lozano MMS, Pastore GM (2003) Electron. J.
Biotechnol. 6: 72–75
Mahadik ND, Kulbhushan BB, Puntambekar US, Khire JM,
Gokhale DV(2004) Production of acidic lipase by a mutant of
Aspergillus niger NCIM 1207 in submerged fermentation
Process Biochem 39:2031-2034
Mahadik ND, Puntambekar US, Bastawde KB, Khire JM, Gokhale
DV (2002) Production of acidic lipase by Aspergillus niger in
solid state fermentation. Process Biochem 38:715–21
Maia MMD, Heasley A, Camargo de Morais MM, Melo EHM,
Morais Jr MA, Ledingham WM et al (2001) Effect of culture
conditions on lipase production by Fusarium solani in batch
fermentation. Bioresour Technol 76:23–27
Melo LLM, Pastore GM ,Macedo GA (2005) Process Biochem
40:3181–3185
Melo LLM ,Pastore GM, Macedo GA (2005) Food Sci. Biotechnol
14:368–370
Mudgetti RE (1986) Manual of industrial biotechnology. American
Society for Microbiology, Washington DC.
Muralidhar RV, Chirumamila RR, Marchant R, Nigam P (2001)A
response surface approach for the comparision of lipase
production by Candida cylindracea using two different carbon
sources Biochem Eng J 9:17-23
Musantra A (1992) Use of lipase in the resolution of racemic
ibuprofen Appl.Microbiol.Biotechnol 38: 61-66
J Biochem Tech (2012) 3(5): S203-S211
Nahas E (1988) Control of lipase production by Rhizopus
oligosporus under various growth conditions J General
Microbiol 134:227-233
Odibo FJC, Okereke UO, Oyeka CA (1995) Influence of culture
conditions on the production of lipase of Hendersonula
toruloidea Bioresource Technol 54:81-83
Park EY, Sato M, Kojima S (2006) Fatty acid methyl ester
production using lipaseimmobilizing silica particles with
different particle sizes and different specific surface areas.
Enzyme Microb Technol 39: 889–896
Paques FW, Pio TF, Carvalho PO, Macedo GA (2008) Braz. J Food
Technol 11: 20–27.
Paques FW, Macedo GA, Quim Nova (2006) 29: 93–99
Raghukumar C, Raghukumar S, Sheelu G, Gupta SM, Nath BN,
Rao BR (2004) Buried in time: culturable fungi in a deep-sea
sediment core from the Chagos Trench, Indian Ocean. DeepSea Research Part-I 51: 1759–1768
Raghukumar C, Raghukumar S (1998) Barotolerance of fungi
isolated from deep-sea sediments of the Indian Ocean. Aquat.
microbiol. Ecol. 15: 153–163
Raghukumar C, Raghukumar S, Sharma S, Chandramohan D (1992)
Endolithic fungi from deep-sea calcareous substrata: isolation
and laboratory studies. In: Desai, B.N. (Ed.) Oceanography of
the Indian Ocean. Oxford IBH Publication, New Delhi
Rao PV, Jayaranam K, Lakshmanan LM (1993) Production of lipase
by Candida rugosa in solid state fermentation2:Medium
optimization and effect of aeration. Process Biochem 28:391395
Reshma MV, Saritha SS, Balachandran C, Arumughan C (2008)
Lipase catalyzed interesterification of palm stearin and rice bran
oil blends for preparation of zero trans shortening with
bioactive phytochemicals. Bioresour Technol 99 (11):5011–
5019
Roth F J, Orpurt P A, Ahearn DG (1964) Occurrence and
distribution of fungi in a subtropical marine environment. Can
J. Bot 42:375–383
Salah RB, Ghamghui H, Miled N, Mejdoub H, Gargouri Y (2007)
Production of butyl acetate ester by lipase from novel strain of
Rhizopus oryzae. J. Biosci Bioeng 103 (4):368–372
Saxena RK, Davidson WS, Sheron A, Giri B (2003) Process
Biochem. 39:239–247.
Schmidt RD ,Verger R, Angew (1998) Chem. Int. Ed. Engl.
37:1608–1633
Sharma RM, Chisti Y ,Banerjee UC (2001) Production, purification,
characterization, and applications of lipases Biotechnol Adv
19:627-662
Shan T, Wu T, Reng Y, Wang Y (2009) Anim. Genet. 40:863–870
Shirazi SH, Rahman SR, Rahman MM (1998) Production of
extracellular lipases by Saccharomyces cerevisiae.World J
Microbiol Biotechnol 14:595-597
Silva WOB, Mitidieri S, Schrank A,Vainstein MH (2005)
Production and extraction of an extracellular lipase from the
entomopathogenic fungus Metarhizium anisopliae. Process
Biochem 40:321-326
Stanbury PF, Whitaker A, Hall SJ (1997) Principles of Fermentation
Technology, 2nd ed Aditya Books (P) Ltd., New Delhi, India
Sugihara A ,Shimada Y, Tominaga Y (1990) Separation and
characterization of two molecular forms of Geotrichum
candidum lipase J Biochem 107:426-430
Sweeney JC, Migaki G, Vainik P M, Conklin RH (1976) Systemic
mycoses in marine mammals. J. am. vet. med. assoc. 169: 946948
Takami H (1999) Isolation and characterization of microorganisms
from deep-sea mud. In: Horikoshi, K., Tsujii, K. (Eds.)
Extremophiles in Deep-Sea Environments. Springer Tokyo pp:
3–26
S211
Takami H, Inoue A, Fuji F, Horikoshi K (1997) Microbial flora in
the deepest sea mud of the Mariana Trench. FEMS
Microbiology Letters 152:279–285
Tan T, Zhang M, Xu J, Zhang J (2004) Optimization of culture
conditions and properties of lipase from Pencillium
camembertii Thom PG-3. Process Biochem.39:1495-1502
Thomas A, Mathew M, Valsa AK, Mohan S, Manjula R (2003)
Optimzation of growth conditions for the production of
extracellular lipase by Bacillus mycoides. Ind. J. Microbiol
43:67-69
Toske S G, Jensen PR, Kauffman CA, Fenical W (1998)
Aspergillamides A and B: modified cytotoxic tripeptides
produced by a marine fungus of the genus Aspergillus. Tetrahed
54: 13459–13466
Wang PY, Chen YJ, Wu AC, Lin YS, Kao MF, Chen JR, Ciou JF,
Tsai SW(2009) Adv. Synth. Catal. 351:2333–2341
Zaunstock B, Molitoris HP (1995) Germination of fungal spores
under deep-sea conditions. Abstr. VI International Marine
Mycology Symposium, Portsmouth
tree, generally grown for medicinal and ornamental purpose could
now be promoted as a potential biofuel crop.
Appendix
Location of Sediment Samples
Sediment
Depth
Latitude
Longitude
Number
(meters)
VIS1
29.64
17°51.264N
83°32.060E
VIS2
265
17°50.814N
84°01.422E
VIS3
52.93
17°50.556N
83°01.228E
VIS4
1
17°46.515N
83°23.943E
VIS5
1
17°42.462N
83°19.234E
VIS6
10
17°30.500N
83°02.300E
VIS7
20
17°30.500N
83°04.400E
VIS8
30
17°30.500N
83°08.000E
VIS9
40
17°29.015N
83°07.480E
VIS10
30
17°29.000N
83°00.816E
VIS11
20
17°29.000N
83°02.205E
VIS12
30
17°29.000N
83°05.086E
VIS13
30
17°27.000N
83°01.314E
VIS14
20
17°27.000N
82°59.295E
VIS15
10
17°27.000N
82°59.398E
KAK1
201.17
16°59.832N
82°58.065E
KAK2
50.87
16°59.805N
82°32.497E
KAK3
108.05
16°59.507N
82°43.923E
KAK4
34.61
16°49.885N
82°25.665E
DIV1
31.28
15°59.943N
81°20.229E
DIV2
88.11
15°59.813N
81°24.737E
DIV3
191
15°59.813N
81°29.045E
DIV4
62.72
15°59.481N
81°22.716E
THA1
242
14°00.276N
80°26.946E
THA2
37.37
14°00.219N
80°20.747E
THA3
46.53
14°00.119N
80°21.586E
THA4
104
14°00.106N
80°24.532E
CHE1
29
13°08.792N
80°24.149E
CHE2
53.6
13°08.768N
80°26.478E
CHE3
99
13°08.490N
80°31.962E
CHE4
195
13°08.149N
80°35.251E
SIN1
192.60
15°00.551N
80°24.985E
SIN2
34.40
15°00.296N
80°12.826E
SIN3
56.19
15°00.199N
80°16.943E
SIN4
106.26
15°00.140N
80°20.764E
CUD1
100
12°12.519N
80°19.101E
CUD2
30
12°11.892N
80°05.725E
CUD3
53.26
12°11.718N
80°15.246E
NAG1
217
11°07.639N
80°07.236E
NAG2
107
11°07.511N
80°05.383E
NAG3
30
11°07.400N
79°57.829E
NAG4
50
11°07.152N
79°59.775E
VIS: Vishakhapatnam; KAK: Kakinada; SIN: Singarayakonda; DIV: Divipoint;
THA: Thamerapatnam; CHE: Chennai; CUD: Cudallore; NAG: Nagapatnam