Download Ranjit Kumar

ISOLATION OF LUMINESCENT BACTERIA
FROM BAY OF BENGAL AND THEIR
MOLECULAR CHARACTERIZATION
Alex Ranjith Kumar
This thesis comprises 30 ECTS credits and is a compulsory part in the Master of Science
with a Major in Resource Recovery – Industrial Biotechnology, 120 ECTS credits
No. 1/2010
Isolation of luminescent bacteria from bay of Bengal and their molecular characterization
Alex Ranjit Kumar, [email protected]
Master thesis
Subject Category:
Technology
University College of Borås
School of Engineering
SE-501 90 BORÅS
Telephone +46 033 435 4640
Examiner:
Elisabeth Feuk-Lagerstedt
Supervisor,name:
Dr. M. Jayaprakashvel
Supervisor,address:
Date:
Department of Biotechnology
AMET University (Under Sec. 3 of UGC Act 196)
Chennai-603112, India
Chennai-603112, India
Company/Organisation or street>
Department of Biotechnology, AMET University, Chennai-603112,
India
2010-09-08
Company/Organisation or street>
Keywords:
Sea water, luminescent bacteria, Bay of Bengal
Client:
ii
Acknowledgement
I am deeply indebted to GOD for the innumerable blessings He has showered on me. His
presence has been a constant throughout my life and has been a great source of solace. His
grace has enabled me to complete this project successfully.
I am very grateful to Elisabeth Feuk-Lagerstedt, SENIOR LECTURER, University of
Boras, Sweden she was truly what the word „ GUIDE’ significant , I revere for her
knowledgement and her help and support throughout my project period, without who I could
never have completed my project or writing thesis. Thanks a lot.
I extend a sincere thanks to Prof. Dr. A. Jaffar Hussain, SPECIAL OFFICER and DEAN,
Life Sciences, AMET University, Kanathur, Chennai for permitting me to work in his
laboratory.
I am grateful to Dr. M. Jayaprakashvel, Assistant Professor, Department of Biotechnology,
AMET University, Kanathur, Chennai for his unwavering support. He was always available
to offer ideas, clarifications and help in my project. His enthusiasm for research is infectious.
And I also thank Dr. R. Muthezhilan, Assistant Professor and HEAD i/c, Department of
Biotechnology, AMET University, Kanathur, Chennai for his constant guidance,
encouragement and support during my project.
I would like to thank Mr. K. Karthik, Visiting Faculty, Department of Biotechnology,
AMET University, Kanathur, Chennai for his support. I thank him for his help in clearing
certain doubts regarding my project whenever the need arose.
I extend my heartfelt thanks to the I and II M.Sc. students at Department of Biotechnology,
AMET University for their immense help, laughs, friendship and of course, great fun!! I
would once again extend my sincere gratitude to all the people involved in the successful
completion of this project. I extend my sincere thanks to Mr. C. Janahardas,
Phototechnician, CAS in Botany, University of Madras for his help in taking photos of
luminescent bacteria.
Lastly, I cannot forget the two most important people in my life, my parents, without whom I
would not have achieved so much and reached this stage in life. I am at a loss of words and
can only say “THANK YOU FOR BEING THERE FOR ME”
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Abstract
Luminescence is the emission of light by an object. Living organisms including certain bacteria
are capable of luminescence. Bacteria are the most abundant luminescent organisms in nature.
Bacterial luminescence has been studied most extensively in several marine bacteria. Bacterial
luminescence is due to the action of the enzyme called luciferase. The luminescent bacteria exist
in nature either as free living bacteria or in symbiotic association ship with certain marine
organisms. Research on luminescent bacteria has always been a fascinating one. In the present
study, twenty free living luminescent bacteria were isolated from Bay of Bengal, India using soft
agar overlay method in sea water complex agar (SWCA). All the 20 strains were characterized for
certain biochemical tests and they were tentatively identified that they are all Photobacterium spp.
The effect of salinity, pH glycerol concentration and heavy metals on the growth and
luminescence of these 20 strains was also studied. In this part of experiment, visual scoring was
done to categorize the luminescence. In case of salinity, it has been found that up to 6% of NaCl
the intense of luminescence was good and thereafter it declined. Further, in some strains it was
completely ceased beyond 9% of salinity. Luminescence was not greatly affected by pH in liquid
medium however; the same was affected in solid medium. The intensity of luminescence has
increased with increasing concentrations of glycerol ranging from 0.3 to 1.2%. All the 20
luminescent bacteria were characterized for their tolerance to heavy metals and antibiotics.
Copper and zinc at 1 mg/ml concentration have inhibited the growth and luminescence of the all
strains. Surprisingly, mercury at the same concentration has inhibited only two strains
(AMET1913 and AMET1920). However, at 2 mg/ml concentration mercury has inhibited the
growth and luminescence of all the 20 strains. Selected six luminescent bacterial strains were also
characterized for their antibiotic susceptibility against six different antibiotics. It has been found
that most of the strains were sensitive to all the six antibiotics tested. Since, the bioluminescence
is regulated by quorum sensing, the effect of culture filtrate extracted with dichloromethane was
also tested for its effect on luminescence. These DCM extracts haven‟t influenced the
luminescence much.
Keywords: Sea water, luminescent bacteria, Bay of Bengal
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Contents
1.
Introduction ...................................................................................................................... 1
1.1 Characteristics of Marine Microorganisms .................................................................. 1
1.2 Luminescent Bacteria ................................................................................................... 2
2. Literature Review ............................................................................................................. 4
2.1 Mechanism of Bioluminescence .................................................................................. 5
2.2 V. fischeri, the most studied luminescent bacterium .................................................... 7
2.3 Molecular characterization of luminescent bacteria .................................................... 7
2.4 Bioluminescence and environmental applications ....................................................... 8
2.5 Other applications of luminescent bacteria ................................................................ 10
3. Methods and Materials .................................................................................................. 10
3.1 Sample collection ....................................................................................................... 10
3.2 Isolation of marine luminescent bacteria ................................................................... 11
3.3 Agar overlay technique .............................................................................................. 12
3.4 Subculture of selected strains ..................................................................................... 12
3.5 Biochemical tests for the identification of the bacterium .......................................... 12
3.5.1
KOH string test: Gram‟s characterization ........................................................ 12
3.5.2
Simple staining for cell morphology ................................................................ 13
3.5.3
Test For Motility ............................................................................................... 13
3.5.4
Catalase Test ..................................................................................................... 13
3.5.5
Citrate Utilization Test ..................................................................................... 14
3.5.6
Indole Test ........................................................................................................ 14
3.5.7
The Methyl Red And Voges-Proskauer Tests .................................................. 14
3.5.8
Growth Of Luminascent Bacteria In Tcbs Medium ......................................... 15
3.6 Isolation Of Genomic DNA From Luminescent Bacteria .......................................... 15
3.7 Agarose Gel Electrophoresis ...................................................................................... 15
3.8 Effect Of External Factors On The Luminascence Of Luminascent Bacteria ........... 16
3.8.1
Effect of salinity (Varying conecntrations of NaCL) ....................................... 16
3.8.2
Effect of pH ...................................................................................................... 16
3.8.3
Effect of glycerol concentration ....................................................................... 16
3.8.4
Effect of heavy metals ...................................................................................... 16
3.9 Preparation of Autoinducers ...................................................................................... 17
3.10
Bioassay for autoinducers ..................................................................................... 17
3.11
Equipments used in the study ................................................................................ 17
4. Results and Discussion ................................................................................................... 18
4.1 Isolation of Marine Luminescent Bacteria ................................................................. 18
4.2 Subculture Of Selected Strains................................................................................... 18
4.3 Biochemical Tests For The Identification Of The Bacterium .................................... 18
4.3.1
KOH string test: gram‟s characterization ......................................................... 18
4.3.2
Simple staining for cell morphology ................................................................ 18
4.3.3
Test for motility ................................................................................................ 19
4.3.4
Catalase test ...................................................................................................... 19
4.3.5
Citrate utilization test........................................................................................ 19
4.3.6
Indole test.......................................................................................................... 19
4.3.7
The methyl red and voges-proskauer tests........................................................ 19
4.3.8
Growth of luminascent bacteria in TCBS medium........................................... 19
4.3.9
Tentative Identification Of The Strains ............................................................ 19
4.4 Isolation Of Genomic Dna From Luminascent Bacteria............................................ 20
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4.5 Effect Of External Factors On The Luminascence Of Luminascent Bacteria ........... 20
4.5.1
Effect of salinity ............................................................................................... 20
4.5.2
Effect of pH ...................................................................................................... 20
4.5.3
Effect of glycerol concentration ....................................................................... 21
4.5.4
Effect of heavy metals ...................................................................................... 21
4.5.5
Bioassay for Autoinducers ................................................................................ 22
5. Summery ......................................................................................................................... 23
6. Conclusion ....................................................................................................................... 23
References ............................................................................................................................... 24
Appendix 1
Appendix 2
Table 1 – Table 7
Figure 1 – Figure 12
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1. Introduction
A large amount of all life on Earth exists in the oceans. Exactly how large the proportion is
unknown, since many ocean species are still to be discovered. While the oceans comprise
about 71% of the Earth's surface, and taking this into account by volume, represent better than
95% of the biosphere (about 300 times the habitable volume of the terrestrial habitats on
Earth).
Marine life is a vast resource, providing food, medicine, and raw materials, in addition to
helping to support recreation and tourism all over the world. At a fundamental level, marine
life helps determine the very nature of our planet. Marine organisms contribute significantly
to the oxygen cycle, and are involved in the regulation of the Earth's climate. Shorelines are in
part shaped and protected by marine life, and some marine organisms even help create new
land.
Over the past decade, marine microorganisms have become recognized as an important and
untapped resource for novel bioactive compounds. Many species are economically important
to humans, including food fish. It is also becoming understood that the well-being of marine
organisms and other organisms are linked in very fundamental ways. The human body of
knowledge regarding the relationship between life in the sea and important cycles is rapidly
growing, with new discoveries being made nearly every day. These cycles include those of
matter (such as the carbon cycle) and of air (such as Earth‟s respiration, and movement of
energy through ecosystems including the ocean). Large areas beneath the ocean surface still
remain effectively unexplored.
Given this fact, the oceans present themselves as an unexplored area of opportunity for the
discovery of pharmacologically active compounds. However, it is important to pursue basic
research on the marine environment in order to permit the continued isolation of unique
microorganisms. There is still limited knowledge of the physiological requirements of most
marine microorganisms, and a greater understanding of their conditions for growth will offer
new insights into the complex world of marine microbiology. Clearly, a greater investment in
the development of marine biotechnology will produce novel compounds that may contribute
significantly towards betterment of human life in the coming years.
1.1 Characteristics of Marine Microorganisms
Since the marine environment is characterized by environmental conditions which are entirely
different from other environments, its microflora is also very much different from that of
others. Marine microflora has its own special characteristics which enable it to survive in that
environment. The main characteristic of marine flora is its capacity to survive and grow in
sea/ocean. Most of the bacterial flora in ocean is Gram-negative rods. The dominance of
Gram-negative bacteria in aquatic environment is due to their cell structure. Aquatic
environments are nutritionally dilute when compared with terrestrial environment. Under such
conditions the outer membrane, especially the lipopolysaccharide (LPS) of Gram-negative
bacteria helps to absorb nutrients.
Various important hydrolytic enzymes are retained within the periplasmic space. Thus, the
possible dilution of enzymes, which may occur when they are excreted out into the
environment, is prevented. In addition LPS gives protection against toxic molecules like fatty
1
acids and antibiotics. A larger proportion of the marine bacteria is motile and pigments
producers. Facultative aerobes dominate sea and there are relatively very few obligate aerobes
and obligate anaerobes. When cultured most oceanic bacteria grow more slowly and form
smaller colonies than those from other environments. Many of them are capable of
proteolysis. As with any other aquatic bacteria, marine bacteria can grow in extremely low
concentrations of nutrients and hence are called oligotrophic bacteria. Generally, the
concentration of organic matter in seawater will be less than mg per liter.
Other interesting features of marine microorganisms are their ability to survive at very low
temperature and at high salinity. The groups exhibiting the above characteristics are referred
to as psychrophiles and halophiles respectively. Marine bacteria are also characterized by
their pressure tolerance, especially those at depths. These forms belong to the group of
barophiles. The generation time of marine bacteria is quite long, ranging from less than one
hour to many months. The shortest generation time, 9.8 minutes, has been reported for
Pseudomonas natriegens at 37°C.
1.2 Luminescent Bacteria
Luminescence is the emission of light by an object. Living organisms including
certain bacteria are capable of luminescence. Bacteria are the most abundant luminescent
organisms in nature. Bacterial luminescence has been studied most extensively in several
marine bacteria (e.g.,Vibrio harveyi, Vibrio fischeri, Photobacterium
phosphoreum, Photobacterium leiognathi), and in Xenorhabdus luminescens, a bacteria that
lives on land. In luminescent bacteria, the general scheme involves an enzyme that is dubbed
“luciferase”. A suite of genes dubbed “lux” genes code for the enzyme and other components
of the luminescent system.
A similarity between the luminescent bacteria concerns the conditions that prompt the
luminescence. A key factor is the number of bacteria that are present. This is also known as
the cell density (i.e., the number of bacteria per given volume of solution or given weight of
sample). A low cell density (e.g., less than 100 living bacteria per milliliter) does not induce
luminescence, whereas luminescence is induced at a high cell density (e.g., 1010 to 1011 living
bacteria per milliliter)
Bacterial luminescence is due to the action of the enzyme called luciferase. Luciferase
catalyses the removal of an electron from two compounds. Excess energy is liberated in this
process. The energy is dissipated as a luminescent blue-green light The bacterial
luminescence reaction, which is catalyzed by luciferase, involves the oxidation of a longchain aliphatic aldehyde and reduced flavin mononucleotide (FMNH2) with the liberation of
excess free energy in the form of a blue-green light at 490nm:
FMNH2 + RCHO + O2 ----> FMN + RCOOH + H2O + light (490nm)
2
Animals can either house these substances in their own bodies or develop
a symbiotic relationship with light-producing bacteria. These bacteria live in a light organ in
the host organism's body. The bacteria produce light all the time, so in order to turn their
lights on and off, some animals can pull their light organs into their bodies. Others cover them
with pieces of skin similar to eyelids. Some organisms also use a fluorescent substance,
like green fluorescent protein, to adjust the color of the light they create. The fluorescent
substance absorbs the blue-green light and emits it as a different color.
Because of all these variations in luciferins, luciferases and how animals use them, many
researchers believe that the ability to make light simultaneously and independently evolved in
multiple forms of life. The fact that there are few bioluminescent animals in freshwater
environments supports this theory. Fresh, inland bodies of water haven't existed as long as the
world's oceans have, so the animals that live there haven't had as much time to adapt to their
surroundings. In addition, the bottoms of most bodies of fresh water aren't dark enough to
require additional sources of light.
Bacteria utilize homoserine lactone in other cell-to-cell communications that are cell-density
dependent. One example is the formation of the adherent, exopolysaccharide-enmeshed
populations, known as biofilms, by the bacterium Pseudomonas aeruginosa. Another example
is the bacterium Agrobacterium that produces diseases in some plants. The phenomenon has
been termed quorum sensing.
Gram-positive and Gram-negative bacteria use quorum sensing communication circuits to
regulate a diverse array of physiological activities. These processes include symbiosis,
virulence, competence, conjugation, antibiotic production, motility, sporulation, and biofilm
formation. In general, Gram-negative bacteria use acylated homoserine lactones as
autoinducers, and Gram-positive bacteria use processed oligo-peptides to communicate.
Recent advances in the field indicate that cell-cell communication via autoinducers occurs
both within and between bacterial species. Furthermore, there is mounting data suggesting
that bacterial autoinducers elicit specific responses from host organisms. Although the nature
of the chemical signals, the signal relay mechanisms, and the target genes controlled by
bacterial quorum sensing systems differ, in every case the ability to communicate with one
another allows bacteria to coordinate the gene expression, and therefore the behavior, of the
entire community.
The lux gene system responsible for bacterial luminescence has become an important research
tool and commercial product. The incorporation of the luminescent genes into other bacteria
allows the development of bacterial populations to be traced visually. Because luminescence
can occur over and over again and because a bacterium's cycle of luminescence is very short
(i.e., a cell is essentially blinking on and off), luminescence allows a near instantaneous (i.e.,
"real time") monitoring of bacterial behavior. The use of lux genes is being extended to
eukaryotic cells. This development has created the potential for the use of luminescence to
study eukaryotic cell density related conditions such as cancer.
Interesting is the fact, that luminescent bacteria do not produce light (or produce it very
weakly) when their cells are in considerable dispersion (e.g. in the sea-water). In contrast,
when many cells are in condensed suspensions (e.g. cultures growing in the liquid,
microbiological mediums) they produce light very efficiently. Previous researches show that
luminescent bacteria produce specific chemical compound - autoinducer which can induce
bioluminescence reactions in bacterial cells if present in environment in actual concentration.
When many bacterial cells are present in the environment, the concentration of autoinducer
3
grows and luminescence is induced very efficiently. Similarly, luminescent bacteria cells
which live in sea-water do not produce light but the cells which live in luminous organs of
marine animals produce it very effectively. It is possible because the quantity of bacterial
cells in these organs reach 1010 cells/ml. Other chemical compounds similar to autoinducer
were isolated from other bacteria species which do not produce light. Perhaps in this case
their role is informational. On the base of concentration of these compounds bacterial cell
"knows" how many other cells are in close nearness.
2. Literature Review
Some bacteria possess a unique ability to produce light and are commonly described as
phosphorescent, a term that implies that they absorb light energy, later releasing it when in the
dark. They are, however, more properly described as luminescent, a term that indicates that
they produce their own light. This ability may be lost just as virulence is sometimes lost on
continued artificial cultivation of pathogenic bacteria; but this ability is normally regained
rather readily if the organisms are cultivated on suitable media. This frequently means
cultivation on neutral or slightly alkaline media prepared from sea water or containing
equivalent amounts of the required salts.
Bacteria able to emit light are common in the marine environment (Baumann and Baumann,
1980). Luminous bacteria constitute a heterogeneous group of microorganisms, mainly
representing the family Vibrionacea. Luminous bacteria occur in the sea as free-living
organisms, as saprophytes, and as symbionts in light organs of certain fish and cephalopods.
The biochemistry of the light reaction associated with luminous bacteria has been extensively
studied, and many reviews on the topic have been published (Hastings, 1968; Cormier and
Totter, 1968; Cormier et al., 1975).
Luminous bacteria can be isolated readily from the marine environment. They have been
found as planktonic forms in seawater and associated with decaying animal material in the
benthos (Nealson and Hastings, 1979). Luminescent bacteria occur in the intestinal tracts of
marine animals (Ruby and Morin, 1979; O'Brien and Sizemore, 1979) and may be associated
with luminous fecal pellets (Raymond and DeVries, 1976). Lesions on the chitinous
exoskeleton of crustaceans can be caused by luminous bacteria (Baross et al., 1978). The
functioning of light organs of certain fishes and cephalopods requires colonization by
bacterial symbionts (Fitzgerald, 1977; Tebo et al., 1979; Hastings and Nealson, 1981).
Despite the wide range of habitats occupied by luminous bacteria, little is known about the
environmental factors affecting their populations (Yetinson and Shilo, 1979).
Squid use the light produced from the bacteria for a behavior known as counterillumination
(Young and Roper, 1977; Young et al., 1980; Jones and Nishiguchi, 2004). Luminescence
emitted from the light organ reduces the squid‟s silhouette to match the intensity and
wavelength of down-welling light (Young and Roper, 1977). This provides squid with a
mechanism that allows them to evade predators by camouflage. All bacteria housed in the
light organs are able to produce light via the lux operon both inside the light organs and in
their free-living state, although intensity of light and differences between strains of bacteria
have never been thoroughly investigated. Likewise, regulation of the lux operon inside the
light organ of squid has only been extensively studied in the Euprymna scolopes–Vibrio
4
fischeri symbiosis, where reduction in the amount of light produced affects symbiotic
competence (Visick et al., 2000).
The taxonomy of these bacteria has been investigated by several workers. Hendrie et al.
(1970) classified marine luminous bacteria into 4 species. Reichelt and Baumann (1973) in an
extensive survey of marine luminous bacteria also classified them into 4 species, and
extended their descriptions and provided more diagnostic traits for their identification Beneckea harveyi, B. splendida biotype I, Photobacterium fischeri, P. logei, P. phosphoreum,
and P. leiognathi. The 4 species of marine, luminous bacteria described in the latest edition of
Bcrgey's Manual (Buchanan and Gibbons, 1974) are: Photobacterium phosphoreum,
Photobacterium mandapamensis, Vibrio Non-Standard Abbreviation. PHB =
polyhydroxybutyrate fischeri, and Lucibacterium harveyi. These last 2 species are
synonymous with Photobacterium fischeri and Beneckea harveyi, respectively, in the work of
Reichelt and Baumann (1973). Photobacterium mandapamensis has been found by Reichelt
and Baumann (1975).
A simple set of diagnostic traits has been devised for the identification of these species (Bang,
S. et al., 1978) which has been recently applied in a number of ecological studies.
Identification of luminous bacteria is based on specific methods (Nauka, 1984; Williams and
Wilkins, 1977, 1984; Mir, 1997). For example, according to the kinetic characteristics of the
enzymatic reaction between luciferase and long-chain aldehydes, these enzymes fall into two
groups with fast and slow kinetics. Fast luciferases are synthesized by representatives of V.
fischeri, V. logei, Ph. phosphoreum, Ph. leiognathi, and A. hanedae. Luciferase with slow
kinetics was found in V. harveyi and V. splendida as well as in the fresh-water species V.
cholerae. This property is useful in distinguishing species of Photobacterium from Beneckea
as the decay kinetics of light emission by luciferase of the former genus have "fast" decay
kinetics while those of the latter genus have "slow" decay kinetics (Hastings et al., 1977).
Luminous marine isolates can be readily identified by application of a relatively few, simple,
diagnostic traits (Baumann, P., Baumann, L. 1983; Baumann, P. et al, 1983). This fact has led
to a number of ecological studies which have established the seasonal fluctuation of luminous
species in sea water (Ruby, E. et al. 1978), their vertical distribution in the water column
(Ruby, E. et al. 1980), and the species specificity of the symbiotic association between
luminous bacteria and marine animals (Nealson, K. et al 1977). Luminous bacteria were
identified in accordance with recent recommendations for identification of bacteria of the
family Vibrionacea (Williams and Wilkins, 1984) and prompt identification of photobacteria
(Nauka, 1984). The following parameters were assessed: morphology, Gram stain (Meditsina,
1973), growth characteristics, and bioluminescence of luminous bacteria at different
temperatures; β-polyhydroxybutyrate uptake by bacterial cells; sugar consumption as
measured by color changes of bromothymol blue, a pH indicator; and enzymatic properties
(Meditsina, 1973).
2.1 Mechanism of Bioluminescence
Bioluminescence is the product of two distinct enzymes, firefly luciferase and bacterial
luciferase. The application of the firefly enzyme in the study of mycobacteria has been
described by other groups (Jacobs et al., 1993). The bacterial luciferase enzyme is a dimer of
approx 80 kDa, consisting of α- and β-subunit (Meighen E A, 1991, Hastings J. W, 1978). It
catalyzes the oxidation of long-chain fatty acids (>7 carbons) and reduced riboflavin,
generating blue-green light (h = 490 nm) m the process.
5
The reaction can be written as follows:
FMNH2 + RCHO + O2 ------> f FMN + H20 + RCOOH + Light
The mechanisms of both of these bioluminescence reactions have been discussed in a recent
review (Grayski, 1987). In the firefly-luciferase reaction, the enzyme luciferase catalyses the
reaction between luciferin (substrate), adenosine triphosphate (ATP), and oxygen, which
leads to the emission of light. The marine bacterial bioluminescent system catalyzes a reaction
between oxygen, a reduced flavin phosphate, and an aldehyde (C, to C,, straight chain)
substrate which results in light emission. Both the firefly and the bacterial bioluminescence
systems have been extensively exploited for environmental monitoring purposes.
Fig. 1 The figure shows how bioluminescence works with help of luceferin and luceferase
Luminous bacteria use molecular oxygen to oxidize reduced flavin mononucleotide (FMNH2)
and a long chain aliphatic aldehyde to yield oxidized flavin mononucleotide, a carboxylic
acid, water and a photon (Hastings and Nealson, 1977). Synthesis of luciferase, the enzyme
that catalyzes this reaction, is subject to several controls. Most species of luminous bacteria
growing in a complex medium must secrete a sufficient concentration of an autoinducer
before luciferase synthesis can begin (Nealson et al., 1970; Eberhard, 1972; Nealson, 1977).
The synthesis of the luciferases of some strains is subject to catabolite repression (Nealson et
al., 1972; Ulitzur and Yashphe, 1975; Ulitzur et al., 1976) and is affected by the
phosphotransferase system (Lin et al., 1976) as well as by other systems (Waters and
Hastings, 1977). In a complex medium, the bacteria remove an inhibitor from the medium, the
presence of which inhibits the synthesis of luciferase (Kempner and Hanson, 1968). The
6
catabolite modulator factor of Ullman et al. (1976) is similar in many of its properties to this
inhibitor. Finally, it appears that oxygen somehow exerts a controlling effect, since some
strains produce maximal amounts of luciferase when they are growing in well aerated media,
while other strains give maximal yields only when the oxygen tension is quite low (Nealson
and Hastings, 1977).
The genes coding for the a- and P-subunits of luciferase, luxA and luxB, have been cloned
from a number of species of marine bacteria, such as Vibrio harveyi, Photobacterium (Vibrio)
fischeri, and Xenorhabdus luminescens (Stewart et al, 1992). This has led to the exploitation
of luciferase as both a measure of cell viabilty, owing to the requirement for cellular-reduced
flavin, and as a reporter gene for the study of promoter activity and gene regulation (Gordon
et al, 1991). The bacterial luciferase genes, with their phenotype of light production, represent
ideal candidates for a reporter system (Meighen, 1991).
The ability of luminous bacteria to emit visible bioluminescence is closely associated with
cell metabolism. This ability underlies the use of luminous bacteria as bioindicators. Various
assays and test methods were developed on the basis of live wild-type or mutant cells of
luminous bacteria, isolated luciferase preparations, and genes of the bioluminescence system
(Santa Maria A et al, 1998; Virta et al, 1998; Aruldoss et al, 1998).
2.2 V. fischeri, the most studied luminescent bacterium
V. fischeri is a Gram-negative bacterium which occurs both free-living and as a symbiont in
the light organs of certain species of fish and squid where it emits light via a process known
as bioluminescence (Nealson, 1977). The onset of bioluminescence in V. fischeri is
characteristically cell density-dependent and Nealson et al. (1970) recognized that spent
culture supernatant from high cell density cultures of V. fischeri, contained a substance (an
„autoinducer‟) which induced bioluminescence when added to cultures of low cell density.
When grown in a closed environment, as found within the light organ or in a laboratory
culture flask, the autoinducer accumulates culminating in the induction of bioluminescence
(Nealson, 1977). Conversely, when free-living, the cell density is low, autoinduction does not
occur and consequently V. fischeri is dark. Therefore, it is the accumulation of autoinducer to
a critical threshold concentration, rather than cell density itself, which triggers the enormous
amplification in light emission observed. The V. fischeri pheromone was later identified as N(3 oxohexanoyl)homoserine lactone ( Eberhard et al. (1981). The V. fischeri structural and
regulatory genes necessary for light production, OHHL synthesis and regulation (the lux
regulon) were located on a 9 kbp DNA fragment (Engebrecht & Silverman, 1984).
2.3 Molecular characterization of luminescent bacteria
The isolation of genomic DNA from a microorganism generally comprises three stages:
cultivation of the cells, disruption to release cell contents, and chemical purification of the
DNA. Two widely used methods for the preparation of bacterial DNA are those described by
Marmur, J., 1961 and Kirby, K. S., 1964, but procedures are frequently modified to suit the
7
particular organisms under study. The best DNA isolation techniques produce good yields of
pure, high molecular weight, largely double-stranded DNA. It may be problematic to obtain
sufficient DNA from some bacteria if they are difficult to grow or to break open, or if they
have small genome sizes. The DNA should be free of contaminating macromolecules, such as
RNA, protein, polysaccharide and chemical compounds, and also any residual plasmid DNA.
Some breakage of the DNA is inevitable during cell lysis and chemical purification, but care
should be taken to minimize mechanical shearing, and to inhibit enzymic degradation,
principally by deoxyribonucleases (DNAses). Fortunately, the latter enzymes are heat labile
and are readily inhibited by a lack of magnesium ions. DNAse activity could result in single
strand breaks or nicks, weakening the DNA strands and making them more susceptible to
shearing and hence to a reduction in molecular weight. Low molecular weight DNA
precipitates as fibers less readily than DNA of high molecular weight, so native DNA with an
intact double helical structure is the required end product.
In the past, identification of marine luminous bacteria based only on a phenotypic approach
was often equivocal because of their highly variable phenotypes. Recent molecular
techniques, including hybridization with luxA probes (Baumann, P., Baumann, L. 1980) and
analysis of protein-coding sequences (Nealson and Hastings, 1977), improved identification
but revealed certain limitations. Hybridization with luxA probes, for example, was highly
speciesspecific for the majority of marine luminous isolates except for V. harveyi. Indeed, the
V. harveyi probe showed cross-reactivity with luxA genes of two closely related species: V.
vulnificus and V. orientalis . Two V. vulnificus biotypes are known thus far (Nealson et
al,1979), biotype 1, a clinical strain, is an opportunistic human pathogen; biotype 2, an
environmental strain, is primarily an eel pathogen, but also an opportunistic pathogen for
humans( Baumann, L., Bang, S.S., Baumann, P. 1980). Some V. harveyi and V. splendidus
strains were recognised pathogens for penaeid larvae and snooks in hatcheries, and thus
careful recognition of these species in sea water samples is necessary. A molecular approach
was also used for determining phylogenetic relationships among type strains of the family
Vibrionaceae on the basis of small subunit rRNA gene sequencing. Comparison of the 16S
rDNA sequences led to construction of a phylogenetic tree for the genus Vibrio and related
genera.
2.4 Bioluminescence and environmental applications
The bioluminescence literature from 1980 to mid-1994 has been reviewed for environmental
applications of bioluminescence measurements. Immunoassay methods which may use
fluorescence, phosphorescence, or in vitro bioluminescence for quantitation were not
addressed. Methods that are principally applicable to medical diagnosis were also not
included unless there was a clear connection to environmental monitoring. The focus was on
in vivo and in vitro bioluminescence methods which have been utilized to elucidate
environmental properties of chemicals, their toxic and mutagenic effects, and to estimate
biomass. The unifying theme in this review was the application of bioluminescence to
environmental monitoring, remedial investigations, and toxicity assessments, and potential
field methods.
8
Luminescence is an easily recognized characteristic for ecological studies; colonies are
readily recognized and counted. One of the principal objectives of environmental monitoring
is to estimate the real danger of contaminants to humans and other organisms. A chemical
analysis of environmental samples may provide a measurement of the total concentration of a
potentially harmful substance; however, these total concentrations may not represent the true
potential impact of the substance to the biota. For example, chemical analysis may involve
extensive extraction from the environmental matrix (i.e. groundwater, soil, etc.). All of the
material detected after sample processing may not, in fact, be available to the target human,
animal, or plant populations. Thus, the measurement of the concentration of a compound in
soil or water does not give a complete assessment of exposure potential because measurement
of total contaminant concentrations does not necessarily reflect how much of a potentially
toxic substance is actually available in a form which can be transported into cells where they
may damage enzymes or deoxyribose nucleic acid (DNA). This issue has been addressed by
Blaise (1991) who has discussed the role of small scale microbiotests in aquatic toxicology.
Bioluminescence has been observed in various insects, fish, and bacteria (Campbell, 1989;
Hastings, 1986). For many of the organisms, the biochemical mechanisms of light emission is
reasonably well understood. For several organisms, the genes coding for the various enzymes
needed for light emission have been mapped, isolated, and cloned (Meighen, 1988). This
technology has been used to transfer bioluminescent properties to normally nonbioluminescent organisms (Ow et al, 1986; De Wet et al, 1987). Bioengineering of the
genetic codes which are responsible for bioluminescence can also have potentially important
environmental applications.
Thomulka et al, (1992, 1993) have discussed the use of P. phosphoreum and V. harveyi to
detect biohazardous chemicals in water. Delistraty (1984) examined the use of the Microtox
assay for assessing the toxicity of synthetic fuel by-product water. Yates et al., (1986) used
Microtox to assess the effects of selenium on the toxicity of cigarette-smoke condensate.
These measurements indicated that selenium could reduce acute toxic effects of cigarette
smoke.
McFeters et al., (1983) utilized the Microtox assay and a two-organism test (algae and
bioluminescent bacteria) for the detection of aquatic toxicants. The two-organism test which
was developed by Tchan et al., (1975) uses P. phosphoreum to monitor the production of
oxygen by the test algae. A toxic effect to either the algae or the bacteria will lead to a
decrease in bioluminescence. For example, a decrease in algal photosynthesis is accompanied
by a decreased production of 4. Since O2 is a necessary substrate for the microbial
bioluminescence reaction, a decrease in luminescence will be observed as a result of toxic
effects on the algae. The results of this study indicated that for a wide variety of toxicants,
there is a good correlation between the two methods. However, the Microtox test was
generally more sensitive than Tchan‟s test, with the exception of some toxicity measurements
with penylurea herbicides (which are especially toxic to algae).
Application of the bacterial lux system to generate light-emitting eukaryotes including yeast
has met with far less success primarily because of difficulties in supplying the flavin and
aldehyde substrates in vivo necessary for the luminescence reaction. Earlier studies have
shown that a fused bacterial luciferase from Vibrio harveyi (luxA-B) can be effectively
expressed in yeast but luminescence was relatively low on addition of decanal (M. Boylan,.
O. Olsson,et al; 1989) The low level of luminescence arose as FMNH2 is not sufficiently high
in the cytoplasm of eukaryotes to saturate the bacterial luciferase; much higher levels are
believed to be present in the mitochondria. Moreover, decanal, the aldehyde used in all
bioassays with bacterial luciferase, has been found to be lethal to yeast, leading to the
9
conclusion that the bacterial luxAB reporter genes cannot be used in these model eukaryotic
organisms (R.P. Hollis, et al; 2001)at least for continuous monitoring of the same cells.
2.5 Other applications of luminescent bacteria
A biosensor is an analytical device that consists of an immobilized biological material in
intimate contact with a compatible transducer, which will convert the biochemical signal into
a quantifiable electrical signal (Gronow 1984). Biosensors are the offspring of the first
successful marriage between biotechnology and modern electronics. The biomolecules are
responsible for the specific recognition of the analyte whereas the physicochemical transducer
supplies an electrical output signal which is amplified by the electronic component (Scheller
& Schubert 1992). The specificity of enzymes is the main reason for their use in biosensors.
Since most of the enzymes employed for use in sensors have been isolated from
microorganisms, it is logical that the organisms themselves should be regarded as potential
biocatalysts (Aston & Turner 1984). In microorganisms, theenzymes remain in their natural
environment,increasing stability and activity (Guilbault 1984; Corcoran & Rechnitz 1985;
Luong et al. 1988; D‟Souza 2001; Verma & Singh 2003). Cell membranes and organelles can
also be used for biosensor construction (Burstein et al. 1986, Verma & Malaku 2001).
Specific binding between antibody and antigen can be exploited in immunobiosensors. To
detect very low concentrations of substances such as drugs, toxins or explosives, receptorbased sensors are very appealing (Prasad et al. 2004).
In this scenario, the present work has been aimed to isolate marine luminescent bacteria and
to characterize them using routine and some molecular tools. It has also attempted to
determine the cellular interaction between them and the influence of environmental
parameters on the growth and luminescence of the isolated luminescent bacteria.
3. Methods and Materials
3.1 Sample collection
100 ml of sea water was collected from the intertidal zones of Kovalam, Kanathur, Muttukadu
in Chennai using sterile containers and brought to the laboratory for further processing (Fig. 1
map; Fig. 2 Sampling a and b). A total of four samples collected and the details are furnished
below
10
Sl.No
Sample
Location
Number of samples
collected
1
S1
Kovalam
1
2
S2
Kovalam
1
3
S3
Kanathur
1
4
S4
Muttukadu
1
3.2 Isolation of marine luminescent bacteria
Serial dilution plating technique
Composition of sea water complex agar (SWCA)
Peptone
Yeast extract
Glycerol
Agar
50% Sea water
5 g
3 g
3 ml
15 gram
It is one of the several non-selective media useful in routine cultivation of microorganisms. It
can be used for the cultivation and enumeration of bacteria which are not particularly
fastidious. Peptone provides the essential nutrients for growth: nitrogen, vitamins, minerals
and amino acids. It forms the principle sources of organic nitrogen, particularly amino acids
and long chained peptides. Yeast extract provide the necessary water soluble substances like
nitrogen compounds, carbon, vitamins and also some trace ingredients necessary for the
growth of bacteria. Glycerol is a carbon source which has great influence on the
luminescence of microorganisms. Bacteriological agar is the solidifying agent.
The four different sea water samples which were collected from three different places were
subjected to serial dilution. 10 ml of sea water sample was mixed with 90 ml of sterile
distilled water in a 250 ml flask to obtain 10-1. 1 ml from this dilution was taken and added to
another 9 ml of sterile distilled water in test tubes fro 10-2 and repeated once similarly to get
10-3 dilution. 0.1 ml from the10-3 dilution was used to spread plate in SWCA medium in Petri
plates. The plates were then incubated for 24 hrs and at every six hours the appearance of
luminescent colonies were observed.
11
3.3 Agar overlay technique
Composition of soft SWCA
Composition
Per (50 ml)
Peptone
0.25g
Yeast extract
0.15g
Glycerol
0.15 ml
Agar
0.3gram
One ml of the previously serially diluted sea water sample (from the10-3 dilution) was takedn
and mixed with 10 ml of soft SWCA and mixed well. Then, this suspension was poured over
pre-solidified regular SWCA plates and kept for incubation at room temperature incubated for
24 hrs and at every six hours the appearance of luminescent colonies were observed.
3.4 Subculture of selected strains
The distinct isolated luminescent colonies of bacteria were marked while observing for
luminescence and were further purified by sub-culturing in SWCA plates. Each such isolate
pure colonies of bacterium were given unique accession number starting with a prefix of
AMET indicating the institute name. Strain numbers AMET1901- AMET1920 were given
and these bacterial strains were stored in sterile sea water in Eppendorf tubes at 4ºC.
Whenever needed, subcultures were made from these stock cultures in SWCA. In all the
experiments, 24 h old bacterial cultures grown in SWCA were used, unless otherwise stated.
3.5 Biochemical tests for the identification of the bacterium
3.5.1 KOH string test: Gram’s characterization
The Gram Nature of the selected strains was determined by performing the KOH
string test (Ciufecu, C. et al, 1986).
This test is an alternative to the classic Gram staining procedure and exploits differences in
the cell wall of bacteria to help determine their Gram character. The reaction depends on the
lysis of the gram-negative cell in the dilute alkali solution releasing cellular DNA to turn the
suspension viscous. This test has the advantage of simplicity, and it can be performed on
older cultures. This can serve as a valuable adjunct to the traditional gram stain method (von
Graevenitz and Bucher 1983).
3% KOH solution was prepared by dissolving 3 g of Potassium Hydroxide in 100 ml of
distilled water. The 3% KOH String Test was done using a drop of 3% Potassium Hydroxide
on a clean grease free glass slide. A visible loopful of cells from a single, well-isolated colony
is mixed into the drop. If the mixture becomes viscous within 45 seconds of mixing and
12
produce a string when lifted using the loop (KOH-positive) then the colony is considered
gram-negative.
3.5.2 Simple staining for cell morphology
Procedure:
- Take a clean grease free glass slide.
- Prepare thin smear of the test bacterial isolate.
- Air dry and Heat fix the smear.
- Cover with Saffranin for 1 minute.
- Drain the dye and rinse under running water.
- Allow to air dry.
- Observe under oil immersion objective 100 x and note the shape of the bacterial
colonies.
3.5.3 Test For Motility
Hanging Drop technique
In this technique, a drop of medium containing cells to be observed is allowed to hang in the
cavity of slide. The advantage of this preparation over the wet mount preparation is the
increased capacity of aeration as the drop is surrounded by an air space. This is the best
method available for the routine use to observe the motility of bacteria. This is because, it is
relatively easy to make and less time consuming. It is essential to differentiate true motility
from the Brownian movement of bacteria. In true motility, the organism changes its position,
while in the Brownian movement; the organism oscillates at its place and does not change the
position in the field.
3.5.4
Catalase Test
The catalase test is a test for the presence of the catalase enzyme. Most organisms posess this
enzyme capable of breaking down hydrogen peroxide. Organisms containing the catalase
enzyme will form oxygen bubbles when exposed to hydrogen peroxide.
Procedure:
1. Place a drop of 3% hydrogen peroxide onto a clean microscope
slide.
2. Touch an isolated colony with an inoculating loop
3. Place the loop, carrying some of the isolate, into the drop of
hydrogen peroxide.
4. Observe the slide for the evolution of bubbles
5. The reaction is positive if oxygen bubbles form rapidly.
13
3.5.5 Citrate Utilization Test
The citrate test utilizes Simmon's citrate medium to determine if a bacterium can grow
utilizing citrate as its sole carbon and energy source. Simmon's media contains bromthymol
blue, a pH indicator with a range of 6.0 to 7.6. Bromthymol blue is yellow at acidic pH's
(around 6), and gradually changes to blue at more alkaline pH's (around 7.6). Uninoculated
Simmon's citrate agar has a pH of 6.9, so it is an intermediate green color. Growth of bacteria
in the media leads to development of a Prussian blue color (positive citrate). Enterobacter and
Klebsiella are citrate positive while E.coli is negative
Procedure:
1. Prepare slants and allow solidifying.
2. Streak the test cultures and incubate for 24 hours.
3. Change in color of medium from green to blue indicates citrate utilization.
3.5.6 Indole Test
The test organism is inoculated into tryptone broth, a rich source of the amino acid
tryptophan. Indole positive bacteria such as Escherichia coli produce tryptophanase, an
enzyme that cleaves tryptophan, producing indole and other products. When Kovac's reagent
(p-dimethylaminobenzaldehyde) is added to a broth with indole in it, a dark pink color
develops. The indole test must be read by 48 hours of incubation because the indole can be
further degraded if prolonged incubation occurs. The acidic pH produced by Escherichia coli
limits its growth.
3.5.7 The Methyl Red And Voges-Proskauer Tests
The methyl red (MR) and Voges-Proskauer (VP) tests are read from a single inoculated tube
of MR-VP broth. After 24-48 hours of incubation the MR-VP broth is split into two tubes.
One tube is used for the MR test; the other is used for the VP test.
MR-VP media contains glucose and peptone. All enterics oxidize glucose for energy;
however the end products vary depending on bacterial enzymes. Both the MR and VP tests
are used to determine what end products result when the test organism degrades glucose. E.
coli is one of the bacteria that produce acids, causing the pH to drop below 4.4. When the pH
indicator methyl red is added to this acidic broth it will be cherry red (a positive MR test).
Klebsiella and Enterobacter produce more neutral products from glucose (e.g. ethyl alcohol,
acetyl methyl carbinol). In this neutral pH the growth of the bacteria is not inhibited. The
bacteria thus begin to attack the peptone in the broth, causing the pH to rise above 6.2. At this
pH, methyl red indicator is a yellow color (a negative MR test)
14
3.5.8 Growth Of Luminascent Bacteria In Tcbs Medium
TCBS medium is a selective medium that allows the selective growth of bacteria belonging to
the genera Vibrio. 100 ml of TCBS agar medium was prepared and poured in petri plates and
20 different strains were streaked and observe the result after 24 hours. Appearance of yellow
color colonies in this medium indicates the bacterial strain as Vibrio spp.
3.6 Isolation Of Genomic DNA From Luminescent Bacteria
Materials Used:
NaCl
1.5 µl
Lysis Buffer
0.75 µl
Protease K
30 µl
Saturated Phenol
1ml
Procedure:
1.5 µl culture was centrifuged and supernatant was discarded. Add 1.5 µl of saturated NaCl
was added to the pellet and mixed well until the pellet is dissolved completely and centrifuge
for 1 min. Discard the supernatant and add 0.75 of distilled water, 0.75 of lysis buffer and 30
µl of protease K, mix well and keep it in the water bath for 30 min .Transfer the supernatant
into another Eppendorf and add saturated phenol .Transfer the aqueous layer to another test
tube and store the DNA at 4 degree Celsius
3.7 Agarose Gel Electrophoresis
Materials used
1% Agarose in 1X TAE buffer
Gel Loading Dye
1X TAE buffer (Electrophoresis buffer)
Filter Paper
Gel casting tray
Electrophoresis tank
30 ml
20 µl
500 ml
Procedure
For a 30 ml 1% agarose gel, add 0.3 g of agarose to 30 ml of 1X TAE buffer. Add 0.5ul of
Etbr.
15
Heat solution in a microwave or boiling water bath until agarose is completely dissolved.
Allow to cool.
Prepare gel casting tray by sealing ends of gel chamber with tape or appropriate casting
system. Place appropriate number of combs in gel tray.
3.8 Effect Of External Factors On The Luminascence Of Luminascent Bacteria
3.8.1
Effect of salinity (Varying conecntrations of NaCL)
SWCA medium was prepared by adding different amounts of NaCl to obtain the final
concentrations of salinity such as 0%, 3%, 6%, 9% and 12%. The medium was poured in
Petriplates and six bacteria were streaked per plate with clear divisions between them.
Likewise all the 20 organisms were tested. The plates were incubated for 24 h and the
intensity of luminescence was assessed by visual scoring. According to the visual scoring
method, the symbol - indicates no luminescence, + dull luminescence, ++ good
luminescence and +++ indicates luxuriant luminescence.
3.8.2 Effect of pH
SWCA medium was prepared with four different pH values such as 5, 7, 9 and 11. The pH of
the medium was adjusted with appropriate acid or base and once pH was adjusted the medium
wass added with respective amount of agar and then sterilized. The medium was poured in
Petriplates and six bacteria were streaked per plate with clear divisions between them.
Likewise all the 20 organisms were tested. The plates were incubated for 24 h and the
intensity of luminescence was assessed by visual scoring as described previously. The same
experiment was done once in broth medium in test tubes without agar.
3.8.3 Effect of glycerol concentration
SWCA medium was prepared by adding different amounts of glycerol to obtain the final
concentrations of salinity such as 0.1%, 0.3%, 0.6% and 0.9%. The medium was poured in
Petriplates and six bacteria were streaked per plate with clear divisions between them.
Likewise all the 20 organisms were tested. The plates were incubated for 24 h and the
intensity of luminescence was assessed by visual scoring. According to the visual scoring
method, the symbol - indicates no luminescence, + dull luminescence, ++ good
luminescence and +++ indicates luxuriant luminescence.
3.8.4 Effect of heavy metals
SWCA medium was prepared as usual and was poured in Petriplates. Each bacterium was
grown previously in SWC broth medium and 24 h old broth culture was used in this
experiment. 100 microliters of each broth culture was swabbed over the smooth surface of
SWCA plates and air dried aseptically in a laminar air flow chamber. Then, wells of size 8
mm diameter were made in the seeded agar plates using sterile cork borer. The heavy metals
such as copper zinc and mercury were prepared at two different concentrations viz., 1 mg/ml
16
and 2 mg/ml. 100 microliters from the all the three heavy metals were taken and poured in
the wells made in bacteria seeded SWCA plates. The plates were incubated for 24 h and the
intensity of luminescence was assessed by visual scoring. Also, the zone of inhibition which
is the indicative of susceptibility or resistance of luminescent bacteria to the particular heavy
metal was recorded.
3.9 Preparation of Autoinducers
Sea Water complex broth was prepared, transfer 10 ml of the medium in each test tube and
inoculate the 20 different culture in each test tubes and leave it for overnight .The culture was
taken and it was transformed into the centrifuge tube and centrifuge for 10 min at 4°C.
Transfer the supernatant in another test tube and an equal volume of Dichloro methane
(DCM) was added and it was shaken well. Once again discard the supernatant and take 5ml of
organic phase. The organic phase, supposed to contain autoinduces of acylate domoserine
lactone (AHL) type, were dried to solidness using slow evaporation method in watch glasses.
Finally, the solvent extracts were re-dissolved in 0.5 ml of DCM.
3.10 Bioassay for autoinducers
SWA complex medium was prepared and poured in petri plates and 3 wells fo 8 mm diameter
were made. In the center well 100 microliter of solvent DCM was added, the top and bottom
wells were loaded with DCM extract of respective bacteria. In a plate two bacteria were
tested. Both these two bacterial autoinducer extracts were loaded in top and bottom wells. In
the side of the wells, both the bacteria were streaked. Shortly, the experiment has carefully
designed to check both the self inductive and cross inductive effects of autoinducers. The
intensity of luminescence was assessed by visual scoring.
3.11 Equipments used in the study
Laminar Air Flow
Mini Centrifuge
Refrigerated Centrifuge
Water Bath
Environmental Shaker
UV – Visible Spectrophotometer
Gel Electrophoresis Apparatus
Glassware
Petri Plates
17
4. Results and Discussion
4.1 Isolation of Marine Luminescent Bacteria
A total of 45 well isolated luminescent colonies were selected (Fig. 3) and subcultured to
purity on SWCA. However, the subsequent observations made for the luminescence have
made to arrive at 20 pure cultures of luminescent bacteria (Figure 4 & Table 1). Majority of
the bacteria were obtained using soft agar overlay method. The serial dilution plating
technique was not an appropriate technique for the isolation of luminescent bacteria from
seawater using SWCA.
The majority of luminescent bacteria inhabit the ocean. Two genera of marine bacteria, Vibrio
and Photobacterium, are among the most abundant luminous bacteria. They can be found in
seawater and in the intestinal tract and on the body surfaces of marine animals. The only
terrestrial luminescent bacterial genus known is Photorhabdus. Members of the Photorhabdus
are mostly insect pathogens that exist in a complex symbiotic relationship with a family of
entomopathogenic nematodes (Engebrecht et al., 1983; Moris et al. 1975). In order to apply
bioluminescence of luminous bacteria to industrial use, isolation of luminous bacteria from
various sources was carried out on the basis of strong light intensity, and 18 strains were
obtained. Eleven of these strains were identified as Photobacterium phosphoreum and seven
as Vibrio fischeri (Makiguchi, et al., 1979).
4.2 Subculture Of Selected Strains
These 20 luminescent bacterial strains were tested thrice for their sustainability to exhibit
luminescence (Fig. 4). These bacteria were stored in sterile natural aged sea water in
eppendorf tubes at refrigerated conditions. It has been found that in growth media without sea
water these organisms don‟t survive. This indicates that these organisms are obligate marine
forms. Regular subcultures were made in SWCA before every experiment.
4.3 Biochemical Tests For The Identification Of The Bacterium
4.3.1 KOH string test: gram’s characterization
All the strains were tested negative for KOH string test. So all of them were belonging to the
group of gram negative bacteria (Table 1).
4.3.2
Simple staining for cell morphology
Microscopic observations have confirmedly suggested that all the 20 bacteria were strainght
and short rods. Interestingly, none of them were vibrio shaped (Table 1).
18
4.3.3 Test for motility
All the 20 luminescent bacteria were found to be actively motile (Table 1).
4.3.4 Catalase test
All the 20 luminescent bacteria were found to be actively motile (Fig. 6 & Table 1).
4.3.5 Citrate utilization test
Only five bacteria were found to show negative results for citrate utilization test while
remaining 15 luminescent bacteria have exhibited positive results (Table 1).
4.3.6 Indole test
All the strains were tested positive for indole test (Table 1).
4.3.7 The methyl red and voges-proskauer tests
All the tested 20 luminescent bacteria were found to exhibit negative results for MR and VP
tests (Table 1).
4.3.8 Growth of luminascent bacteria in TCBS medium
None of the tested 20 luminescent bacteria have either grown or produced yellow color
colonies in TCBS agar which is very selective for Vibrio spp. So, it has been confirmed that
none of the 20 strains were belonging to the genera Vibryo (Table 1).
4.3.9 Tentative Identification Of The Strains
With reference to the keys provided in the Bergys Manual of Determinative Bacteriology and
the results of biochemical tests that are summarized in table1, all the 20 strains were
tentatively identified as Photobacterium spp. The strains have exhibited different results for
citrate utilization tests. However, with the tests that have been done in the present study, it
could be concluded only that the strains may belong to Photobacterium spp (Table 1). Further
biochemicals and molecular taxonomical studies are required to confirm the identity of these
organisms.
Marine luminous bacteria comprise gram-negative motile rods, the single, most unique trait of
which is the emission of light. Beijerinck in 1889 recognized the unique nature of
bioluminescence and proposed that all light-emitting bacteria be placed into a single genus,
Photobacterium. Taxonomic studies have since revealed new luminous bacterial species
possessing a large number of phenotypic characters common to members of the
Enterobacteriaceae and Vibrionaceae. As described in Bergey's Manual of Determinative
Bacteriology (Buchanan et al;, 1974) and by Hendrie et al. (1970), there are three genera and
five species of luminous bacteria, Vibrio cholerae biotype albensis, Vibrio fischeri,
Lucibacterium harveyi, Photobacterium phosphoreum, and Photobacterium mandapamensis.
19
4.4 Isolation Of Genomic Dna From Luminascent Bacteria
Genomic DNA from all the 20 luminescent bacteria were isolated in pure form and were run
on agarose gels (Fig. 7). All the strains have equal mobility rate in a 0.7% agarose gel is again
the indicative of the uniform identity of these organisms. They all may belong to same species
of Photobacterium genera.
4.5 Effect Of External Factors On The Luminascence Of Luminascent Bacteria
4.5.1 Effect of salinity
It has been found that up to 6% of NaCl concentration the intense of luminescence was good
and thereafter it declined. Further, in some strains it was completely ceased beyond 9% of
salinity. Interestingly, the strains AMET 1901, AMET1905, AMET1908, AMET1915,
AMET1918 and AMET1920 exhibited immediate luminescence at 3% of NaCl within 5 hours
after incubation (Fig. 8 & Table 2).
Kenneth et al. (1996) have made a study which evaluated the optimal range of sodium
chloride (salt) in a soil and water mixture using Vibrio harveyi, a bioluminescent marine
bacterium, for terrestrial toxicity testing. Their results suggest that the salt range for this
toxicity test with soil is between 7 and 11 percent with the greatest bioluminescence at 9 and
10%. The influence of salt was determined by the amount of bioluminescence in the reaction
mixture. The effect of temperature and salinity on numbers of luminescent bacteria present in
waters of the Mystic (Conn.) River estuary was evaluated by . Counts decreased with
decreasing salinity; none were detected at freshwater stations. A population maximum of 35
per ml was noted at the highest salinity station Pyrocystis lunula is a unicellular, marine,
photoautotrophic, bioluminescent dinoflagellate. Experiments determined if acute changes in
salinity had an effect on the organisms‟ ability to re-establish bioluminescence, or on the
bioassay's potential to detect sodium dodecyl sulfate (SDS) and copper toxicity. Lowering the
salinity from 33 to 27‰ or less resulted in a substantial decrease in re-establishment of
bioluminescence, while increasing the salinity to 43 or 48 ‰ resulted in a small decline.
Salinity had little influence on the bioassay's quantification of Cu toxicity, while the data
showed a weak negative relationship between SDS toxicity and salinity (Jaquelyn et al.
2003). Salinity effect on luminescent bacteria has influenced the pathogenecity of the
organisms. Aquaculture luminescent pathogens such as Vibrio harveyi and Photobacterium
phosphoreum when exposed to low salinities (10,15 ppt) for 12 h before use in immersion
challenge experiments with Penaeus monodon larvae resulted in significantly enhanced
mortalities (P < 0.05). This may account in part for the seasonality of luminous bacterial
disease outbreaks (Prayitno and Latchford, 1995).
4.5.2 Effect of pH
Luminescence was not greatly affected by pH in liquid medium however; the same was
affected in solid medium. pH 7 and 9 were found optimum for the favorable sustenance of
luminescence by luminescent bacteria. Interestingly, all the isolates have exhibited
considerable luminescence in broth with pH 11. Strains AMET1903, AMET1904 and
AMET1913 have exhibited remarkable luminescence in all these range of pH (Fig. 9 & Table
3).
20
Pyrocystis lunula is a unicellular, marine, photoautotrophic, bioluminescent dinoflagellate.
This organism is used in the Lumitox®bioassay with inhibition of bioluminescence reestablishment as the endpoint. Experiments determined if acute changes in pH had an effect
on the organisms‟ ability to re-establish bioluminescence, or on the bioassay's potential to
detect sodium dodecyl sulfate (SDS) and copper toxicity. The re-establishment of
bioluminescence itself was not very sensitive to changes in pH within the pH 6–10 range,
though reducing pH from 8 to levels below 6 decreased this capacity (Jaquelyn et al. 2003).
pH effect on luminescent bacteria has influenced the pathogenecity ofaAquaculture
luminescent pathogens such as Vibrio harveyi and Photobacterium phosphoreum Exposure of
luminous bacteria to acid pH (5.5) significantly reduced their pathogenicity toward penaeid
prawn larvae (P < 0.05). These results imply that environmental factors may play a key role in
disease outbreaks (Prayitno and Latchford, 1995).
4.5.3 Effect of glycerol concentration
The intensity of luminescence has increased with increasing concentrations of glycerol
ranging from 0.3 to 1.2%. However, 0.3% which was used in the composition of SWCA was
not enough to induce pronounced luminescence in all the strains. At increased concentrations
over 0.3%, all the strains have exhibited luxuriant luminescence (Fig. 10, Table 4).
4.5.4 Effect of heavy metals
All the 20 luminescent bacteria were characterized for their tolerance to heavy metals and
antibiotics. Copper and zinc at 1 mg/ml concentration have inhibited the growth and
luminescence of the all strains. Surprisingly, mercury at the same concentration has inhibited
only two strains (AMET1913 and AMET1920). However, at 2 mg/ml concentration mercury
has inhibited the growth and luminescence of all the 20 strains (Fig. 11 & Tables 5, 6).
Selected six luminescent bacterial strains were also characterized for their antibiotic
susceptibility against six different antibiotics. AMET 1901 was found resistant to Amikacin at
30 microgram concentration. AMET 1905 was found to be highly resistant to Amikacin at 30
microgram, nalidixic acid at 30 microgram and cifroflaxacin at 5 microgram concentrations.
Rest of the strains were susceptible to all the tested six antibiotics at varied degrees (Fig. 12 &
Table 7)
Agricultural activities and human industrialization are mainly responsible for the release of
heavy metals into the environment, especially the air and the water. The first step towards the
effective management of water resources is the assessment of pollution levels. Biosensors for
the detection of pollutants in the environment can complement analytical methods by
distinguishing bioavailable from inert, unavailable forms of contaminants. A bioassay system
for detecting heavy metals in water using bioluminescent bacteria, Vibrio harveyi and Vibrio
fischeri has been developed, which offers the advantages of simplicity and rapidity for
screening heavy metals in water sources. Bioluminescence was found to be species specific
and strain specific. Mercury, zinc and copper showed definite microbial toxicity and
inhibition of bioluminescence. The inhibition range for each strain of a species was
standardized and its reproducibility verified. The utility of the biosensors to detect heavy
metals in tap water was demonstrated with samples supplemented with Hg (II) (Seema and
Nair et al., 2005).
21
4.5.5 Bioassay for Autoinducers
Since, the bioluminescence is regulated by quorum sensing, the effect of culture filtrate
extracted with dichloromethane was also tested for its effect on luminescence. These DCM
extracts haven‟t influenced the luminescence much. Moreover, the evaporation of DCM itself
has inhibited the luminescence production considerably.
Quorum sensing is a mechanism of intercellular communication active in many species of
bacteria. It is used by the bacteria to measure the density of their own population within their
environment and to regulate their gene expression and behavior accordingly. For instance, it
is used by many pathogenic species such that when they first invade the host, and are at low
density, they behave in ways that allow them to evade the immune system. If they achieve
high density, and are apparently overwhelming the body‟s defenses, then they change their
behavior and progress to a full-blown disease state. In Vibrio fischeri, quorum sensing
controls bioluminescence, the ability of the bacteria to produce light, an exciting visual
phenomenon for the student lab. The mechanism of quorum sensing involves an autoinducer
synthase, LuxI in V. fischeri, which makes the small autoinducer molecule. The autoinducer
builds up in the medium and at high concentrations will bind to a transcription regulator,
LuxR in V. fischeri, which will then alter the gene expression. We can work with this system
in E. coli, which is easier to handle and more predictable in the lab. We use four E. coli
strains that carry plasmids, extrachromosomal DNA elements, that carry various parts of the
V. fischeri lux genes. None of these strains can luminesce because they are lacking one or
more parts of the lux regulatory system. Two of the strains have the entire system except for
one gene, either luxI or luxR. The other two strains contain only luxI or only luxR (Popham
and Stevens. 2006).
The phenomenon of quorum sensing is a common regulatory mechanism used by a number of
bacteria. During the process of quorum sensing, a bacterial species takes a population census
and thereby induces specific cellular functions only at a high cell density. An intercellular
signaling molecule, commonly termed the autoinducer, is produced and subsequently sensed
by the bacterial cells. Autoinducers can be thought of as pheromones: chemicals produced by
an individual that can be sensed, and interpreted as a specific piece of information, by other
individuals within a population. The quorum sensing response was observed in the
luminescent marine bacterium Vibrio fischeri in the early 1970s and now serves as a model
system for understanding quorum sensing in Gram negative proteobacteria. It has been
determined that two genes are essential for this type of regulatory scheme: luxI, which
encodes an autoinducer synthase called LuxI; and luxR, which encodes an autoinducerdependent activator of the luminescence genes called LuxR. The autoinducer molecule
produced by LuxI is an acylated homoserine lactone (3-oxo-hexanoyl-homoserine lactone). V.
fischeri cells are permeable to the autoinducer, therefore the compound accumulates within
the cells and in the surrounding environment at equal concentrations. When the autoinducer
reaches a critical threshold concentration, LuxR-autoinducer complexes begin to form and the
genes responsible for cellular luminescence (the lux operon) are activated. Quorum sensing
thus constitutes an environmental sensing mechanism that allows the bacteria to respond to
changes in their population density (Popham and Stevens, 2006)
22
5. Summery
Luminescent Bacteria were isolated from various region of the coast. All 20 strains were
characterizes for salinity, pH, and glycerol concentration. Heavy metal resistance zones were
found only for copper, zinc, and mercury. All other heavy metals tested showed resistance.
AMET1902, AMET1903, AMET1910, and AMET1918 showed maximum zone size of
2.2cm. AMET1901 to AMET1906 showed a maximum zone inhibition of 1.2cm and 2.5cm
against Amikacin(AK) and Ciprofloxacin respectively. In general all the tested isolates were
sensitive to all six antibiotics being tested. Majority of the isolates were rod shape, catalase
positive, gram negative, and motile.
6. Conclusion
Looking into the depth of microbial diversity, there is always a chance of finding
microorganisms producing novel enzymes with better properties and suitable for commercial
exploitation. The multitude of physic-chemically diverse habitats has challenged nature to
develop equally numerous molecular adaptations in the microbial world. Microbial diversity
is a major resource for biotechnological products and processes. Thus the strains isolated from
the different region has good beneficial potential such as heavy metal tolerance and antibiotic
sensitivity
23
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28
Appendix 1
Source
Gram’s
Nature
Citrate
Utilization
Indole
+
+
+
+
+
+
+
+
Catalase
+
-
+
+
+
+
+
+
Motility
Positive
Positive
Positive
Positive
Positive
Positive
Positive
Positive
Cell
Shape
Motile
Motile
Motile
Motile
Motile
Motile
Motile
Motile
MR
VP
Slant: Alkaline
TSI
Gas: -
Gas:
++
+++
Slant: Alkaline
-H2S:
Butt: Acidic
Slant: Alkaline
Gas: -H2S: +++
Butt: Acidic
Slant: Alkaline
Gas: -H2S: ++
Butt: Acidic
Slant: Alkaline
Gas: -H2S: +++
Butt: Acidic
Slant: Alkaline
Gas: -H2S: +
Butt: Acidic
Slant: Alkaline
Gas: -H2S: ++
Butt: Acidic
Slant: Alkaline
Gas: -H2S: +++
Butt: Acidic
Slant: Alkaline
H2S:
Butt: Acidic
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
Butt: Acidic
Gas: -H2S: +++
Nil
Nil
Nil
Nil
Nil
Nil
Nil
Growth
on TCBS
agar
Nil
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp.
Tentative
identification
Nil
Table 1. Biochemcial tests for the tentative identification of luminescent bacteria isolated from Bay of Bengal Sea Water
Strain code
Gram
Negative
Straight
short
rods
+
Gram
Negative
Straight
short
rods
+
Gram
Negative
Straight
short
rods
Positive
Gram
Negative
Straight
short
rods
Motile
Gram
Negative
Straight
short
rods
S1
Gram
Negative
Straight
short
rods
Straight
short
rods
Gram
Negative
AMET1901
S1
S1
S1
S2
S2
S2
S2
S2
Straight
short
rods
Gram
Negative
AMET1902
AMET1903
AMET1904
AMET1905
AMET1906
AMET1907
AMET1908
AMET1909
Gram
Negative
Straight
short
rods
AMET1910
AMET1912
AMET1913
AMET1914
AMET1915
AMET1916
AMET1917
AMET1918
AMET1919
AMET1920
S3
S3
S3
S3
S4
S4
S4
S4
S4
S4
Gram
Negative
Gram
Negative
Straight
short
rods
Gram
Negative
Straight
short
rods
Straight
short
rods
Gram
Negative
Straight
short
rods
Gram
Negative
Gram
Negative
Straight
short
rods
Straight
short
rods
Gram
Negative
Straight
short
rods
Gram
Negative
Gram
Negative
Straight
short
rods
Straight
short
rods
Gram
Negative
Straight
short
rods
Motile
Motile
Motile
Motile
Motile
Motile
Motile
Motile
Motile
Motile
Positive
Positive
Positive
Positive
Positive
Positive
Positive
Positive
Positive
Positive
+
+
+
-
-
-
+
+
+
+
+
+
+
+
+
+
+
+
+
+
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
Slant: Alkaline
Butt: Acidic
Gas: -H2S: +++
Slant: Alkaline
Butt: Acidic
Gas: -H2S: +++
Slant: Alkaline
Butt: Acidic
Gas: -H2S: ++
Slant: Alkaline
+++
Butt: Acidic
Gas:
-H2S:
Slant: Alkaline
Butt: Acidic
Gas: -H2S: +
Slant: Alkaline
Butt: Acidic
Gas: -H2S: ++
Slant: Alkaline
Butt: Acidic
Gas: -H2S: +++
Slant: Alkaline
Butt: Acidic
Gas: -H2S: +++
Slant: Alkaline
Butt: Acidic
Gas: -H2S: +++
Slant: Alkaline
Butt: Acidic
Gas: -H2S: +++
Nil
Nil
Nil
Nil
Nil
Nil
Nil
Nil
Nil
Nil
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
Photobacterium sp
3(8)
Table 2. Effect of different concentrations of NaCl on the luminescence of luminescent
bacteria
Luminescence in different concentration of NaCl (%)
Isolate code
0
3
6
9
12
AMET1901
++
++
++
-
-
AMET1902
++
++
++
-
-
AMET1903
++
++
+++
-
-
AMET1904
++
++
+++
-
-
AMET1905
++
++
++
-
-
AMET1906
++
+++
++
-
-
AMET1907
++
+
++
-
-
AMET1908
++
+
++
-
-
AMET1909
++
+++
++
-
-
AMET1910
++
+
+
-
-
AMET1911
++
+
++
-
-
AMET1912
++
+++
++
-
-
AMET1913
++
+++
++
-
-
AMET1914
++
+++
++
-
-
AMET1915
++
++
++
-
-
AMET1916
++
++
++
-
-
AMET1917
++
++
++
-
-
AMET1918
++
++
++
-
-
AMET1919
++
++
++
-
-
AMET1920
++
+++
+++
-
-
-
No luminascenmce;
+
Dull luminescence
++
Good luminescence;
+++
Luxuriant luminescence
4(8)
Table 3.Effect of different pH on the luminescence of luminescent bacteria
Isolate code
Luminescence of bacteria in different pH
5
7
9
11
AMET1901
+
+
+
+
AMET1902
+
++
+
+
AMET1903
+
+
++
+
AMET1904
+
+
++
+
AMET1905
+
+
+
+
AMET1906
+
+
+
+
AMET1907
+
+
+
+
AMET1908
+
+
+
+
AMET1909
+
+
+
+
AMET1910
+
+
+
+
AMET1911
+
+
+
+
AMET1912
+
+
+
+
AMET1913
+
+
++
+
AMET1914
+
+
+
+
AMET1915
+
+
+
+
AMET1916
+
+
+
+
AMET1917
+
+
+
+
AMET1918
+
+
+
+
AMET1919
+
+
+
+
AMET1920
+
+
++
+
-
No luminascenmce;
+
Dull luminescence
++
Good luminescence;
+++
Luxuriant luminescence
5(8)
Table 4.Effect of heavy metals (1 mg/ml) on the luminescence of luminescent bacteria
Isolate Number
Heavy metal (1 mg/ml) tolerance spectrum
Copper
Zinc
Mercury
AMET1901
S(1.8 )
S(1 )
R
AMET1902
S (2.2)
S(1.6)
R
AMET1903
S (2.2)
S(1.5)
R
AMET1904
S(2)
R
R
AMET1905
S(2)
S(1.9)
R
AMET1906
S(2)
S(1.9)
R
AMET1907
S(1.8)
S(1.5)
R
AMET1908
S(1.9)
S(1.2)
R
AMET1909
S (2)
S(1.1)
R
AMET1910
S (2.2)
S(1.3)
R
AMET1911
S (1.9)
S(0.6)
R
AMET1912
S (2)
S(0.6)
R
AMET1913
S (2.2)
S(1.2)
S(1 )
AMET1914
S (1.9)
S(1.2)
R
AMET1915
S (0.6)
S(0.7)
R
AMET1916
S (1.9)
S(2)
R
AMET1917
S (2)
S(1.7)
R
AMET1918
S (2.2)
R
R
AMET1919
S (2.1)
S(1.1)
R
AMET1920
S (0.8)
S(2)
S(0.5)
R- resistant ; S – Susceptible.
Values in parentheses are zone of inhibition in cm
6(8)
Table 5.Effect of heavy metals (2 mg/ml) on the luminescence of luminescent bacteria
Isolate Number
(Mercury
Copper
AMET1901
S(1.5)
S(1 )
AMET1902
S(1.2)
S(1.1)
AMET1903
S(1.4)
R
AMET1904
S(1.3)
S(1.6)
AMET1905
S(1.1)
S(1.3)
AMET1906
S(1.5)
S(1.1)
AMET1907
S(1.3)
S(1.3)
AMET1908
S(1.4)
S(1.6)
AMET1909
S(1.2)
S(2)
AMET1910
S(1.1)
S(1.7)
AMET1911
S(1.2)
S(1.6)
AMET1912
S(1.2)
S(1)
AMET1913
S(1.2)
R
AMET1914
S(1.1)
S(2)
AMET1915
S(1.2)
S(1.1)
AMET1916
S(1.4)
R
AMET1917
S(1.3)
R
AMET1918
S(1.1)
R
AMET1919
S(1.5)
S(1.6)
AMET1920
S(1.2)
R
R- resistant ; S – Susceptible.
Values in parentheses are zone of inhibition in cm
7(8)
Table 6.Effect of different concentrations of glycerol on the luminescence of
luminescent bacteria
Isolate code
Luminescence in different concentrations of glycerol (%)
0.1%
0.3%
0.6%
0.9%
AMET1901
+
+
+++
+++
AMET1902
+
+
+++
+++
AMET1903
+
+
+++
+++
AMET1904
+
+
+++
+++
AMET1905
+
+
+++
+++
AMET1906
+
+
+++
+++
AMET1907
+
+
+++
+++
AMET1908
+
+
+++
+++
AMET1909
+
+
+++
+++
AMET1910
+
+
+++
+++
AMET1911
+
+
+++
+++
AMET1912
+
+
+++
+++
AMET1913
+
+
+++
+++
AMET1914
+
+
+++
+++
AMET1915
+
+
+++
+++
AMET1916
+
+
+++
+++
AMET1917
+
+
+++
+++
AMET1918
+
+
+++
+++
AMET1919
+
+
+++
+++
AMET1920
+
+
+++
+++
++
No luminascenmce;
Good luminescence;
+
+++
Dull luminescence
Luxuriant luminescence
8(8)
Table 7.Antibiotic susceptibility/resistance of Luminescent Bacterial strains
Isolate
code
Antibiotic sensitivity spectrum
Amikacin
Nitrofurantoin Natillin
Nalidixic
30 mcg
300 mcg
acid
30 mcg
Ceftazidime Ciprofloxacin
30 30 mcg
5 mcg
mcg
AMET
R
S (1.6)
S (1.5)
S( 1.7 )
S (1 )
S( 2.5 )
1901
AMET
S( 1.4 )
S (1.6)
S (1.2)
S( 1.4 )
S (1.5 )
S( 0.8 )
S (1.2 )
S (1)
S (1.4 ) S (1.2 )
S( 1.7 )
S (2)
S (1.3 )
S (1.2 )
S( 1.4 ) S (1 )
S (1.2 )
S (1.8)
R
S (1.2)
S (1.2 ) R
S (0.8 )
R
S (1.2 )
S (1.6)
S(1.4 )
S (1 )
S (1.7 )
1902
AMET
1903
AMET
1904
AMET
1905
AMET
S (1)
1906
R- resistant ; S – Susceptible.
Values in parentheses are zone of inhibition in cm
Appendix 2
Figure 1. View of sampling site, Kanathur, Chennai
Figure 2. Map showing the sampling sites in the study
area, East Coast of India, Bay of Bengal Sea
1(8)
2(8)
Figure 3. View of luminascnet bacterial colonies in
darkness on the Sea Water Complex Agar medium
Figure 4. Isolated pure cultures of luminescent
bacteria in normal light on the Sea Water Complex
Agar medium
3(8)
Figure 5.. Isolated pure cultures of luminescent bacteria in
dark on the Sea Water Complex Agar medium
Figure 6. Biochemical tests for identification: catalase test
+ indicates positive reaction; - indicates negative reaction
4(8)
Figure 7. Isolation of genomic DNA from
luminescent bacteria
0 % NaCl
3 % NaCl
6 % NaCl
Figure 8. Effect of different concentrations of NaCl
on the luminescence of luminescent bacteria
5(8)
pH 5
pH 7
pH 9
pH 11
Figure 9. Effect of different pH on the luminescence
of luminescent bacteria
6(8)
0.1% glyceol
0.6% glyceol
0.9% glyceol
0.9% glycerol
0.3% glyceol
Figure 10. Effect of different concentrations of
glycerol on the luminescence of luminescent bacteria
7(8)
Figure 11. Effect of heavy metals on the luminescence
of luminescent bacteria
Top: 0.1 mg/mL of Heavy metal;
Bottom : o.2 % of heavy metal
8(8)
1
4
2
5
3
6
Figure 12. Antibiotic susceptibility/resistance of six
Luminescent Bacterial strains
1. AMET 1901; 2. AMET 1902; 3. AMET 1903;
4. AMET 1904; 5. AMET 1905; 6. AMET 1906