Isolation of Luminescent Bacteria from Bay of Bengal and Their Molecular Characterization

Home , Bioluminescent bacteria, Luminescent bacteria

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: Department of Biotechnology AMET University (Under Sec. 3 of UGC Act 196) Chennai-603112, India Chennai-603112, India Company/Organisation or street> Client: Department of Biotechnology, AMET University, Chennai-603112, India Date: Company/Organisation2010-09-08 or street> Keywords: Sea water, luminescent bacteria, Bay of Bengal

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”

iii 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

iv 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

v 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 Table 1 – Table 7 Appendix 2 Figure 1 – Figure 12

vi 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

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 long- chain 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 non- bioluminescent 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, receptor- based 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 5 g Yeast extract 3 g Glycerol 3 ml Agar 15 gram 50% Sea water

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.

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)

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 30 ml Gel Loading Dye 20 µl 1X TAE buffer (Electrophoresis buffer) 500 ml Filter Paper Gel casting tray Electrophoresis tank

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 re- establishment 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 autoinducer- dependent 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 References

Aruldoss, J.A. and Viraraghavan, T. B. 1998. Environ. Contam.Toxicol., vol. 60, no. 3, pp. 456–463.

Baltimore: Williams and Wilkins, 1984. Bergey‟s Manual Systematic Bacteriology, vol. 1.

Birger, M.O., Spravochnik po mikrobiologicheskim ivirusologicheskim metodam issledovaniya (Handbook of Methods in Microbiology and Virology), Moscow: Meditsina, 1973, p. 73.

Baumann, P., Baumann, L. 1983. Genus Photobacterium. In: Krieg, N. R. (ed.), Bergey's Manual of Systematic Bacteriology, 9th ed. Baltimore: Williams and Wilkins Co., in press.

Baumann, P., Furniss, A. L., Lee, J. V. 1983. Genus Vibrio.

Baumann, P., Gauthier, M., Baumann, L. 1983. Genus Alteromonas. in: Krieg, N. R. (ed.), Bergey's Manual of Systematic Bacteriology, 9th ed. Baltimore: Williams and Wilkins Co., in press

Baumann, P., Baumann, L. 1977. Biology of the marine enterobacteria: Genera Beneckea and Pholobacterium. Annual Review of Microbiology 31:39-61.

Baumann, P., Baumann, L, 1980. The marine Gram-negative eubacteria, in: Start, M. P., Stolp, H., Triper, H G.. Balows, A., Schlegel, H. G. (eds.), The prokaryotes: A handbook on habitats, isolation, and identification of bacteria. New York, Heidelberg, Berlin: Springer- Verlag. In press.

Bang, S. S., Baumann, P., Nealson, K. H. 1978. Phenotypic characterization of Photobacterium Iogei {sp. nov.), a species related to P. fischeri. Current Microbiology 1:285- 288.

Baumann, L., Baumann, P., Mandel, M., Allen, R. D. 1972, Taxonomy of marine bacteria: the genus Beneckea. Journal of Bacteriology 107: 268-294.

Beebe, W. & M. Vander Pyl, 1944. Eastern Pacific expeditions of the New York Zoological Society. XxX111. Pacific Myctophidae (fishes). Zoologica, Vol. 29, pp. 59-95.

Baguet, F., 1975. Excitation and control of isolated photophores of luminous fishes. Prog. Neurobiol., Vol. 5,pp. 97-125.

Baguet, F., B. Christophe & G. Marechal, 1980. Luminescence of Argyropelecus photophores electrically stimulated. Camp. Biochem. Physiol. Ser. A, Vol. 67, pp. 375-381.

Baguet, F. Kc G. Marechal, 1976. Bioluminescence Nicol, J.A.C., 1958. Observations on luminescence in pelagic animals. J. Mar. Bioi. Assoc. U.K., Vol. 37,pp. 705-752. Blaise, C. (1991) Microbiotest in Aquatic Ecotoxicology: Characteristics, Utility, and Prospects. Environ. Toxicol. Wat. Qual. 6, 145-155.

24 Boylan M., J. Pelletier, E.A. Meighen. 1989. Fused bacterial luciferase subunits catalyze light emission in eukaryotes and prokaryotes, J. Biol. Chem. 264, 19151918. Cormier, M. J., Lee, J., Wampler, J. 1975. Biolumiuescence: recent advances. Ann. Rev. Biochem. 44, 255-272

Cormier, M. J., Totter, J. R. 1968. Bioluminescence: enzymic aspects. Photophysiology 4, 315- 353

Campbell, A.K. 198. Living Light: Biochemistry, Function and Biomedical ApplicationsEssays in Biochem. Vol 24, 41-81

Campbell, D.L.; Lawton, L.A.; Beattie, K.A.; Codd, G.A.1994. Comparative Assessmentof the Specificity of the Brine Shrimp and Microtox Assays to Heptatoxic (Microcyctin- LRContaining) Cyanobacteria. Environ. Toxicol. Water Qual.: An Intern. J. 9, 71-77.

Delistraty, D. 1984. Bioluminescent Toxicity Assay of Synfuel By-Product Waters. Bull.Environ. Contam. Toxicol. 32, 613-620.

Gitel‟- zon, I.I., Ed., Novosibirsk Nauka, 1984. Svetyashchiesya bakterii (Luminous Bacteria),

Gordon, S., Parish, T., Roberts, I. S , and Andrew, P. W. 1994. The application of luciferase as a reporter of environmental regulation of gene expression in mycobacteria. Lett Appl Mlcroblol. 19, 336-340.

Grayeski, M. L. 1987. Chemiluminescence Analysis. Anal. Chem. 59, 1243A-1256A.

Hastings, J. W. : Bioluminescence. 1968. Ann. Rev. Biochem. 37, 597- 630 (1968)

Hastings, J. W., Riley, W. H., Massa, J. 1965. The purification, properties, and chemiluminescent quantum yield of bacterial luciferase. J. biol. Chem. 240, 1473- 1481

Hastings, J. W., Nealson, K. H. 1977. Bacterial bioluminescence. Annual Review of Microbiology 31:549-595.

Hastings, J. W., Nealson, K, H. 1980. The symbiotic luminous bacteria. In: Start-, M. P., Stolp, H., Truper, H. G., Balows, A., Schlegel, H. G. (eds.), The prokaryotes: A handbook onhabitats, isolation, and identification of bacteria. New York, Heidelberg, Berlin: Springer- Verlag. In press.

Hastings, J. W 1978. Bacterial btolummescence an overview. Methods Enzymol 57,125-135

Hastings, J. W., Nealson, K. H. 1981. The symbiotic luminous bacteria, p. 1332-1346. In M.P. Starr, H. Stolp, H. G. Truper, A. Balows, and H.G. Schiegel (eds.), Prokaryotes, Vol. II. Springer-Verlag, New York.

Herring, P.J., 1977. Oral light organs in Sremoplyx, with some observations on the bioluminescence of hatchetfishes. In, A voyage of discoverye,d ited by M. Angel, Pergamon Press, Oxford, pp. 553-567.

Herring, P. J., 1982. Aspects ofbioluminescence offishes. Ocean. Mar. &al. Annu. Rev., Vol. 20, pp. 415-470. 25 Herring, P. J., 1983. The spectral characteristics of luminous marine organisms. Prac. R. Sac. London Ser.B, Vol. 220, pp. 183-217.

Herring, P. J., 1987. Systematic distribution of bioluminescence in living organisms. J. Bi&m. Chemj~um.,Vol. 1, pp. 147-163.

Herring, P. J. & J. G. Morin, 1978. Bioluminescence in fishes. In, Bioluminescence in action, edited by P. J. Herring, Academic Press, London, pp. 273-329.

Hollis R.P., C. Lagido, J. Pettitt, A.J. Porter, K. Killham, G.I. Paton, L.A. Glover. 2001. Toxicity of the bacterial luciferase substrate, n-decyl aldehyde, to Saccharomyces cerevisiae and Caenorhabditiselegans, FEBS Lett. 506 , 140–142.

Holt, J.G., Ed., Baltimore: Williams and Wilkins, 1977. The Shorter Bergey’s Manual of Determinative Bacteriology, Mir, 1997, vol. 1.

Krieg, N. R. (ed.), Bergey's Manual of Systematic Bacteriology, 9th ed. Baltimore: Williams and Wilkins Co., in press.

Kempner, E. S., Hanson, F. E.1968. Aspects of light production by Photobaeteriumfischeri. J. Bacteriol. 95, 975-979

Kirby, K. S. 1964. Isolation and fractionation of nucleic acids. Progr. Nucl. AcidsRes. 3,1.

Lin, P., Saier, M. H., Nealson, K. H. Regulation of the synthesis of the bioluminescent system in Beneckea harveyi. Fed. Proc. 35,

Meighen, E A. 1991. Molecular biology of bacterial btolummescence MtcroblolRev. 55, 123- 142.

Marmur, J. 1961. A procedure for the isolation of deoxyribonucleic acid from microorganisms J Mol. Biol. 3,208-218

Meighen, E.A. 1988. Enzymes and Genes from the lux Operons of Bioluminescent BacteriaAnn. Rev. Microbial. 42, 151-176.

Meighen, E.A. 1991. Molecular Biology of Bacterial Bioluminescence Microb. Rev., March1991, 123-142.

McFeters, G.A.; Bond, P.J.; Olson, S.B.; Tchan, Y.T. 1983. A Comparison of MicrobialBioassays for the Detection of Aquatic Toxicants. Water Res. 12, 1757-1762.

Nealson, K. H., Eberhard, A., Hastings, J. W. 1972. Catabolite repression of bacterial bioluminescence: Functional implications. Proc. Nat. Acad. Sci. USA 69, 1073-1076

Nealson, K. H., Hastings, J. W. 1976. Low oxygen is optimal for luciferase synthesis in some bacteria. Arch. Microbiol. 112, 9-16 (1977)361

Nealson, K. H., Hastings, J. W. 1979. Bacterial bioluminescence; its control and ecological significance. Microbiological Reviews 43:496-518.

26 Nicol, J. A. C., 1960. Spectral composition of the light ofthe lantern fish, Myctophumpunctatum. J. Mar. Bial.Assoc. U.K., Vol. 39, pp. 27-32.

Nicol, J. A. C., 1967. The luminescence of fishes. Symp. Zool. Sot. London, Vol. 19, pp. 27- 55.

Nicol, J. A.C., 1969. Bioluminescence. In, Fish Physiology, edited by W. S. Hoar, Academic Press, New York, pp. 355-400.

Nicol, J.A. C., 1978. Bioluminescence and vision. In, Bioluminescence in action, edited by P. J. Herring, Academic Press, London, pp. 367-398.

Olsson O., A. Escher, G. Sandberg, J. Schell, C. Koncz, A.A. Szalay, Engineering of monomeric bacterial luciferases by fusion of luxA and luxB genes in Vibrio harveyi, Gene 81 1989. 335–347.

Ow, D.W.; Wood, K.V.; DeLuca, M.; De Wet, J.R.; Helinski, D.R.;„Howell, S.H. 1990.Transient and Stable Expression of the Firefly Luciferase Gene in Plant Cells and TransgenicPlants. Science 234, 856-859.

Reichelt, J.L., Nealson, K., Hastings, J.W. 1977. The specificity of symbiosis: pony fish and luminescent bacteria. Archives of Microbiology 112:157-161.

Ruby, E. G., Morin, J. G. 1978. Specificity of symbiosis between deep-sea fishes and psychrotrophic luminous bacteria. Deep-Sea Research 25:161-167.

Reichelt, J. L., Baumann, P. 1973. Taxonomy of the marine luminous bacteria. Archiv for Mikrobiologie 94:283 330.

Reichelt, J. L, Baumann, P. 1975. Photobacteriurn mandapamensis Hendrie etal., a later subjective synonym of Photobacterium leiognathi Boisverl etal. International Journal of Systematic Bacteriology 25:208-209.

Reichelt, J. L., Baumann, P., Baumann, L. 1976. Study of genetic relationships among marine species of the genera Beneckea and Photobacterium by means of in vitro DNA/DNA hybridization. Archives of Microbiology 110:101-120

Runv, E. G., AND K. H. NEALSON. 1977. Pyruvate production and excretion by the luminous marinebacteria. Appl Environ. Microbiol 34:164-169.

Runv, E. G., AND K. H. NEALSON. 1978. Seasonal changes in species composition of luminous bacteria in nearshore seawater. Limnol. Oceanogr. 23:530- 533.

Santa Maria, A., Lopez, A., Diaz, M.M., et al., Ecotoxicol. Environ. Safety, 1998, vol. 39, no. 3, pp. 164–167.

Thomulka, K.; McGee, D.; Lange, J. 1993 Detection of Biohazardous Materials in Water by Measuring Bioluminescence Reduction with the Marine Organism Vibrio hmveyi. J. Environ. Sci. Health, A 28(9), 2153-2166.

Tchan, Y.T,; Roseby, J.E. and Funnell, J.R. 1975 A New Rapid Specific Bioassay Method for Photosynthesis Inhibiting Herbicides. Soil Biol. Biochem. 7, 39-44.

27 Virta, M., Lineri, S., Kankaanpaa, P., et al., Appied. Environmental Microbiol., 1998, vol. 64, no. 2, pp. 515–529.

Young, R.E., Kampa, E.M., Maynard, S.D., Mencher, F.M., Roper, C.F.E., 1980. Counterillumination and the upper depth limits of midwater animals. Deep Sea Res. 27A, 671–691. Young, R.E., Roper, C.F.E., 1977. Intensity regulation of bioluminescence during countershading in living midwater animals. Fish. Bull. 75, 239–252.

28 Appendix 1 Strain code Strain 1.BiochemcialTable of the for luminescent tests tentative bacteria identification from Water isolated Sea Bengal of Bay AMET1909 AMET1908 AMET1907 AMET1906 AMET1905 AMET1904 AMET1903 AMET1902 AMET1901

S2 S2 S2 S2 S2 S1 S1 S1 S1 Source

Negative Gram Negative Gram Negative Gram Negative Gram Negative Gram Negative Gram Negative Gram Negative Gram Negative Gram Nature Gram’s

rods short Straight rods short Straight rods short Straight rods short Straight rods short Straight rods short Straight rods short Straight rods short Straight rods short Straight Shape Cell

Motile Motile Motile Motile Motile Motile Motile Motile Motile Motility

Positive Catalase Positive Pos Positive Positive Positive Positive Positive Positive itive

Utilization Citrate + + + + + + + + -

Indole + + + + + + + + +

- MR ------

- VP ------

Butt: Acidic Butt: Alkaline Slant: TSI Gas: Acidic Butt: Alkaline Slant: Gas: Acidic Butt: Alkaline Slant: Gas: Acidic Butt: Alkaline Slant: Gas: Acidic Butt: Sla Gas: Acidic Butt: Alkaline Slant: Gas: But Alkaline Slant: Gas: Acidic Butt: Alkaline Slant: Gas: Acidic Butt: Alkaline Slant: Gas: nt: Alkaline nt: t: Acidic t:

------H H H H H H H H H 2 2 2 2 2 2 2 2 2 S: +++ S: +++ S: +++ S: ++ S: +++ S: + S: ++ S: +++ S: S: ++ S:

Nil Nil Nil Nil Nil Nil Nil Nil Nil agar TCBS on Growth

Photobacterium Photobacterium Photobacterium Photobacterium Photobacterium Photobacterium Photobacterium Photobacterium Photobacterium Photobacterium iden Tentative tification

sp sp sp sp sp sp sp sp sp.

AMET1910 S3 Gram Straight Motile Positive + + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: +++ AMET1912 S3 Gram Straight Motile Positive + + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: +++ AMET1913 S3 Gram Straight Motile Positive + + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: ++ AMET1914 S3 Gram Straight Motile Positive - + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: +++ AMET1915 S4 Gram Straight Motile Positive - + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: + AMET1916 S4 Gram Straight Motile Positive - + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: ++ AMET1917 S4 Gram Straight Motile Positive + + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: +++ AMET1918 S4 Gram Straight Motile Positive + + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: +++ AMET1919 S4 Gram Straight Motile Positive + + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: +++ AMET1920 S4 Gram Straight Motile Positive + + - - Slant: Alkaline Nil Photobacterium sp Negative short Butt: Acidic rods Gas: -H2S: +++

3(8)

Table 2. Effect of different concentrations of NaCl on the luminescence of luminescent bacteria Isolate code Luminescence in different concentration of NaCl (%) 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; + Dull luminescence ++ Good luminescence; +++ Luxuriant luminescence

8(8)

Table 7.Antibiotic susceptibility/resistance of Luminescent Bacterial strains

Isolate Antibiotic sensitivity spectrum code Amikacin Nitrofurantoin Natillin Nalidixic Ceftazidime Ciprofloxacin 30 mcg 300 mcg 30 mcg acid 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 ) 1902

AMET S (1.2 ) S (1) S (1.4 ) S (1.2 ) S( 1.7 ) S (2) 1903 AMET S (1.3 ) S (1.2 ) S( 1.4 ) S (1 ) S (1.2 ) S (1.8) 1904 AMET R S (1.2) S (1.2 ) R S (0.8 ) R 1905 AMET S (1.2 ) S (1.6) S(1.4 ) S (1) S (1 ) S (1.7 ) 1906

R- resistant ; S – Susceptible. Values in parentheses are zone of inhibition in cm

Appendix 2 1(8)

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 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 2 3

6 4 5

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