Characterization of Bacterial isolates from Kashmir cave, Pakistan, and their Potential Applications.
By
Sahib Zada
Department of Microbiology Quaid-i-Azam University Islamabad, Pakistan 2017
Characterization of Bacterial isolates from Kashmir cave, Pakistan, and their Potential Applications.
A thesis
Submitted in the Partial Fulfillment of the
Requirements for the Degree of DOCTOR OF PHILOSOPHY
IN
MICROBIOLOGY
By
Sahib Zada
Department of Microbiology Quaid-i-Azam University Islamabad, Pakistan 2017
DECLARATION
The material contained in this thesis is my original work and I have not presented any part of this thesis/work elsewhere for any other degree.
Sahib Zada
DEDICATED
TO
My Ammi, Abbu and
Late Talha khan
CONTENTS
S. No. Title Page No.
1. List of Abbreviations i
2. List of Tables ii
3. List of Figures iii
4. Acknowledgements vi
5. Summary viii
6. Chapter 1: Introduction 1
7. Chapter 2: Review of Literature 18
8. Chapter 3: Paper 1 60
Paper 2 99
Paper 3 124
Paper 4 147
9. Chapter 5: Mn oxidation by Cavernicoles
a. Abstract 170
b. Introduction 171
c. Methodology 180
d. Results 185
e. Discussion 201
f. References 204
10. Conclusions 206
11. Future Prospects 207
List of Abbreviations
A Adenosine ATP Adenosine-5’-triphosphate BLAST Basic Local Alignment Search Tool BLASTX BLAST search using a translated nucleotide query °C Degree celsius Ca Calcium DNA Deoxyribonucleic acid e.g. Exempli gratia, for example et al. et alii/alia, and others Fe Iron Fig. Figure FTIR Fourier Transform Infra-Red spectroscopy HEPES N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid LBB Leucoberbelin Blue LDPE Low density polyethylene MCO Multi Copper Oxidase MEGAN 5 MEtaGenome ANalyzer Mn Manganese MOB Manganese Oxidizing Bacteria NaCl Sodium Chloride NCBI National Center for Biotechnology Information OD Optical Density PCR Polymerase Chain Reaction pH Power of Hydrogen RDP Ribosomal Database Project rRNA Ribosomal ribonucleic acid SEM Scanning Electron Microscopy SOD Superoxide Dismutase TBE Tris base, boric acid EDTA UV-Vis Ultra Violet Visible VOC Volatile Organic Compounds X-RD X-ray Powder Diffraction Zn Zinc
List of Tables No. Title Page No. 1 List of largest and most studied caves world wide 9 2 List of caves in Pakistan 10 3 Cave microbes isolated from different caves in different country 13 through out the world 4 Different caves are studied for microbial diversity in different 37 countries 5 List of minerals obtained from smast-5 floor and smast-7 wall of 72 Kashmir smast samples 6 Concentration of elements in sample collected from cave floor and 72 outside cave soil (control) 7 Viable Cell Count of bacterial consortium in different media 109 compositions incubated at 37°C. 8 Mn(II) oxidizing Bacteria Isolated from Kashmir cave soil and 186 speleothem.
List of Figure
S.No. Page Title No. 1 Different types of caves and its formation including dissolution and 4 weathering. 2 Solutional cave formation 5 3 Limestone cave formation 5 4 Distribution of major groups of microbial communities in cave 12 environments by 16S rRNA gene sequencing 5 The process of calcium carbonate precipitation by bacteria 30 6 Bacteria serving as nucleation site for CaCO3 precipitation in the 31 sand particle 7 Distribution of major groups of microbial communities in cave 36 environments by 16S rRNA gene sequencing 8 Kashmir cave (smast), Nanseer Buner, Khyber Pakhtunkhwa, 65 Pakistan. White arrows show location of the cave, black arrow 105 shows entrance to the cave. 9 XRD patterns of Kashmir smast (a) from the floor and (b) from the 71 wall along-with the matched peaks of the mineral ICSD (Inorganic Crystal Structure Database) 10 Quantitative analysis of minerals, A. Wall soil sample smast-7, B. 73 Floor soil sample smast-5 11 Infrared spectra of Smast-7 wall 73 12 TGA (Thermogravimetric Analysis) plots of Kashmir smast 74 (sample-5 Floor and sample-7 wall) 13 FE-SEM micrograph & EDS spectra of (a) smast-7 wall and (b) 75 smast-5 floor 14 Nutrient agar plate showing the zones of inhibition against the 76 clinical isolates 15 Phylogenetic tree of all four species with related sequences in NCBI 77-78 16 Effect of time of incubation, pH and temperature on the growth and 79 antimicrobial activity of B. licheniformis KC2-MRL against M. luteus, S. aureus, Klebsiella and E. coli 17 Zone of inhibition of our four antibiotic producing strains (Serratia 80 sp. KC1-MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3- MRL and Stenotrophomonas sp. KC4-MRL) against selected antibiotics in mm. 18 Comparison of FTIR spectra of control (Bacitracin) and the 81 antibacterial compound produced by B. licheniformis KC2-MRL 19 Fourier-transform infrared spectra from control and different media 112 after incubation at 37°C for one month 20 Scanning electron microscopy of low-density polyethylene samples 114 under specified treatment after incubation with bacterial consortia at 37°C for one month. 21 Limestone cave (Kashmir Smast Pirsai Mardan, Pakistan) 126 22 Evolutionary relationships of taxa Lysinibacillus sphaericus KC5- 130 MRL. 23 Effect of size of inoculum on the growth of Lysinibacillus 131 sphaericus KC5-MRL and production of lipase. 24 Lipolytic, amylolytic and proteolytic activity of Lysinibacillus 132 sphaericus KC5-MRL at 30°C 25 Stability of crude extracts of lipase, amylase and protease of 134 Lysinibacillus sphaericus KC5-MRL at different pH 26 Stability of crude extracts of lipase, amylase and protease of 136 Lysinibacillus sphaericus KC5-MRL at different temperature 27 Effects of detergents on the lipolytic, amylolytic and proteolytic 137 activity of isolate Lysinibacillus sphaericus KC5-MRL 28 Effects of metal ions on lipase, amylase and protease activity of 138 isolate Lysinibacillus sphaericus KC5-MRL 29 Effects of organic solvents on lipase, amylase and protease activity 139 of Lysinibacillus sphaericus KC5-MRL 30 Effect of inhibitors on lipase, amylase and protease activity of 139 Lysinibacillus sphaericus KC5-MRL 31 Molecular Phylogenetic analysis by Maximum Likelihood method 154 32 Calcium precipitates induced by bacteria in crystals form A) 155 Paracoccus Limosus, B) Brevundimonas naejangsanensis. 33 Compound microscopy of precipitates produced at 25°C. 155 34 Compound microscopy of precipitates produced at pH 5. 156 35 Compound microscopy of precipitates produced after 20 days of 156 incubation. 36 Electron microscopy at different wavelength (A) at 500 μm, (B) at 157 200 μm, (C) at 100 μm, (D) 50 μm, (E) 1 micro meter. 37 FTIR analysis of CaCO3 with control. 158 38 XRD analysis of the polymorphs of Calcium carbonate. 165 39 Mn cycle of oxidation states found in nature 175 40 Four possible mechanisms of Mn+2 oxidation by bacteria. 176 41 Enzymatic pathway of Mn(II) oxidation 177 42 Kashmir Smast (Cave) entrance zone 180 43 Speleothems isolated from Kashmir smast (Cave) 181 44 Initial screening of Mn(II) oxidizing bacterial strains from cave soil. 187 45 Stereoscopy of the isolates. 188 46 DNA bands of Mn(II) oxidizing isolates 188 47 Phylogenetic anaylsis by Maximum Likelihood method 189 48 Growth curves of Bacillus pumilus C3 at 30oC and 25oC (No 190 o o Mn600, 30 C and No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30oC), after adding 1 and M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600) 49 Growth curves of Bacillus Safensis C6 at 30oC and 25oC (No 191 o o Mn600, 30 C and No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30oC), after adding 1 and M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600). 50 Growth curves of Bacillus pumilus C7 at 30oC and 25oC (No 192 o o Mn600, 30 C and No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30oC), after adding 1 and M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600). 51 Growth curves of Bacillus cereus C8 at 30oC and 25oC (No Mn600, 193 o o 30 C and No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30oC), after adding 1 and M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600). 52 Growth curves of Bacillus acidiceler C11 at 30oC and 25oC (No 194 o o Mn600, 30 C and No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30oC), after adding 1 and M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600). 53 Variation in Mn(II) oxidation at different pH and Ca+2 ion 195 concentration. 54 196 Mn(II) oxidation capacity and Mn(III, IV) oxide concentration as a 197 function of reaction time in C rich media K-medium. The ages of the 198 Mn oxide are from 4 h to 36 h. 199 55 Effect of metals on Mn oxide production by cavernicoles after 24 h 200 of incubation.
Acknowledgements Praise to ALMIGHTY ALLAH, whose blessings enabled me to achieve my goals. Tremendous praise for the Holy Prophet Hazrat Muhammad (Peace Be upon Him), who is forever a torch of guidance for the knowledge seekers and humanity as a whole. I have great reverence and admiration for my research supervisor, Dr. Fariha Hasan, Department of Microbiology, Quaid-i-Azam University, Islamabad, Pakistan, for her scholastic guidance, continuous encouragement, sincere criticism and moral support throughout the study. Her guidance helped me in all the time of research and writing of this thesis, with her patience and immense knowledge. I do not find enough words to express my heartfelt gratitude for Dr. Yuanzhi Tang, Professor in Department of Earth Science and Technology, Georgia Institute of Technology, Atlanta Georgia,USA. She supervised me during my studies in Georgia Tech University during IRSIP. This experience would not have been as valuable without the guidance, support and inspiration provided by her. I am impressed by her scientific thinking and politeness. I am also thankful to Alex, Sheliang, and Emily Saad Postdoctoral Research Associates at Atmospheric Science and Technology Department, for their care and immense help during my entire stay at Georgia Tech. I would also like to thank Higher Education Commission, Pakistan, for providing me grant under the Project “International Research Support Initiative Program (IRSIP)”. I am extremely grateful to the entire faculty at the Department of Microbiology, Quaid- i-Azam University, Islamabad. Many thanks to Dr. Frank Stewart and Josh at Georgia Tech for their kind help at Stewart lab. Extremely thankful to Dr. Muhammad Rafiq for his help during whole study. I extend my great depth of loving thanks to all my friends and lab mates (seniors and juniors) especially Imran Khan, Irfan Khan, Wasim Sajjad, Abdul Haleem, Arshad, Abdul Haq, Ghufran, Matiullah, Wasim (Bhatti), Waqas, Hameed Wazir, Dr. Akram, Dr. Malik, Amir Nawab(Ustaz Sb), Shakir, M. Aziz, Amin, Asim, Adnan, Saeedullah Jan, Afzal, Rafiqullah Umair, Akhtar Nadhman, Barkat, Faiz, Sana, Maliha, Khansa, Nazia, Naosheen, Aiman and Hifsa Saleema for their help throughout my study. I would like to thank my class fellows Sanaullah, Sultan, Rashid, Sohail, Pervaiz Ali, Aziz Khan, Ahmad Sadiq Hasan and Tariq for their help and support. A special thanks to my sweet USA memories makers Waqas Waheed, Amir Shafiq, Abdullah, Nauroz, Etizaz, Sherjeel Ahmad Usman,Usama(Abubakar) Ismail Khan, and Danial (PSA Sadar). A non-payable debt to my loving Ammi, Abbu, brothers and sisters for bearing all the ups and downs of my research, motivating me for higher studies, sharing my burden and making sure that I sailed through smoothly. Completion of this work would not have been possible without the unconditional support and encouragement of my loving family members. I would like to acknowledge my uncle Fazal Dad and Amir Nawab Khan for their moral and financial support. Finally, I express my gratitude and apology to all those who provided me the opportunity to achieve my endeavors but I missed to mention them personally.
Sahib Zada
Characterization of Bacterial isolates from Kashmir cave, Pakistan, and their Potential Applications.
PhD thesis by
Sahib Zada Department of Microbiology, Faculty of Biological Sciences, Quaid-i-Azam University, Islamabad 2017
Summary
Cave is a natural origin large underground enclosed place mostly in a hillside or inside the sea. Complete darkness inside the cave environment. Caves are of two types, sea caves and ground caves. Sea caves are generally short in length about 5 to 50 meters may exceed 300 meters. Due to low pressure and low oxygen concentration inside cave environment breathing is high. In large cave systems the air exchange is so high that air speed is about 80 miles per hour due to that, atmospheric gases are present inside cave, while ground caves are many miles in length.
Humans used caves for different purposes in early era for temporary shelter, for the celebration of rituals of passage, buried their wealth, source of different minerals, paleolithic painting, treasure hunting and as a historical landmark. Cave hosts diverse microbial and eukaryotic communities which have a very rich ecology. Pakistan has many caves which are still unexplored biospeleologically. The largest cave of Pakistan is Pir Ghaib Gharr cave in Baluchistan. In current study samples were collected aseptically from Kashmir smast Mardan/Buner. Samples were transported into Microbiology Research Lab, Quaid i Azam University Islamabad for further analysis. The geochemical analysis of the samples was carried out to determine the mineralogical composition of the samples. Bacterial strains were isolated by culture depended method using different media in laboratory. Different strains were selected on the basis of colony morphology. The isolated strains were further characterized and analyzed for different applications. The geochemical analysis is explained paper wise. A total 34 bacterial strain were isolated from samples collected from samples of Kashmir Cave on the basis of colony morphology. These isolates were further studied for their potential applications summarized in following.
Geochemical analysis and production of antibiotic: Bacterial strains having the ability to inhibit the growth of other bacteria were isolated from soil samples collected from Kashmir Smast (smast is Pushto for cave), Khyber Pakhtunkhwa, Pakistan. The study includes mineralogical and geochemical analyses of soil sample collected from the cave, so as to describe the habitat from which the microorganisms have been isolated. Total bacterial count of the soil sample was 5.25104 CFU mL−1. Four bacterial isolates having activity against test organisms Micrococcus luteus, Klebsiella sp., Pseudomonas sp., and Staphylococcus aureus were screened out for further study. Two of the isolates were found to be Gram-positive and the other two Gram-negative. The four isolates showing antibacterial activity were identified as Serratia sp. KC1- MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3-MRL, and Stenotrophomonas sp. KC4-MRL on the basis of 16S rRNA sequence analysis. Although all isolates showed antibacterial activity, only Bacillus licheniformis KC2- MRL was selected for further study due to its large zone of inhibition. Anti‐ bacterial activity of B. licheniformis KC2-MRL was optimum when grown in nutrient broth adjusted to pH 5 and after 24 hours of incubation at 35oC. The extracted antibacterial compound was stable at pH 5–7 and 40oC when incubated for 1 hour. The strain was found resistant against cefotaxime (ctx). Atomic-absorption analysis of the soil sample collected from the cave showed high concentrations of calcium (332.938 mg kg−1) and magnesium (1.2576 mg kg−1) compared to the control soil collected outside the cave. FTIR spectrum of the concentrated protein showed similarity to bacitracin. The antibacterial compound showed activity against both Gram-negative and positive test strains. Mineralogy of Kashmir Smast is diverse and noteworthy. Different geochemical classes identified by X-ray diffraction were nitrates, oxides, phosphates, silicates, and sulfates. Weathered cave limestone contributes notably to the formation of these minerals or compounds. FTIR spectroscopic analysis helped to identify minerals such as quartz, clinochlore, vermiculite, illite, calcite, and biotite.
Biodegradation of polyethylene: Low density polyethylene (LDPE) is used for making common shopping bags and plastic sheets and is a significant source of environmental pollution. The present study was aimed at testing the ability of bacterial strains identified as Serratia sp. KC1-MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3-MRL and Stenotrophomonas sp. KC4-MRL isolated from a limestone cave to degrade polyethylene. These strains were isolated from soil of Kashmir Smast, a limestone cave in Buner, Pakistan. These strains showed antibacterial activity against Micrococcus luteus, Klebsiella sp., Pseudomonas sp., and Staphylococcus aureus. The pieces of LDPE plastic were incubated along with bacterial strains for a period of one month and then analyzed. Degradation was observed in terms of growth of microorganisms used in consortia, chemical changes in the composition of LDPE by fourier-transform infrared spectroscopy, and changes in physical structure of LDPE by scanning electron microscopy. Maximum growth (107×105 CFU/ml) at 28°C and subsequent change in chemical and physical properties of plastic were observed in the presence of calcium and glucose. The cave-soil sample had a very high concentration of calcium. The microscopy showed adherence of bacteria with lots of mechanical damage and erosion on the surface of plastic films incubated with bacterial consortia. The spectroscopy showed breakdown and formation of many compounds, as evident by the appearance and disappearance of peaks in LDPE treated with bacterial consortia as compared to the untreated control. We conclude that antibiotic-producing cave bacteria were able to bring about physical and chemical changes in LDPE pieces and degradation of LDPE was enhanced in media augmented with calcium.
Bio-mineralization of CaCO3: Cave bacterial strains make significant contribution in the precipitation of calcium carbonate (CaCO3). In this section of study, it is shown that the CaCO3 precipitation is due to result of microbial metabolic activities. The isolated strains were used observe the CaCO3 precipitation by using B4 medium. A total of three bacterial strains showed the capability of CaCO3 precipitation on the selected medium. Bacterial cells with mineralization potential were molecularly identified through 16S rRNA gene sequencing as Bacillus toyonensis, Paracoccus Limosus and Brevundimonas diminuta. The most precipitates were observed at temperature and pH o of 25 C and 5. The precipitated CaCO3 was further confirmed by Scanning Electron Microscopy (SEM), X-ray powder diffraction (X-RD), and Fourier Transform Infra Red spectroscopy (FTIR) analysis.
Potential of enzymes production: All isolates were screened for different enzymes and isolate KC5- MRL was found to produce three industrially important enzymes; lipase, protease and amylase. The bacterial isolate was identified as Lysinibacillus sphaericus KC5-MRL (Accession No. KF010827). Optimum pH for the growth of Lysinibacillus sphaericus KC5- MRL, was around 7 and grew best at 35ºC. The optimum activity of lipase was observed at 30°C after 24 hr of incubation and pH 5 (42.23 U/ml). Maximum lypolytic activity (181.93 U/ml) was observed when 8% inoculum was used. Amylolytic activity of Lysinibacillus sphaericus KC5-MRL was optimum (15 U/ml) after 24 hr of incubation at 30°C. Proteolytic activity of L. sphaericus KC5-MRL was found to be 59 U/ml, after 48 hr at 30°C. Highest stability (42%) of lipase was observed at pH 10. pH stability of amylase showed highest activity at pH 7 i.e. 99.4%, whereas, protease stability was highest at pH 8. L. sphaericus KC5- MRL lipase, protease and amylase were stable at 35ºC and with residual activity as 118%, 104% and 107%, respectively. Triton X-100 and sodium dodecyl sulfate (SDS) stimulated the lipase and protease activities, whereas, Triton X-100 and T-80 stimulated ++ + ++ ++ amylolytic activity. Mg , NH4 and Ca stimulated the lipase activity and Zn + ++ ++ + showed highest inhibitory effect on lipase activity. Hg , Mg , Zn and NH4 reduced amylase activity, whereas, Na+ and Ca++ showed stimulatory effect. Hg+, Zn++, Ca++
+ + + and NH4 reduced protease activity but Na and Mg stimulated protease activity. Chloroform, formaldehyde, methanol and benzene stimulated amylase activity. Nitobenzen, methanol, benzene, and acetone stimulated protease activity. Ethylenediamine tetraacetic acid (EDTA) showed stimulatory effect on lipase. EDTA, Trisodium citrate TSC, mercaptoethanol, Phenyl acetaldehyde (PAA) and PMSF reduced amylase activity. All the modulators reduced protease activity except TSC and PMSF which stimulated its activity. The study concludes that these enzymes can be used for different purposes in various industries such as food and detergent.
Studies for Mn Oxidation by Cave Microbes: Mn (II) oxides are present abundantly in every environment, and very active in biogeochemical cycle of nutrients, carbon, contaminants, and other elements. It believes that bacteria play a key role in Mn oxides precipitation in environment. Manganese oxidizing bacteria (MOB) are reported mostly from marine or other aquatic environment, and few from terrestrial. The current knowledge is on the precipitation of Mn oxides by five Kashmir cave bacterial isolates B. pumilus C3, B. safensis C6, B. pumilus C7, B.cereus C8, and B. acidiceler C11. These Mn(II) oxidizing bacterial strains were isolated and purified on carbon rich K- medium. The Mn(II) oxidation by these isolated bacterial strains was enzymatically controlled reaction. The activity of Mn(II) oxidation was optimum at pH 5-7 and a temperature of 25-30oC and was lost at high temperature. Calcium ion (Ca+2) concentration affected the Mn(II) oxidation dramatically, while the Zn and Cu ions had no such high effect on the growth and Mn(II) oxidation. This demonstrates that cave bacteria are involved in the production of biogenic manganese oxides in cave environment. Introduction Cave is a natural large underground enclosed place mostly in a hillside or inside the sea. Complete darkness inside the cave environment. Caves are of two types, sea caves and ground caves. Sea caves are generally short in length about 5 to 50 meters may exceed 300 meters (Burcham, 2009). Due to low pressure and low oxygen concentration inside cave environment breathing is high. In large cave systems the air exchange is so high that air speed is about 80 miles per hour due to that, atmospheric gases are present inside cave, while ground caves are many miles in length (Barton et al., 2007). Cave study started in 17th century but later, before the mid of nineteenth century, the research value of cave environment focused its contribution to many other studies of science including geology, archaeology, chemistry, biology and geography. Visit to cave environment is difficult due to darkness, dangerous and no pre-indications (Crane and Fletcher, 2015). The cave ecosystem have constant temperature, high humidity and low nutrient availability with slightly acidic pH (Biswas, 2010). The internal maximum temperature of cave may rise to 10oC while the humidity is almost 100%. The scientists were of the opinion that caves have no life due to darkness, limited nutrients, acidic condition and presence of H2S in air, later it was observed that they were not right (Engle et al. 2008). The scientific study of cave is known as speleology and biospeleology is the study of life in cave (Moore and Nicholas, 1967). The study of caves covers the cave features, make-up, structure, history, life forms and its formation. The process by which the caves are form is called speleogenesis. Speleology is interconnected study with chemistry, biology, geology, physics, meteorology and cartography. A French scientist named Edouard-Alfred Martel (1859-1938) was the first who studied the modern speleology who is considerd the father of modern speleology. He was the founder of an association called Societe de Speleologie founded in 1895 the first ever association for cave study. The advancement of cave study is linked with caving sport because of taking interest by public and awareness, mostly spelelological field works are conducted by sport cavers. Cave study is known with different names in different countries like spelunking in USA and Canada and in some areas cave study is called potholing (Engle et al. 2008). Humans used caves for different purposes in early era for temporary shelter, for the celebration of rituals of passage, buried their wealth, source of different minerals, paleolithic painting, treasure hunting and as a historical landmark (Hayden and Villenueve, 2011). Types of caves
On the basis of cave formation and rock type, caves have been classified into several types. The most common types of caves are formed in limestone rocks (calcareous), and in basaltic rocks (lava tubes). Other cave types are limited in range and include those in quartzite, granite, ice, talus, gypsum and sandstone (Palmer, 1991).
The two general types of caves (terrestrial and underwater) are shaped by the dissolution and characterized as limestone caves. Limestone caves are shaped due to the rock dissolution in weak acids, which exist naturally in ground water. Weak acid is used to solubilize these rocks just like sugar in tea, but dissolves steadily and slowly. Cracks are formed as a result of ground water which then increases in size and form a full cave.
These caves may overflow with water when groundwater increases in a level higher than a cave. In some cases, the whole cave or some part of it fills with water (Barton and Jurado, 2007).
Solutional cave: Solutional caves are formed in soluble rocks like limestone, chalk, dolomite, salt beds, marble or gypsum. This type of cave is mostly formed in terrestrial environment. The largest cave in the world is solutional cave. The soluble rocks dissolve by the action of rain and ground water charged with carbonic acid (H2CO3) and natural organic acid present in environment. Solutional caves are adorned with
CaCO3 formed via precipitation, which contain stalactites, stalagmites, flowstones, helictites, soda straws and columns. Speleothems are the secondary minerals deposited in solutional caves, whereas the sulfurous fumes formed by the reaction of H2S gas and oil reservoirs results in the formation of secondary types of solutional caves. The H2S gas react with water and form sulfuric acid (H2SO4) which dissolves the rocks and as a result the solutional cave is formed (Burcham, 2009). Ice cave: Any cave which have significant amount of perennial ice inside cave and have at least a portion at temperature less than 0oC throughout the year. An Englishman, Edwin Swift Balch in 1900, described for the first time an ice cave and suggested the word “glacier” for this type of cave. Now the term “ice cave” is commonly used for the cave containing year-round ice. Cave formed in ice is properly called glacier cave. The surface of the rocks is thermally insulated due to which the surface temperature is nearly constant (Ford and William, 1989). Lava cave (Lava tube cave): The formation of lava tube cave mechanism is not clearly understood. The discovery of lava tube is one of the major mechanisms of building shield volcanoes by separating the flowing lava and dispensing it away from the vent (Swanson et al., 1971). The strong indirect evidences of huge lava tubes are thought to be present on moon (Greeley, 1971) have given the importance to understand the phenomenon of the formation of lava tube. Different theories exist about the lava tube formation. Ollier and Brown (1965) suggested that lava tubes require laminar flow of the lava. Lava tubes are mostly formed in basaltic lava flows. The basaltic lava deposits differ in viscosity, heat and gas contents. As the flow become cooler, the gas evaporates, slow in motion, and more viscous than pahoehoe flows. This flow cracks the crust into huge pieces of clinker which are pushed and buried by the flow. This type of flow forms cave, the clinker may offer routes for microbes to cave. Lava tube cave range in size from tiny, finger size tunnels to cave having length more than 6000 m and diameter of 17 m (Howarth, 1973). Sea cave: This type of cave is also called littoral cave, and is formed by wave action in zones of weakness in sea cliffs. Sea caves are present everywhere in the world. Coasts of Norway are rich of sea caves mostly 100 feet above sea level (Rabbe, 1988). The largest sea caves were discovered in the west coast of United State, the Hawaiian Island and Shetland Islands; while the world’s largest one is Matainaka cave having a length of 1.5 km located at Otago coast, New Zealand (Barth, 2013). Sea caves are formed in sea side rocks i.e. sedimentary, metamorphic and igneous, having a weak zone. Sea Caves are formed by mechanical erosion produced by wave actions which break the rocks. Some caves are formed in carbonate rocks in littoral zones and expand by littoral process but its origin is by dissolution which are called hybrid caves (Mylroie and Mylroie, 2013). Rain water, carbonic and organic acid leached from soil also play key role in cave formation by weakening the rocks.
Cave genesis The process of cave formation is called speleogenesis and the caves are formed by different geological processes like combination of different chemical reactions, dissolution, mechanical weathering, melting of ice or glacier, corrosion by water, tectonic forces, microorganisms, pressure, atmospheric influences and also by excavating (Engel et al., 2004). Dissolutional caves are formed by water flowing on and through limestone rocks which can be formed from epigenic (top-down process) or hypogenic (bottom-up process). Caves formed in basalt or sandstone rocks are categorized as pseudokarst (Palmer, 2007). Erosional caves are formed by mechanical scouring or wave action rather than by dissolution. Dissolution caves can transit to erosion caves with the passage of time. Sea caves develop by erosion along sea cliffs and anchialine caves form by dissolution along sides. Lava tube caves are different than dissolution caves which are formed from cooled crust around the flowing lava.
Fig. 1.1. Different types of caves and its formation including dissolution and weathering (White and Culver, 2000).
Fig. 1.2. Solutional cave formation (Cooper et al., 2007).
Fig. 1.3. Limestone cave formation (Williams, 2016). Traditionally, cave genesis is mostly from dissolution of minerals by water. Rock solubilization is a fundamental process in cave formation but not adequate geological prerequisite for cave development. The potency and velocity of mineral dissolution by water depend on climatic parameters e.g. speleogenesis is faster in humid and warm conditions (Priesnitz, 1974). The solubilization process is mineral specific i.e. in limestone cave the carbonate (calcite and aragonite), and dolomite [CaMg(CO3)2] are the most soluble minerals. Role of cave microbes in minerals deposition The role of microbes in the decomposition of minerals is directly or indirectly and utilize these decompose minerals for their growth. In acidic condition microbial dissolution of sulfide minerals produce more than 1,000 times Acid Rock Drainage than chemical reaction alone. These microbes take part in minerals precipitation, adsorption or metals release, and decomposition/formation of organic-metallic compounds. Cave microbes use soluble minerals as oxidizing agent, electron acceptors/donors in redox reactions, and involved in metabolism (Adams, 2005).
In cave pyrite (FeS2) is oxidized by ferric iron or oxygen and microbes can enhance the rate of pyrite oxidation. Williamson et al, (1994) derived a law which state that abiotic oxidation of pyrite in enhance by increasing the concentration of oxygen and slightly by decreasing pH. The oxidation of pyrite by microbes begins to exceed at about pH 3.5 – 4. The Oxidation of pyrite of microbial origin by ferric iron is many times faster than abiotic oxidation by oxygen at pH 2 (Nordstorm and Alpers, 1999). The temperature also plays an important role in biotic and abiotic rate of oxidation. The acids are produce only in the presence of iron fraction. Other minerals like gypsum and carbonate present in the site also contribute in weathering of rock pile stability or instability. Acid formation is a result of oxidation of pyrite which contains iron and sulfur. The following reaction illustrates the reaction of pyrite (solid) and oxygenated water.
2+ 2- + FeS2 + 3.5O2 + H2O Fe + 2SO4 + 2H
According to this mechanism the formation of reduced iron and acid takes place. This environment (Reduced iron and presence of Oxygen) favor the growth of acidophilic, autotrophic microbes (Iron and Sulfur oxidizers) A number of iron and sulfur oxidizer bacteria are reported from caves which include Acidithiobacillus species are very common known as Thiobacillus ferrooxidans, Leptospirillum ferrooxidans and sulfolobus acidocaldarius and some colorless sulfur-oxidizing bacteria (Ehrlich, et al, 1991).
These microbes play a key role in biogeochemical cycles and minerals leaching, the bacterium T. ferrooxidans is a model bacterium for iron oxidation studies (Johnson, et al, 1993). The mechanism of acid production is believed that iron oxidizers (T. ferrooxidans) catalyze by oxidizing iron (Fe+2) to ferric iron (Fe+3) which act as oxidant of pyrite, and create more reduced iron, sulfate and acid in huge amount as describe in below reaction (Evangelou and Zhang, 1995).
2+ + 3+ 4Fe + O2 + 4H → 4Fe + 2H2O
3+ 2+ 2- + FeS2 + 14Fe + 8H2O → 15Fe + 2SO4 + 16H
In biologically acid production the Fe2+ act as rate limiting factor during these processes (Singer and Stumm, 1970). There is also a completion for oxygen and other nutrients among iron oxidizers bacteria including T. ferrooxidans and other microbes of the habitat. Reports supported the idea that the genera i.e. Thiobacillus, Leptospirillum, Sulfolobus and some color colorless sulfur-oxidizing bacteria, with Thiobacillus sp. are involved in the oxidation of sulfide into sulfur (Ehrlich et al,, 1991; Wichlacz and Unz, 1981; Harrison, 1984). The rate limiting step is mediated by the enzyme sulfate oxidase (enzyme cofactors are Mo and Fe).
CO2 +2H2S → CH2O+2S+H2O
2CO2 + H2S + 2H2O → 2(CH2O) + H2SO4 Oxidation of Sulfur and Related Compounds
2S+3O2+2H-OH → 2H2SO4 (Thiobacillus thiooxidans)
12FeSO4+3O2+6H-OH → 4Fe2(SO4)3+4Fe(OH)3 (T. ferrooxidans)
Microbes oxidized sulfur at a variety of pH 8.5 to 1.9, lower pH favor the oxidation of sulfur by different microbes, once the pH is lower by the production of sulfate by some microorganisms this may involve in the activation of other group of microbes and take part in the oxidation. Therefore succession of microbes takes place as the pH of soil is lowered by the production of sulfate (Ehrlich, et al. 1991, Harrison, 1984). The acid production from the oxidation of pyrite by microbes also increases the dissolution of minerals present in cave. The lithological studies of cave rocks revealed that cave rocks compose of carbonate minerals (calcite) and silicate minerals (plagioclase, K-feldspar, hornblende, biotite, augite). Microbes used the oxidized metal as electron acceptors during autotrophic processes these metal may include arsenic, cadmium, calcium, copper, iron, magnesium, manganese, phosphate, potassium, selenium, sulfur, uranium, and zinc.
Different studies can be merged to find out the effect of microbes on rocks weathering and possible role in mineralization. The study of cave rock and cave microbiology should also helpful in strategies of increase rock stability and decreasing or increasing water penetration into the rock.
Caves of the world Caves hides the beauty and splendor of nature like the depths of ocean. Caves are present mostly in mountain or under the earth. Caves are located everywhere in the world. The largest cave system in the world is Mammoth cave located in Kentucky, USA. According to the National Speleological Society (NSS) survey in January 2016, the enlisted caves are the world largest caves systems (Gulden, 2016).
Table. 1.1. List of largest and most studied caves world wide Length Cave Name Location Discovery Reference (km) Mammoth cave 651.8 Kentucky, USA 1791 Sistema Dos Ojos Cave 335.0 Quintana Roo, Mexico 1987 Jewel cave 289.8 South Dakota, USA 1900
Sistema Ox Bel Ha cave 257.1 Quintana Roo, Mexico 1996
Optymistychna cave 236.0 Korolivka, Ukraine 1966 Gulden, 2016
Wind cave 229.7 South Dakota, USA 1881 Lechuguilla cave 222.6 New Maxico, USA 1900
Clear water cave 215.3 Sarawak, Malaysia 1978
Fisher Ridge cave 200.5 Kentucky, USA 1981 Holloch cave 200.4 Moutathal, Switzerland 1875
List of largest caves in Pakistan Like other countries Pakistan has many caves which are still unexplored biospeleologically. Pakistan Cave Research and Caving Federation is a national body established by Mr. Hayatullah Khan Durrani who also represented Pakistan in Union of International Speleology (UIS), and British Caving Federation (BCA).
Table. 1.2. List of caves in Pakistan Cave Name Length (km) Location Pir Ghaib Gharr cave >1.20 Balochistan
Kashmir smast (cave) 0.188 Mardan KPK Gondrani cave Unknown Bela, Balochistan Bhaggar cave Unknown Azad Kashmir, Pakistan Juniper Shaft cave Unknown Balochistan Mughagull Ghara cave Unknown Balochistan Mughall saa cave Unknown Balochistan Mangocher cave Unknown Balochistan
Microbial community in cave environment The common types of caves throughout world are limestone caves made in limestone rocks and basalt rocks. Caves have poor nutrients oligotrophic ecosystem (below 2 mg of TOC per liter), having low temperature, complete darkness and high humidity. The average number of microbes in cave ecosystem is 106 cells/gram of rock (Barton and Jurado, 2007). Photosynthetic organisms are found only in the entrance zone of cave but some are also present inside due to artificial light mounted by public. Due to light absence in cave the primary production of organic substances by microbes are negligible. But still there are different chemolithoautotrophic processes occur and microbes are potentially involved in such processes. Microorganism in such environment used hydrogen, nitrogen or other organic compounds as well as reduced form of metals found in the cave rocks around like iron and manganese (Gadd, 2010, Northup and Lavoie, 2001). The allochtonous matters source is water which leaking through cracks in rocks from soil above the rocks, or air which transfers the organic particles. Besides these plant roots, anthropogenic activities, remnants of mammals may provide organic compounds which promote the growth of heterotrophic microbes. Microbial community structure of caves is influenced by many factors include pH, availability of nutrients, light, oxygen, different metals compounds, water, and susceptibility of substrates to colonization (Engle et al. 2010; Jones and Bennett, 2014). Due to low nutrients availability, low temperature, high humidity highly adoptive and extremophiles microbes can only survive in cave environment (Rothschild and Mancinelli, 2001). Most microbial communities depend on the energy and carbon fixation of photosynthesis. Ageless darkness avoid the phototrophs colonization in cave environment (Barton and Jurado, 2007). The chemoautotrophs are the subordinates of the caves climates which import energy into caves by fixing carbon (Sarbu, et al., 1996). Bacteria, archaea and fungi are the most founded biodiversity in caves and omnipresent in cave habitats such as stream water, soil, rocks surfaces and sediments (Engel, et al., 2004). Inside caves environment many bacterial phyla have been reported by using 16S rRNA genes sequencing (Ortiz, et al., 2013). The most abundant taxa on caves walls are proteobacteria, Actinobacteria and Acidobacteria (Barton, et al., 2007; Pašić, et al., 2010; Cuezva, et al., 2012). Bacterial diversity of cave soil and sediment could be comparable (Ortiz, et al., 2013), while the rock surfaces have lowest microbial community (Macalady et al., 2007; Yang et al., 2011). Cave microbiota is highly variable due to microhabitats of cave. In a single cave different bacterial diversity and rocks composition were observed possibly due to the geochemistry of rocks (Barton et al., 2007). Nutrients availability and disturbance also affect the microbial diversity inside a cave. Microbes and organic matter could be pipe in into caves by air flow, water seepage, and may be by entrance of human and animals into caves (Shabarova et al., 2013). Eight different microbial phyla were identified by Pasic et al, (2010) dominated by Proteobacteria followed by Actinobacteria, Nitrospira, Acidobacteria, Chloroflexi, Gemmatimonadales, Verrucomicrobia and Planctomycetales (Fig. 1.4).
Fig. 1.4. Distribution of major groups of microbial communities in cave environments by 16S rRNA gene sequencing (Pasic et al., 2009) Sulfidic caves environments are appreciable for the chemotrophic organisms and colonized by a thick filamentous microbial mat. These microbes oxidize sulfur for the energy requirement (Boston et al., 2006). The sulfur oxidation process by microbes is characterized by water rich in H2S source and limited in oxygen (Chen et al., 2009). (Engel, 2007) have reported that the ε-Proteobacteria along with γ-, β-, and α- proteobacterial are the most sulfur oxidizer in sulfidic cave system. Some nitrite and ammonia oxidizers were also isolated from Movile cave Romania (Chen et al., 2009). Many methylotrophic microbes such as Methylothenera, Methylophilus and Methylovorus were also reported in the same research. These methylotrophic were related to degradation of chitin like substances or methanogenesis.
Table. 1.3. Cave microbes isolated from different caves in different country through out the world
Caves Country Microbes Proponents/ Year Nullabor Caves Australia α, β, γ and δ-Proteobacteria and novel microorganisms Holmes et al. 2001
Altamira Cave Spain Proteobacteria, Plantomycetales, Schabereiter-Gurtner et al. 2002 Cytophagal/Flexibacter/Bacteroides, Acidobacterium, Cuezva et al, 2009 Actinobacteria and green-sulfur bacteria Reed Flute Guilin, Guangxi, China Knoellia sinensis and K. subterranea (Actinobacteria) Groth et al. 2002
Lower Kane Cave Wyoming, USA ε-Proteobacteria Engel et al. 2003 Lechuguilla Spider Carlsbad Caverns National α, β and γ-Proteobacteria, Enterobacteriaceae, Northup et al. 2003 Park New Mexico, USA Xanthomonas, Bacillus/Coliform group and Lactobacillaceae Llonin La Garma Asturias, Northern Spain Proteobacteria, Actinobacteria, Gram-positive bacteria Schabereiter-Gurtner et al. 2004
Kartchner Caverns Arizona, USA Bacillus, Brevibacillus, Rhizobium, Sphingomonas, Ikner et al. 2006 Staphylococcus, Pseudomonas, and other uncultured β-Proteobacteria
Engel et al, (2004) assessed three aphotic springs in cave with regards to bacterial diversity using 16S rDNA phylogenetic analysis and found the bacterial diversity was very low but the most dominant taxonomic group in sulfidic cave spring was Epsilonproteobacteria (68%) affiliated bacteria. The other affiliations of the bacterial strains were with Gammaproteobacteria (12.2%), Betaproteobacteria (11.7%), Deltaproteobacteria (0.8%), and the Acidobacterium (5.6%) and Bacteriodetes/Chlorobi (1.7%) divisions shown in (Fig. 1.4). Cave microbes and biomineralization The role of cave microbes in cave ecosystem is valuable since these microbes are able to colonize rock surfaces and to use organic and inorganic compounds as energy sources. These microbes are interesting for the study of biomineralization process due to stable microclimatic conditions in the cave, where bioinduced mineral fabrics are conserved without important diagenatic modifications (Canaveras et al., 2006; Sanchez-Moral et al., 2006). Caves rocks and soil host a wide range of metals, among which iron is one of the dominants. Oxidized iron is found in deposits while reduced form of iron is present in cave water. Iron oxidizing microbes such as Leptothrix, Gallionelle and Siderooxidans were found to reside in sediments (Peck, 1986), water (Moore, 1981), and in speleothems (Kasama and Murakami, 2001). These microbes induce iron mineralization at the point of contact of seeps and springs with oxygen. Bacterial mineralization was found four times more on ferrohydrite speleothems (Kasama and Murakami, 2001). Although the role of bacteria in biominralization and iron ores formation has rarely been studied (Wu et al., 2009; Baskar et al., 2012). Carbonates precipitation by cave microbes In the history of earth, microbes play an important role in mineral deposition and are active in biomineralization (Ehrlich, 1996). Bacterial strains isolated from different environments were found to possess the capability of precipitating calcium carbonate in both laboratory and natural condition (Morita, 1980; Rivadeneyra et al., 1993). The phenomena of calcium carbonates precipitation in cave environment has been reported in many research studies in last two decades (Castanier et al., 1999; Forti, 2001; Barton and Northup, 2007). Precipitation may be enhanced by removal of crystallization inhibitors by bacteria (Bosak and Newman, 2005), shifting of pH of micronutrient environment around the microbes, or nitrogen release and fixation (Castanier et al., 1999; Hammes and Verstraete, 2002; Cacchio et al., 2004). It has also been reported by
several researchers that bacterial cell pump the Ca2+ ion from inside to outside medium to avoid the toxic effect of calcium concentration (Cacchio et al., 2004; Cai and Lytton, 2004). Calcium precipitation has been correlated with metabolic process and with microbial cell wall structure (Mastromei et al., 1999).
It has been proposed that precipitation of calcium carbonate by microbes could be considered to avoid or prevent emission of CO2 (Sharma et al., 2008). It has been reported that bacteria play an important role in crystal nucleation, (Sanchez-Moral et al., 2012). Several studies on calcium carbonate precipitation reported different mechanisms for the microbial mediated carbonate precipitation. Minerals adsorption to surface of cell could trigger microbial mediated biomineralization, in which bacterial cell act as nucleus for precipitation (Buczynski and Chafetz, 1991; Rivadeneyra et al., 2010).
Manganese oxidation by cavernicoles
Manganese is the fourth widely distributed element in the Earth’s crust, and occurs in different oxidation states from -3 to +7. Manganese is important element for all living things. Only three form of Mn are significant for living things i.e +2, +3, and +4. The +2 form of Mn is soluble in earth crust while +3 and +4 are in precipitated form. The oxidation/ reduction of Mn is catalyzed by microbes. The Mn oxides are reduced by many anaerobic bacterial strains either by acid production or by production of reducing substances such as sulfides or these microbes use the oxidized metal as an electron acceptor in respiration (Lovley 1991; Nealson and Myers 1992; Nealson and Little 1997). Many aerobic microorganisms are reported to catalyze the Mn+2 oxidation (Ehrlich 1984; Ghiorse 1984). Microbes accelerate the Mn+2 oxidation five times as compared to abiotic Mn oxidation (Nealson et al. 1988; Tebo 1991; Wehrli, 1990). In terrestrial environment, a continuous oxidation reduction cycle of Mn takes place in oxic/anoxic condition. In such environment, Mn acts as a redox shuttle in oxidation reduction of organic carbon (Nealson and Myers 1992).
Applications of cavernicoles The cave microbes have many industrial applications. Cavernicoles are used for the preservation of ancient testimonials and sculpture via the identifications of microbial species which precipitate the protective coating of calcite. Cave environment is
nutrients limited in which the microbes compete for nutrients and fight for survival. Due to this ability of cavernicoles, these microbes produce metabolites against other microbes. Therefore, these microbes are used for the production of novel antimicrobial compounds. Besides antibiotics these microbes also produce antifungal and anticancer compounds which are the basic need of today medical science. Cavernicoles play a significant role in biomineralization. These microbes precipitate calcium carbonate five times more than outside microbes. The rate of manganese oxidation by cave microbes is more significant than other environment microbes. The strange character of cave microbes is the ability of bioremediation. The cave microbes are also a possible good source of beneficial compounds like extremozymes (Singh et al., 2011), biosurfactants (Banat et al., 2010), antitumorals (Chang et al., 2012), exopolysaccharides (Nicolaus et al., 2010), radiation-protective drugs (Singh and Gabani, 2011). Other applications of these microbes are they take part in biospeleogenesis and biocementation.
Aim and Objective The aim of the study was to conduct initial assessments of bacteria which can produce antibiotics, industrially important enzymes, degrade plastic, have ability to mineralize manganese and carbonate and geochemistry of caves in Pakistan. The objectives of the study were To collect soil, guano and speleothem samples from Kashmir cave using standard microbiological procedures To analyze soil, guano and speleothem samples geochemically. To isolates microbes and screen for the production of antibacterial compounds and industrially important enzymes. To study the efficiency of isolated bacterial strains to degrade polyethylene plastic.
To screen the cave bacteria for their ability to mineralize manganese and carbonate precipitation.
Literature Review
Cave
Cave is a natural underground hollow space large enough for an individual to enter. Most caves are formed naturally by weathering of limestone rocks and extend deep underground. Initial reports about study of cave trace dates back to 17th century and mid of 19th century. Biospeleology is a study of microbial life and its interaction with all other branches of natural sciences i.e. chemical sciences, geological sciences, materials sciences, environmental sciences and also describes microbes’ interaction with the minerals in the caves. Globally, the ratio of caves is very less and its access is limited due to danger, hard environment, darkness and no preindications (Crane and Fletcher, 2015).
Cave environment has a stable constant interior temperature, low availability of nutrient, excess of humidity and acidic pH (slightly) (Biswas, 2010). The humidity raises up to 100%, while the maximum temperature may raise to 10oC inside the cave. Initially, scientists thought that due to low content of nutrient, no source of light and acidic condition, caves are not suitable to host life but the hypothesis was proved wrong later (Engle, 2015).
For the geological study of mankind and earth, caves provide a way of collecting information and to determine Earth’s biological history, paleontologists utilized fossil data. In the past, caves used to be a hiding and living place for people, while in present era caves are utilized for vegetable and fruit storage and mushroom growth. Fermentation of wine and cheese required constant temperature and dark environment thus caves are a suitable reservoir for that. Due to constant temperature and high humidity people in the past used caves as hospitals, to treat patients with respiratory illnesses.
Since 1940 bat feces are being used as fertilizer. Scientists are using natural resources and habitat of cave for human benefits. For protection and conservation of cave resources for the upcoming generations, cave study is very important (Burcham, 2009).
Edouard-Alfred Martel, a French scientist, was the pioneer of cave study and
considered as father of speleology. Societe de Speleologie, the first association related to the study of cave, was founded by Edouard-Alfred Martel and his colleague in 1895.
Geochemistry of cave
Caves are the most stable component of the environment which acts as a reservoir for different material like physical, biological and chemical deposits. The most important chemical deposits are speleothems. Besides, these caves have unique and ancient biology which very rarely are interrupted by humans. Ancient people used caves as a shelter and for the search of different important substances not available from other places on Earth but caves were rich source of these substances. From prehistoric to present times, endorsed the idea of uses of caves for different minerals and other important compounds. Some evidences suggested that ~30,000 years ago humans used caves for searching of pigments like iron and manganese oxides and hydrooxides for different ornamental uses like paints. Later on the cave mining was expanded for the mineralogical studies (Larocca, 2005).
Most of the caves are composed of calcium carbonate, a few natural cavities have been studied for the minerals in detail, about 350 cave minerals have been investigated in which some are new to science (Onac and Forti, 2011).The presence of such new unique compounds may be due to the corrosion of rock material with water before entering into the caves and remain as sediment over there. Different mechanisms are involved in different caves for mineralogical reactions these mechanisms include; evaporation, dehydration, oxidation, sublimation, hydration, deposition of particles from aerosols and vapors, and segregation. The driving parameters for these reactions are temperature and pH of the cave environment. Another most important driver of such reactions is microorganisms. The stable environment of cave allows the formation of huge crystals alongwith more common small numerous aggregates of different compounds (Onac and Fort, 2011).
More common compounds like calcite and gypsum and less common like vanadate are identified from different caves along with other more than 250 minerals (Hill and Forti, 1997). About 4000 years ago Assyrians around Tigris River used Niter from caves for food preservation. Similarly, Native Americans search mirabilite, epsomite and gypsum in Mammoth cave, USA (Farnham, 1820; Broughton, 1972; Tankersley, 1996).
Recently, halite caves of Atacama were mined for salt supply to the country (De Waele et al., 2009). Another most important material called Guano (phosphate rich source) used as natural fertilizer was extracted from caves worldwide (Abel and Kyrle, 1931; Badino et al., 2004; Onac et al., 2007).
It must be emphasized that the genesis of many of the minerals found in caves is unrelated to the existence of the cave itself; they were often brought in by owing or seeping water or were exposed by the corrosion of the cave walls. By definition “a cave mineral is a secondary deposit precipitated inside a human- sized natural cavity”, where secondary means, a mineral derived from a primary mineral existing in the bedrock or cave sediment through a physicochemical reaction (Hill and Forti, 1997). The generic term that encompasses all secondary deposits formed in caves is speleothem (cave).
As cave speleothem is composed of 90% of calcite and aragonite therefore caves have attracted people for important mineral extraction. For the first time, White (1961) studied cave for minerals. About 350 types of minerals were identified from different caves (Hill and Forti, 1997; Back and Mandarino, 2008).
Calcium carbonate is one of the major chemical compounds found in caves. Besides, many other chemical compounds are found in cave. Many of the compounds were first identified inside cave and later they were detected outside the cave as well (Garavelli and Quagliarella, 1974; Martini, 1978, 1980a, 1980b, 1983, 1992; Bridge and Robinson, 1983; Back and Mandarino, 2008). The main reason for this is the interaction of water with different chemical compounds prior entry into the cave. Basically four main types of solutions which interact with cave bedrocks may include Connate (water and minerals ions trap in pores of sedimentary rocks), Juvenile (water rich with volatile fluids), Meteoric (water derived from precipitation of snow and rain) and seawater. Temperature and pH of the interaction point also greatly influence the concentration of mineral species and dissolution.
It is a known fact that the major carrier for minerals and chemicals into the caves is water, but some sources like lava, vents, thermos mineral compounds etc. also plays role in mineral ion transportation into the cave, which help in the generation of different minerals inside cave by different minerogenic and biological mechanisms (White,
1997; Onac, 2005; Forti et al., 2006). The driving force for such reactions and processes are temperature, relative humidity, and carbon dioxide concentration, and are greatly enhance by endemic microorganisms.
Dissolution/Re-precipitation
The dissolution and re-precipitation processes involve the dissolution of minerals with water current prior enter into caves and after entry, the caves temperature control the evaporation and the minerals precipitate again. When water vapors evaporate, rocks 2+ + 2+ + 2- interact with them and release of Ca , Na , Mg , K and SO4 ions occur which diffuse into solution. When these mineral filled solutions enter into caves, this water evaporate and minerals precipitate again. By this mechanism the formation of speleothem also takes place with the soluble minerals such as halite, epsomite, mirabilite other sulfates (De Waele et al., 2009) and in limestone and/or gypsum cavities throughout the world (Forti, 1996a). Similarly, the precipitation of opal found in large quantity in quartzite caves of Venezuela (Forti, 1996b).
The mineral dissolution and re-precipitation mechanism also occur in volcanic cavities. When the temperature of lava decreases, water solubilizes the minerals by entering into the cavities via fissures or porous surfaces and dissolves various salts. When this solution exposed to hot environment of caves, it rapidly evaporates and allows the re- precipitation of dissolved mineral.
Geochemical reactions in caves
The weathering processes involve the aerobic and anaerobic reaction mechanisms leading to deposition of mineral ions and other compounds. The most important and abundant compounds are oxides and hydroxides, sulfates, carbonates and nitrates. The mechanisms involved in these depositions are oxidation/reduction (redox), hydration/dehydration and double replacement reactions.
Oxidation/Reduction
When the ground water penetrates in rocks, it changes the environment to anoxic and slightly acidic which favors the conversion of pyrite, sulfide, and limestone to a number of oxides and hydroxides by oxidation. The oxidation of hydrogen sulfide to sulfuric
acid is also reported from sulfidic caves, which extremely decrease the pH of the cave environment and formation of sulfur speleothem occurred (Forti and Mocchiutti, 2004).
Comparatively, the reduction process in cave is less common than oxidation (White, 1976). The reduction reactions are occurred anaerobically, in which the organic compounds are reduced for energy uptake. In some caves the geological boundary is developed which switching off and on the oxidation and reduction mechanism called ‘bohnerz’ (Seemann, 1970; Forti, 1987; Onac, 1996).
Most of the redox reactions are mediated by microbes e.g. reduction processes involved in the reduction of sulfur, manganese, iron and nitrogen.
Hydration/Dehydration
Hydration and dehydration is the condition of incorporation and removal of water molecule from any mineral which affects the state of that particular mineral. Caves are considered one of the humid environments containing more than 85% humidity in most of the caves. In some caves the evaporation process increase in part of the cave or in whole cave which lead to removal of water molecules from some hydrated minerals and convert into different form. Similarly, in humid part of the caves some dehydrated minerals may absorb water molecule from surrounding and covert into hydrated form of minerals which may affect their crystalline structure.
Common example of hydration and dehydration is hydrated mirabilite
(Na2SO4·10H2O) and dehydrated thenardite (Na2SO4) (Bertolani, 1958). Similarly, the epsomite (MgSO4.7H2O) convert into hexahydrite (MgSO4·6H2O) by the release of one water molecule and further release of water molecule convert it into kieserite
(MgSO4·H2O) (Bernasconi, 1962; White, 1997). Example of some other hydration and dehydration processes are bassanite (CaSO4·0.5H2O) hydrated into gypsum
(CaSO4·2H2O) (Forti, 1996). Martini (1980a) also reported the conversion of mbobomkulite (monoclinic mineral of the chalcoalumite (Ni, Cu)Al4[(NO3)2,
SO4]2(OH)12 • 3H2O group) into hydrombobomkulite (A monoclinic sky blue mineral containing aluminum, copper, hydrogen, nickel, nitrogen, oxygen, and sulfur).
Double replacement reactions
In double replacement reaction, the interchange of a part of two compounds with each other, which lead to the formation of two other compounds. As most of the caves like limestone caves are composed of carbonates so frequently carbonate is involved from one side and strong or weak acid at other side in such reactions. Strong acid favors the oxidation processes and carryout the formation of sulfuric acid from pyrite of H2S (Pohl and White, 1965; Hill, 1987), and phosphoric acid and nitric acid in environments where guano is present (Hill and Forti, 1997; Onac, 2000).
The most common final mineral products (as a result of double displacement) are nitrate, phosphate and sulfate. Such minerals are involved in the generation of a number of speleothems like, moon milk to flowstone and a number of crystals like helictites to euhedral. Similarly, gypsum is the common mineral precipitated by double exchange reactions.
Karst
The karstification and speleothem formation typically occurs when underground water molecules react with carbonate rocks. This phenomenon can be expressed by a single chemical reaction as;
2+ - CaCO3 (solid) + H2O + CO2 (gas) <------>Ca (aqueous) + 2HCO3 (aqueous)
In this reaction the gaseous CO2 reacts with limestone and causes its dissolution, while on the other hand reaction occurs to precipitate the calcite and aragonite to form speleothem and release the CO2 into environment. Approximately, 95% of speleothems (calcite and aragonite) are precipitated in this manner, and it is known that about 97% of deposits are composed of calcite and aragonite in caves (Dreybrodt, 1988; Hill and Forti, 1997; Palmer, 2007; Onac, 2011).
This phenomenon is high in carbonate caves, but same process occurs for mineral deposition in caves other than carbonate.
The first condition is almost always available in a cave while the second one totally depend on the dissolution of Ca2+, Mg2+ etc. in the water seeping into the caves. The second condition almost acts as a limiting condition for the formation of speleothem in caves.
Microbe - mineral interaction
It is a known fact that microbes are found everywhere in the world. Ability of microbes to adapt to the environment and play key role in biological and geological processes made them unique from all other biological species. It is also known that microorganisms have their impact on the development of secondary chemical deposits in caves (Shaw, 1997). In early stage of cave explorations it was believed that the speleothems e.g. stalactites, stalagmites, coralloids, pool fingers, etc. have the ability to grow like plants (Tourneford, 1704). The understanding of microbial – rocks interaction clearly demonstrates the role of microbiota in the development of caves and deposition of minerals (Northup and Lavoie, 2001; 2004; Barton, 2006; Jones, 2010).
Studies on the microbial role demonstrate that enzymes produced by microorganisms are directly involved in the biomineralization of minerals which lead to the precipitation and accumulation. These enzymes may change pH of the surrounding or enhance the reaction speed (Northup et al., 1997; Boston et al., 2001).
Different types of microbes are found in the caves. These may include chemolithotrophs and chemoheterotrophs. Mainly microbes are busy in redox reactions (Sasowsky and Palmer, 1994). Chemolithotrophs obtain their energy from oxidation of inorganic materials, while chemoheterotrophs carry out oxidation of organic matter to obtain energy.
Microorganisms have the same mechanisms for weathering and mineralization inside caves irrespective of nature of the cave but some mechanisms and processes are limited to some specific caves. Some reactions require large amount of silica and thus these processes are restricted to volcanic caves. In some volcanic caves of Korea (Kashima et al., 1989) the formation of coralloids and helictates are mainly due to the presence of specific diatom Meolosira. The presence of Meolosira sp. is found only in twilight zones of caves. Onac et al. (2001) reported the role of microorganisms in speleothem deposition and precipitation of silica as Opal, opal-CT and quartz.
In addition, microbes have been found to enhance weathering of basaltic rocks into amorphous silica moon milk in lave tube caves Pico Island (Azores) and Rapa Nui Island. The organic richness of deposits indicates that microorganisms control
weathering processes (Forti, 2005; Calaforra et al., 2008). The caves of Pico Island have the largest opal flowstone which is formed from silica moon milk (Forti, 2001). Generally, the role of microorganisms in cave formation; weathering and mineral precipitation is an established fact. These are complex processes and involve microbial enzymes with a variety of mechanisms and reactions. These processes are complex and take a long period of time. A number of different reactions like dissolution, double displacement reaction, redox reactions are responsible for formation of phosphate, sulfate, nitrate and other minerals.
Precipitation of guano related minerals
It is found that all type of caves contain organic deposit of bat guano although traces of some other animals are also found. Guano is rich organic complex material host a number of different reactions mostly of biological origin which lead to the formation of different acids like nitric acid, phosphoric acid and sulfuric acids (Forti, 2001). These acids react with carbonate and other sediments which produce more than 100 different secondary minerals (Onac, 2005, 2011b). Some important compounds directly produce in guano are guanine and urea.
- In cave environment there are two sources of NO3 ions, one is the decaying forest material in humid zones and the second one is deposit of guano which contain N2.
The Process of Cave formation (Speleogenesis)
The process of cave formation is a conventional procedure of mineral dissolution by ground water. This process is dependent on climatic parameters; speleogenesis is mostly rapid in humid and warm climate (Priesnitz, 1974). The solubility is minerals specific. The most soluble rock type is limestone which is composed of calcite, aragonite, and dolomite. At higher concentration of CO2, the carbonates dissolve rapidly, the reactions are:
CO2 (g) ↔ CO2 (aq) -----(1)
CO2 (aq) + H2O ↔ H2CO3-----(2)
2+ - CaCO3 (s) + H2CO3(aq) ↔ Ca + 2HCO3 -----(3)
Equation (3) is a partial reaction, which combines with the equations (4) and (5) and cause calcite dissolution (Plummer et al., 1978). This process can also be summarized in one equation (6).
+ 2+ - CaCO3 (s) + H ↔ Ca + HCO3 -----(4)
2+ - - CaCO3 (s) + H2O ↔ Ca + HCO3 + OH -----(5)
2+ - CaCO3 (s) + CO2 (aq) + H2O ↔ Ca + 2HCO3 -----(6)
According to equation 4 and 6, the pH and CO2 concentration in water are the two key factors influencing the kinetics of limestone dissolution. These factors are at equilibrium at the atmosphere in systems, CO2 concentration depends on pCO2 (sea level value is ca. 39Pa, which is corresponding to the 0.039% concentration) and temperature (solubilization of CO2 is lower at increase temperature). Thus for cave development on bare rocks free of soil and vegetation, temperature climate would provide ideal environment. While in case of covered rocks, biological processes in soil are the vital source of CO2 instead of atmosphere. In this situation temperature, humidity and primary production levels are the factors which determine the concentration of soil CO2 and fluxes (Lloyd and Taylor, 1994; Raich and Schlesinger,
1992; Rustad and Fernandez, 1998). The CO2 proportion in soil gases are normally range from 0.2-11% but up to 17% is also reported in tropical regions (Derbyshire, 1976; Hashimoto et al., 2004; Köhler, 2009; Kursar, 1989; Liu et al., 2010). While in moderate climate the average range is from 0.1-3.5% but values up to 10.2% are also reported occasionally (Bekele et al., 2007; Davidson et al., 2007; Derbyshire, 1976;
Jassal et al., 2005). Thus, microbial and plant respiration is capable to raise the CO2 concentration more than hundred time as compared to atmospheric conditions and to speed up the speleogenesis (Gabrovsek et al., 2000). Acid production by microbes also accelerates the minerals solubility.
The above discussed conventional speleogenesis mostly depends on water and carbonic acid dissolution in the very common type of rocks. Along limestone rocks other forms of karst are also exist. In barokarst speleogenesis the process is enhanced by pressure corrosion, while hot springs and steam play a key role in thermokarst formation. Another specific type of limestone rock is hypokarst where the cave formation starts
from inside of bedrock, this type of speleogenesis is happen at the edge of oxic and anoxic zones in the presence of high value of H2S in ground water. This type of speleogenesis is called Sulfuric Acid Speleogenesis (SAS) and was first proposed by
Egemeier in 1970’s (Egemeier, 1981). In this mechanism the H2S is evaporate into the cave environment and oxidized to sulfuric acid (H2SO4) on the surfaces of wet bedrock which cause the renewal of carbonate by gypsum, which can be easily dissolve in water that considerable raise the void volume (Hill, 1990; Palmer, 1991).
H2SO4 + CaCO3 + H2O→CaSO4.2H2O + CO2
Different famous caves are formed by this mechanism in world e.g., Lechuguilla (USA), Frasassi (Italy), Novaya Afonskaya (Georgia). Light sulphur isotopes have been isolated from gypsum in above studied sites which hint the involvement of microorganisms in such processes. While the role of microbes in this type of speleogenesis is still under debate, Engle and his colleagues have reported that sulfur oxidizing microbes having affiliation with β-, γ-, and ε-Proteobacteria may play role in
H2S oxidation and carbonates dissolution than abiotic processes under certain conditions (Engel et al., 2004). Microbes use H2S as an energy source. For example, in sulfidic caves, the microbes oxidize H2S and form sulfuric acid, which further react with carbonate and causes dissolution of rocks (Engel, et al., 2004, Macalady, et al., 2007).
Factors Affecting Speleogenesis
There are at least eight different factors which are involved in solubility of calcite, dolomite and rate at which solution occurs, which are: CO2 concentration in solution, pH, organic matters oxidation, temperature, pressure, concentration of added salts, rate of solution flow, and degree of solution mixing. Calcium carbonate is more soluble if
CO2 concentration is raised, pH is decreased, O2 and organic compounds are increased, temperature is lowered, pressure is raised, salts concentration is increased, rate of flow is increased and increased the degree of mixing (Thrailkill, 1968).
Concentration of CO2 is a single key factor which affect the solution because CO2 react with water to form carbonic acid (H2CO3). The air normally has a pressure of
1atmosphere, and has a partial pressure of 0.0003 atm of CO2. Rain water dissolve
small amount of calcite during rain. Water having oxygen and organic compounds and possess 0.1 atm of carbon dioxide are capable to dissolve a lot of calcite (Moore, 1981).
Microbial calcium carbonates precipitation (MCP)
The phenomena of calcium carbonates precipitation have been studied from last two decades (Barton and Northup, 2007; Castanier et al., 1999; Forti, 2002). Microbially induced calcium carbonate precipitation is an environmental friendly process to protect decayed ornamental stones. This process relies on microbially induced carbonate precipitation of limestone. Unlike that from lime-water treatment, the carbonate cement formed under bacterial influence appears to be highly coherent (Metayer-Levrel et al., 1999). This method has been used for the durability of cementitious materials (Ramachandran et al., 2001; Muynck et al., 2008). Cave environment is favorable for the chemical precipitation (Castanier et al., 1999; Ehrlich, 1998) while the role of microbes in this process is nevertheless discussed. The carbonates precipitation is biologically induced and controlled mechanism which is reported from caves environment (Barton et al., 2001; Boquet et al., 1973; Cañaveras et al., 2001; Douglas and Beveridge, 1998). Different factors are involved in Carbonates precipitation by bacteria e.g., elimination of mineralization inhibitors by bacteria (Bosak and Newman, 2005), changing in microenvironment pH via autotrophic process, or by fixation or release of nitrogen (Cacchio et al., 2004; Castanier et al., 1999; Hammes and Verstraete, 2002). Some scientists believe that calcium carbonate precipitation by microbes is active pumping of calcium ions (Ca+2) from inside of cell into outside medium, to avoid the toxicity of calcium concentrations (Cacchio et al., 2004; Cai and Lytton, 2004). A mutant strain from which a ChaA gene was knocked out, lost the expression of Ca+2 ion efflux protein and also stopped growth on a medium having carbonate, as opposed to the wild type (Banks et al., 2010). These authors proposed that bacterial cell play a key role in initial crystal nucleation, which was further confirmed by Sanchez-Moral et al (2012), who reported that bacterial cell would promote initial steps of deposition. Rusznyak et al. (2012) reported some microbes having the capability of specific crystal formation in harvesting experiments were isolated from speleothems of Herrenberg Cave (Germany). The carbonate accumulation was described to be less favorable for the growth of microbes (Sanchez-Moral et al., 2012).
Microbial cells excrete polysaccharides and wide range of amino acids outside which also may influence the formation of crystals, even placed in abiotic carbonate environment (Braissant et al., 2003). Altogether, for the initiation of carbonate crystal formation, their shapes, and precipitation, microorganism play a vital role.
Mechanisms of microbial calcium carbonate precipitation
The carbonate precipitation is biologically induced and controlled mechanism which is reported from caves environment (Barton et al., 2001). Microbial carbonate precipitation (MCP) occurs as a byproduct of common microbial metabolic processes, such as: 1. Photosynthesis (McConnaughey and Whelan, 1997), 2. Urea hydrolysis (Stocks-Fischer et al., 1999) and 3. Sulfate reduction (Castanier et al., 1999). Bacterial cells have negative surface charge, which act as a scavenger of cations, containing Ca+2 and Mg+2 bind on to cell surfaces, which making microbes as an ideal nucleation sites for crystals e.g., MCP (Stocks-Fischer et al., 1999; Ercole et al., 2007). Another advantage of MCP is its capability of sequestration of atmospheric carbon dioxide through CaCO3 formation (Barath et al., 2003). Bio-precipitation of calcium carbonate technologies have many importance e.g. used in sand columns consolidation (Nemati and Voordouw, 2003), for repair of limestone monuments (Fujita et al., 2000) and to a smaller extent for remediation of cracks in concrete (Ramachandran et al., 2001).
Fig. 2.1. The process of calcium carbonate precipitation by bacteria (Braissant et al. 2009)
The second mechanism by which bacteria precipitate the calcium carbonate is microbially induced calcium carbonate precipitation (MICCP). Ureolytic bacteria hydrolyze urea and form wide rang of carbonates in short period of time. In urease controlled reaction, 1 mol of urea is hydrolyzed inside microbial cell and form 1 mol of ammonia and 1 mol of carbonate, which hydrolyzed spontaneously to form additional 1 mol of ammonia and carbonic acid:
CO(NH2)2 + H2O NH2COOH + NH3
NH2COOH + H2O NH3 + H2CO3
These products equilibrate in water to form bicarbonate, 1 mol of ammonium and hydroxide ions which give rise to pH increase
+ -2 H2CO3 2H + 2CO3 +4 - NH3 + H2O NH + OH +2 -2 Ca + CO3 CaCO3
Bacterial communities have the ability to change the pH of microenvironment to
alkaline via physiochemical reactions. Calcium ions are also concentrated by bacterial cell surfaces in precipitation (Fortin et al., 1997). At neutral pH, positively charged particles could make a layer on bacterial cells surfaces due to its negative charge which favor the heterogenous nucleation (Douglas and Beveridge, 1998; Bäuerlein, 2003). Calcium carbonates precipitates commonly develop on the outside surfaces of bacterial cells by successive stratification (Pentecost and Bauld, 1988; Castanier et al., 1999) and bacterial cells can be embedded in developing crystals (Rivadeneyra et al., 1998; Castanier et al., 1999).
Fig. 2.2. Bacteria serving as nucleation site for CaCO3 precipitation in the sand particles (Source: DeJong et al., 2010).
Possible reactions in urea-CaCl2 medium to precipitate CaCO3 at the cell surface can be summarized as follows:
Ca2+ + Cell → Cell−Ca2+
− 3− 2− Cl + HCO + NH3 → NH4Cl + CO3
2+ 2− Cell−Ca + CO3 → Cell−CaCO3
Manganese oxidation by bacteria
Manganese oxidation by cavernicoles
The Earth’s crust is composed of about 0.1% manganese (Nealson, 1983). In Earth’s crust manganese secured fifth position in transition metals (Tebo et al., 2004). Manganese is present in 7 different oxidation states extending from 0 to +7 while naturally it is present in +II, +III and +IV states (Tebo et al. 1997, 2004). Mn have higher redox potential than iron due to which the reduction of Mn is easier than Fe, and tough to oxidize than Fe (Kirchner and Grabowski, 1972).
Oxyhydroxides are found in abundant after sulfate and carbonate minerals. Many reports are available which showed the presence of iron and manganese in abundant form in caves (Hill and Fort, 1997; White et al., 2009; Gazquez et al., 2011) to irregular surfaces on the walls usually on top of visibly altered carbonates (Northup et al., 2003;
Spilde et al., 2005; Gazquez et al., 2012a). With carbonate and silicate speleothems, sulfur compounds, oxides of iron and manganese and saltpeter, the microbial associations have been reported (Northup et al. 1997). The cavernicoles play important role in mineral precipitation either actively by producing enzymes or other metabolites which change the microenvironment (Danielli and Edington. 1983), or passively by acting as nucleation site (Went, 1969). Cave microbes also play an important role in cave dissolution by producing the acid as a byproduct (Ehrlich, 1996). Speleothems and cave microbes have many interactions due to which Cunningham et al. (1995) called speleothems a biothems. In speleothem formation the microbes play a key role.
Manganese compounds are present in caves as in clastic deposit form layer on wall or speleothems (Gascoin. 1982; Hill. 1982) or as crust (Jones. 1992; Moore. 1981). In cave, manganese is present in the form of birnessite very common (Hill and Forti, 1997), and some low quantity of crystals oxides and hydroxides like pyrolusite, chalcophanite, cryptomelane, hausmannite, romanechite, rancieite, todorokite and rhodochrosite are also reported (Onac et al.. 1997a; Onac et al.. 1997b). From karst solution a cavity, the manganese is also isolated (Jones, 1992). Peck (1986) and Northup et al. (2003) provided evidences that manganese oxides reported from caves are almost biogenic in nature. Mn(IV) oxide are present in aquatic as well as in terrestrial environment (Post, 1999). Mn boosts the enzymatic activity like DNA polymerase and phosphoenol pyruvate carboxykinase that is why it is present in a minute quantity in all living organisms and is an important element for all living kingdoms (Beyer et al., 1986). Manganese is a strong redox agent and plays an important part in aerobic biological redox reactions e.g. Mn superoxide dismutase, and Mn pseudocatalase (Beyer and Fridovich. 1987; Dubinina. 1978). Mn (II) is the soluble form of manganese or form a complex compound with organic or inorganic ligands, while Mn(III) is unstable in aquatic condition and readily change to Mn(II) and MnO2 and Mn(IV) forms insoluble oxides and oxyhydroxides. Oxidation of Fe and Mn from reduced stats in aqous and oxic forms produce a H+ which leads to acidification of the environment.
+2 + 2Mn +O2(g)+2H2O → 2MnO2(s)+4H
+2 + 4Fe +O2(g)+10H2O → 4Fe(OH)3(s)+8H
Microorganisms of caves are capable of production extracellular polymeric substances (EPSs) and some other metabolites with acidic functional groups, which stress the pH lowering due to metal oxidation. Due to deprotonation of organic functional group, the cell walls of bacteria have negative charges at low pH which may act as nucleating sites for the cation like iron and manganese (Fein, 2009).
Microbes have a key role in the formation of insoluble Mn(III, IV) from the soluble Mn(II) in natural environments (Tebo et al., 1997). The microbial processes speed up the oxidation of Mn(II) to Mn(IV) five times faster than surfaced catalyzed reactions (Nealson et al., 1988; Tebo et al., 2004), and the presence of Manganese oxides in soil, sediments and aquatic environment are thought to be of biological origin (Nealson et al., 1988; Tebo, 1991; Wehrli et al., 1995).
Microbial diversity of cave
Caves are natural underground cavities large enough for a human being to enter, mostly in empty spaces in rocks. Caves are considered as an extreme environment, which is unsuitable for the life due to extreme abiotic conditions. They form ecological niches for specific group of microbes (Schabereiter-Gurtner et al., 2004). Most caves in the world are limestone caves formed from limestone rocks, while lava tube caves are in basalt rock. An oligotrophic ecosystem is present inside caves. Caves are characterized by complete darkness, low temperature, and high humidity almost 100%. Due to oligotrophic conditions the average number of microbes growing is 106 cells/g of rock (Barton and Jurado, 2007).
Microbes in any environment are dependent on energy and carbon fixation of photosynthetic organisms, but in case of cave environment where darkness persists throughout, prevents the phototrophs colonization. Restricted nutrients and energy can enter into caves through entrance, sinkhole, underground hydrology, and drip waters (Barton and Jurado, 2007) and the aphotic and oligotrophic environments only allow for the survival and functioning of species adapted to the oligotrophic conditions, which
clarify the dominancy of chemoautotrophic microbes in cave environments which fixes carbon and import energy into cave food web (Sarbu et al., 1996; Chen et al., 2009).
Photosynthetic activity occurs only at the entrance of the caves, and sometimes inside the caves due to artificial light. Perpetual light absence prevents the production of primary organic compounds by phototrophic microbes. Cavernicoles use alternate method for the assimilation of carbon associated with chemoautotrophy. In such dark condition, the primary organic compound is produced by chemolithoautotrophic microbes, which not only derive energy by binding with hydrogen, nitrogen, or volatile organic matters but also from oxidation of reduced metal ions (e.g manganese, iron) available in cave rocks (Gadd, 2010; Northup and Lavoie, 2001). The only source of allochthonous compound in cave is leaking of water via cracks in rocks from soil present above it, streams depositing sediments of clay on the walls and floor, or through air which carries organic compound particles. Besides these, the other sources of organic matters are remains of human and animal activity or plant roots. The presence of these organic compounds enhances the growth of heterotrophic organisms.
Bacteria and archaea are the dominant domains of cave environment and are present in different cave habitats such as sediments, rock surfaces, soils and stream waters (Engel et al., 2004; Barton and Jurado, 2007). 16S rRNA gene sequencing have been helpful to identify many bacterial phyla in caves (Engel et al., 2004; Barton et al., 2007; Ortiz et al., 2013b). According to 16S rRNA gene sequencing, the Proteobacteria, Acidobacteria and Actinobacteria are the dominant bacterial taxa on cave walls (Barton et al., 2007; Pašic ́ et al., 2010; Cuezva et al., 2012). Bacterial colonization in sediments of cave could be comparable to that in overlying soils (Ortiz et al., 2013b), but the surfaces of cave rocks are colonized by lowest diversity natural microbial communities (Macalady et al., 2007; Yang et al., 2011). From comparative study of geographical distinct caves, it was supposed that surfaces of cave rocks could be colonized by common microbial phyla which are merely found in other habitats, which suggest a specific bacterial linage in cave environments (Porca et al., 2012).
In past, traditional culturing techniques were used for the study of microbial diversity. By using such traditional methods, microbial diversity was not found convincingly due to the inability to grow the microorganisms (Torsvik and Ovreas, 2002). This problem was overcome by molecular techniques. This is plausibly due to the fact that microbes
that grow in vitro generally comprise a small portion of environmental populations (Donachie et al., 2007). Nowadays, the most common and accurate methods for the determining microbial diversity are based on molecular markers, including small (16S rRNA) and large (23S rRNA) ribosomal RNA genes subunits, as well as functional genes such as soxB (active in sulfur oxidizing microbes), amoA (active in ammonia oxidizing microorganisms), RuBisCO (gene found in chemoautotrophic organisms) and genes critical for cell function, i.e., “housekeeping genes” like rpoB, recA or gyrB (Holmes et al., 2004).
Fig. 2.3. Distribution of major groups of microbial communities in cave environments by 16S rRNA gene sequencing (Pasic et al., 2010)
Table. 2.1. Different caves are studied for microbial diversity in different countries
Caves Country Microbes Proponents/Year Nullabor Caves Australia α, β, γ and δ-Proteobacteria and Holmes et al., 2001 novel microorganisms Altamira Cave Spain Proteobacteria, Plantomycetales, Schabereiter-Gurtner Cytophagal/Flexibacter/Bacteroides, et al., 2002 Cuezva et Acidobacterium, Actinobacteria and al., 2009 green-sulfur bacteria Reed Flute Guilin Guangxi Knoellia sinensis and K. Groth et al., 2002 China subterranea (Actinobacteria) Lower Kane Wyoming, USA ε-Proteobacteria Engle et al., 2002 Cave Lechuguilla Carlsbad Caverns α, β and γ-Proteobacteria, Northup et al., 2003 Spider National Park New Enterobacteriaceae, Xanthomonas, Mexico, USA Bacillus/Coliform group and Lactobacillaceae
Llonin La Asturias, Northern Proteobacteria, Actinobacteria, Schabereiter-Gurtner Garma Spain et al., 2004 Gram-positive bacteria Kartchner Arizona, USA Bacillus, Brevibacillus, Rhizobium, Ikner et al., 2006 Caverns Sphingomonas, Staphylococcus, Pseudomonas and other uncultured β-Proteobacteria
Applications of cave microorganisms
Recognition of microbes in caves changed our perception about life in cave (Barton and Northup, 2006). Biospeleology has similarly mirrored the meteoric rise of microbiology as science, with new insights suggesting that cave microbes may be play a role in process as varied as speleothem deposition to cavern development (Canaveras et al., 2006; Engel et al., 2004). Microbes have wide range of activities in cave environment from obvious slimy goop to the more subtle calcite deposition or changing in the structure of rocks surfaces. The importance of cave microbes may be of interest to speleologists, the applications of this study go well beyond caves. The cave microbes degrade the ancient, prehistoric paintings within cave environment (Schabereiter- Gurtner et al., 2002). Besides this contribution of cavernicoles in degradation and conservation of these paintings, but these microbes also have a potential role in major biogeochemical processes occurring inside cave environment.
Biocementation
Cave microbes have great role in biocementation processes occurring in cave environment. These microbes colonize on carbonate surfaces which lead to deposition of calcite. The work had a key role in ancient marble monuments and statues preservation, where microbes could deposit a coating of calcite to prevent ancient structures from erosion (Laiz et al., 2003).
Mineral precipitation
In caves, energy is produced by different microbial processes which are used for their survival. Although these are small reactions which produce nitrates, sulfate, and carbonate, but play a key role in cave ecosystem. These microbes also play an important role in minerals precipitation from which the interaction between autotrophic and heterotrophic can be studied. Heterotrophic microbes are present at the entrance gate of cave while autotrophic are at the depth of cave, for the growth of autotrophic microorganisms’ different reactions occur like volcanism, serpentanization and radiolysis.
These microbes not only precipitate calcite but also oxidize soluble manganese (Mn+2) to Mn(III, IV) which are the insoluble or precipitated form of manganese. Biogenic Mn oxides are strongest naturally found oxidants and play a key role in oxidation/reduction of many organic and inorganic compounds. Biogenic Mn oxides have high sorptive ability due to which adsorb many ions, controlling the bioavailability of many toxic and essential elements. Biogenic Mn oxides recognized as “scavengers of the sea” (Goldberg, 1954). The biogenic Mn oxides are also used in waste water treatment to oxidize or reduce toxic metals.
Biodegradation
Cave microbes can degrade the complex aromatic compounds, such as benzothiazole and benezenesulfonic acid for growth which are used in the manufacture of plastic and are environmentally dangerous contaminants (Bennett and Barton, 2006). Due to this talent of cavernicoles, these microbes are inoculated into contaminated environments to degrade and clean pollutants rapidly and to restore the natural habitats in a process known bioremediation.
Production of enzymes and novel antibiotics
Cavernicoles are adapted to the extreme starvation condition because of this ability these microbes also have the ability of to harbor other essential biomolecules. Similarly cave environment have microorganisms which produce ethanol for fuel, extremozyme which are environmentally friendly and are used in paper procession and even use in stonewashing of jeans. These microbes also have the ability to produce novel antibiotics and anticancer metabolites (Onaga, 2001). Cave microbes are possible sources of many useful compounds: biosurfactants (Banat et al, 2010), antitumors (Chang et al, 2011), exopolysaccharides (Nicolaus et al, 2010), radiation protective drugs (Singh and Gabani, 2011). Scientists are doing more and advance research to discover novel antimicrobial compounds from a new site of microbes. The innovation gap over the past 30 years, however, two new antibiotics has been introduced: oxazolidinone linezolid in 2000 and cyclic lipopeptide daptomycin in 2003 (Hamad, 2010). Microbes are present in soil which plays a key role in regulation of microbial communities in soil, water, sewage and compost. There are hundreds of antibiotics are produced naturally
in a pure form, among these a few are non-toxic and are used in daily life (Zinsser, 1988). Biomarkers for extraterrestrial life
Recently NASA once again focusing on exploration of past life and life forms on Moon, Mars and other extra-terrestrial habitats by simulation models of caves environment and geochemistry and biology (NASA report, 2014). Finally, one of the philosophical questions of humanity regards our place in the Universe: Are we alone? Is life on Earth unique? Cave microbiology can not only answer questions about the limits of life, but also help us to identify the geochemical signatures of life. Such signatures are capable of surviving geologic uplift, which allows them to be detected on the surface of planets, such as Mars (Boston et al., 2001). While such ideas may seem an extraordinary application of cave geomicrobiology, NASA has recently undergone a dramatic refocus by gearing its activities to returning humans to the moon and exploration of Mars and world’s beyond to find evidence of past life, activities in which cave geomicrobiology may play an important role (White House Press Release, 2004).
Geochemical and mineralogical analysis of Kashmir cave (smast), Buner, Pakistan, and isolation and characterization of bacteria having antibacterial activity Sahib Zada1†, Abbas Ali Naseem2†, Seong-Joo Lee2, Muhammad Rafiq1,Imran Khan1, Aamer Ali Shah1, Fariha Hasan1* 1Department of Microbiology, Quaid-i-Azam University, Islamabad, Pakistan 2Department of Geology, Kyungpook National University, Daegu 702 – 701, Korea
ABSTRACT Bacterial strains having the ability to inhibit the growth of other bacteria, were isolated from soil sample collected from Kashmir smast (‘smast’ is Pushto language word means cave), Khyber Pakhtunkhwa, Pakistan. The study includes mineralogical and geochemical analyses of soil sample collected from the cave, so as to describe the habitat from where the microorganisms have been isolated from. Total bacterial count of the soil sample was 5.25×104 CFU/ml. Four bacterial isolates having activity against test organisms (Micrococcus luteus, Klebsiella sp., Pseudomonas sp. and Staphylococcus aureus) were screened out for further study. Two of the isolates were found to be Gram positive and the other two Gram negatives. The four isolates showing antibacterial activity were identified as Serratia sp. KC1-MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3-MRL and Stenotrophomonas sp. KC4-MRL on the basis of 16S rRNA sequence analysis. Although, all isolates showed antibacterial activity but only Bacillus licheniformis KC2-MRL was selected for further study on the basis of size of zone of inhibition. Antibacterial activity of the B. licheniformis KC2-MRL was optimum when grown in Nutrient broth adjusted to pH 5 and after 24 hours of incubation at 35oC. The extracted antibacterial compound was stable at pH 5-7 and 40oC when incubated for 1 hour. The strain was found resistant against cefotaxime (ctx). Atomic absorption analysis of the soil sample collected from the cave showed high concentration of calcium (332.938 mg/kg) and magnesium (1.2576 mg/kg) as compared to the control soil collected from outside the cave. FTIR spectrum of concentrated protein showed similarity to that of bacitracin. Interestingly, the antibacterial compound showed activity against both Gram negative and positive test strains. Mineralogy of Kashmir smast (cave) is diverse and noteworthy. Different
geochemical classes identified by X-ray diffraction were as nitrates, oxides, phosphates, silicates and sulfates. Weathered cave limestone contribute notably to the formation of these minerals or compounds. FTIR spectroscopic analysis helped to identify minerals such as quartz, clinochlore, vermiculite, Illite, calcite and biotite.
Key words: Bacillus licheniformis KC2-MRL, Kashmir smast, mineralogy, antibacterial activity
INTRODUCTION
Caves are characterized as having very low nutrient availability constant low temperatures, and high humidity. Caves can either be terrestrial or aquatic, usually oligotrophic innature i.e. nutrient limited. Some may be rich in specific natural minerals or due to exposure to nutrient containing sources, therefore, different caves will have different types of microorganisms inhabiting various ecological niches. Fauna, environmental factors, temperature and organic matter, describes the caves’ biotic activities such as nutrient cycling and geomicrobiological activities including formation/alteration of cave structures (Adetutu and Ball, 2014). Cave organisms have evolved some extraordinary abilities to survive and live in this inhospitable environment (Engel et al., 2005; Simmons et al., 2008; Northup and Lavoie, 2009). Cave microbial flora is rich in different types of microorganisms having some diverse and unique characteristics (Groth et al., 1999). The most abundant organisms observed in caves are filamentous and belong to the Actinobacteria group, followed by coccoid and bacilli forms (Cuezva et al., 2009). Some pathogenic microorganisms have also been reported from the Altamira cave (Jurado et al., 2006). Luong et al. (2010), for the first time, reported the recovery of Aurantimonas altamirensis from human medical constituents, other than from cave. E. coli and S. aureus, the disease causing bacteria, have also been isolated from caves (Lavoie and Northup, 2005) and species of Pseudomonas, Sphingomonas and Alcaligenes sp. (Ikner et al., 2007) and Inquilinus sp. (Laiz et al., 1999). Caves can be source of novel microorganisms and biomolecules such as enzymes and antibiotics that may be suitable for biotechnological purposes (Tomova et al., 2013). The influence of particular nutrients in antibiotic biosynthesis, was determined by the
chemical structures of antibiotic substances (Pereda et al., 1998). Rigali et al., (2008), provide evidence that certain substrates and oligotrophic conditions will lead to increased induction of secondary metabolites. Nitrogen from various sources may incorporate in antibiotic molecules as precursors or their amino groups can transfer to specific intermediate products (Doull and Vining, 1990; Cheng et al., 1993). Nutrient deficiency is responsible for onset of antibiotic biosynthesis (Demain et al., 1983; Doull and Vining, 1990; Sanchez and Demain, 2002). When carbon or nitrogen source is a limiting factor, growth is rapidly reduced and antibiotic biosynthesis occurs in the stationary phase. In other cases, antibiotic production is associated with the growth phase. Due to the oligotrophic environment in cave ecosystems, microorganisms present in the cave compete for nutrients and produce antibiotics against other microbes. Microbial resistance systems against wide spectrum, standard antibiotics, metabolic by-products such as organic acids, and lytic agents such as lysozyme. Besides these antibiotics, other biologically active compounds like exotoxins and bacteriocins were described be Riley and Wertz (2002) and Yeaman and Yount (2003). The continuous efforts of scientists are to discover new antibiotics and new source microorganisms. Cave microorganisms can be used for the production of potential new antibiotics. Antibiotic producing microbes mostly belong to the genera Penicillium, Streptomyces, Cephalosporium, Micromonospora, Bacillus (Park et al., 1998) and Pseudomonas species followed by the entero, lactobacilli and streptococci (Berdy, 2005). More than 8000 antibiotics are known to exist, and hundreds are discovered yearly (Brock and Madigan, 1991), however only a few prove to be commercially useful. About 17% of these antibiotics are produced by molds and 74% by Actinomycetes (Zhang et al., 2008). Bacillus sp. mostly form peptides and phenazines, which are heterocyclic and derivatives of fatty acid but the production of macrolactones is very rare (Berdy, 2005). Gramicidins, polymixins, bacitracins and some other antibiotics are formed non- ribosomally (Nissen-Meyer and Nes, 1997; Hancock and Chapple, 1999). A great number of antibiotics have been isolated from various microorganisms. Studies are still being conducted to isolate and identify novel antibiotics effective against pathogenic fungi and bacteria. The number and species of microorganisms in soil vary in response to environmental conditions such as nutrient availability, soil texture, and type of vegetation cover (Atlas and Bartha, 1998). The soil composition and texture play important role in harboring microbes with unique characteristics. Thus it is
important to know about the composition, soil type, structure and texture of the soil from where the microorganisms are isolated for research purpose and for the production of metabolites, such as antibiotics. Microbial species adapt to caves by interacting with minerals there (Barton and Jurado, 2007). The Geochemistry and metal analysis of the cave environment can influence the synthesis of antibiotics by cave bacteria, as metal ions are known to affect the synthesis of microbial metabolites in vitro. Tanaka, et al., (2010) made a connection between rare earth elements, scandium and/or lanthanum, and increased activation of the expression of nine genes belonging to nine secondary metabolite–biosynthetic gene clusters of Streptomyces coelicolor A3(2). Investigations on the effect of several metal ions indicated that Cu2+, Mn2+ and Fe2+ stimulated AK-111-81 biosynthesis by Streptomyces hygroscopicus, depending on their concentration (Gesheva et al., 2005). Divalent ions stimulated the production of polyenes (Georgieva-Borisova, 1974; Liu et al., 1975; Solivery et al., 1988; Park et al., 1998) and Fe2+ and Mn2+have been found to favour niphimycin production. Soil texture and structure also strongly influence the activity of soil biota. For example, medium-textured loam and clay soils enhance activity of microbes and earthworms, whereas fine textured sandy soils, with lower water retention potentials, are not very favorable. Alterations in pH of the soil can affect metabolism of species, enzyme activity and availability of nutrients, and thus are often lethal (Singh and Mishra, 2013). The aim of the present study was to isolate microbes from the cave having antibacterial activity and characterization of the producer as well as the product and geochemistry of the cave, in order to understand the environmental conditions under which these microorganisms are living and producing compounds inhibitory for other microbes.
MATERIALS AND METHODS Sampling site and collection of soil samples Soil samples were collected from Kashmir smast (cave) (‘smast’ in local language means cave), Nanser, Buner, Khyber Pakhtunkhwa (GPS coordinates 34o25’42.12”N 72o13’10.82”E) (Fig. 3.1.1). The cave is 188 m long, with average height and width ~28 m and ~25 m, respectively. The Kashmir smast is a series of natural limestone caves (probably of marine origin), most part is of stalagmite, located in the Babozai Mountains in between Mardan and Buner in Northern Pakistan. According to study on a rare series of bronze coins and artifacts found in the region, the caves and their
adjacent valley probably comprised a sovereign kingdom in Gandhara which maintained at least partial independence for almost 500 years, from 4th century AD to the 9th century AD (Ziad, 2006). It is a limestone cave, with internal temperature around 10oC, having pH of 5 and the internal surface of cave was muddy due to dripping of water from the top. The only source of water was drip water. Two soil samples were collected from wall and ground surface of the cave in sterile Falcon tube under aseptic conditions. The sample was collected from the dark end of the cave about 188 m from entrance. This cave is located far away from human access so human intervention is negligible. The samples were then brought to the laboratory in an ice box and stored at 4°C for further processing. These soil samples were screened for the antibiotic producing isolates within 24 hours. Mineralogical Analysis Soil Analysis by Atomic Absorption For the quantitative analysis of elements (Ni, Cr, Co, Cu, Zn and Pb) in the soil sample, Atomic Absorption (AA240FS Fast Sequential Atomic Absorption Spectrophotometer) spectrophotometry was performed. To prepare the sample for this analysis, soil digestion was performed.
Fig. 3.1.1. Kashmir cave (smast), Nanseer Buner, Khyber Pakhtunkhwa, Pakistan. White arrows show location of the cave, black arrow shows entrance to the cave. (Pakistan full map from: http://www.mapsofworld.com/pakistan/; site map: google Earth)
Sample preparation for mineralogical analysis 1 g of cave and control soil (outside cave used as a control) each were ground separately, to make it more fine and were then mixed in 15 ml aqua regia, and was ° heated at 150 C and left overnight, then 5 ml of HClO4 was added and again heated at 150°C. The solution almost became dry until brown fumes were produced. Whatman filter paper (No. 42) was used for filtration and the volume was made up to 50 ml using double distilled water (FAO/SIDA, 1983). XRD (X-ray diffraction spectroscopy) X-ray powder diffraction (XRD) is a rapid analytical technique used for phase identification and characterization of unknown crystalline materials (e.g. minerals, inorganic compounds) and identification of fine-grained minerals such as clays and mixed layer clays that are difficult to determine optically (Geochemical Instrumentation and Analysis). XRD patterns were obtained from the samples using (X’Pert-APD Philips, The Netherlands) with an X-ray generator (3 kW) and anode (LFF Cu). The Cu Kα radiation was administered at a wavelength of 1.54 Å. The X-ray generator tension and current were 40 kV and 30 mA, respectively. The step-scan data were continuously collected over the range of 5 to 80°2θ. Quantitative Analysis Mineral proportions were calculated using SIROQUANT, a commercially available MS-Windows program for standardless mineral quantification. Weight percent mineral phase contents were estimated. Using calculated hkl mineral library files, refinement stages were optimized for the smallest possible χ2 goodness-of-fit parameter for the associated Rietveld peak pattern match (Taylor, 1991; Taylor and Clapp, 1993). Thermogravimetirc Analysis (TGA) Thermogravimetric (TGA) analysis determines weight loss due to heating, cooling, records change in mass from dehydration, decomposition, and oxidation of a sample with time and temperature (Voitovich et al, 1994). TGA was performed on Setaram TGA. Instruments incorporated high-resolution thermogravimetric analyser (series Q500) in a flowing nitrogen atmosphere (60 cm3 min–1). Approximately, 35 mg of sample underwent thermal analysis, with a heating rate of 5°C/min, within the range of from 25 to 1000°C. With the isothermal, isobaric heating program of the instrument the furnace temperature was regulated precisely to provide a uniform rate of decomposition in the main decomposition stage.
FE-SEM and EDS (Field Emission-Scanning electron microscopy with Energy- dispersive X-ray spectroscopy analysis) Field-emission cathode in the electron gun of a scanning electron microscope provides narrower probing beams at low as well as high electron energy, that results in improved spatial resolution and minimized sample charging and damage (Stranks et al., 1970). FE- SEM with EDS analysis of the samples were performed for the determination of thickness, structure uniformity and elemental composition measurement, using S-4800 and EDX-350 (Horiba) FE-SEM (Hitachi, Tokyo, Japan). Samples were spread on a glass plate that was fixed onto a brass holder, and coated with osmium tetraoxide
(OsO4) using a VD HPC-ISW osmium coater (Tokyo, Japan) prior to FE-SEM analysis. Fourier Transform Infrared Spectroscopy (FTIR) About 2 mg of the soil sample was mixed with 40 mg of KBr in ratio 1: 20 using mortar and pestle. KBr powder was dried at 120oC in an oven to avoid the broad spectral peak. A 1X 13 mm pellet was prepared. The pellet was placed in a holder and introduced in the infrared beam for analysis through Fourier Transform Infrared Spectrometer (Jasco FT/ IR – 620). Microbiological Studies Total viable heterotrophic bacteria (CFU/ml) For isolation of bacteria from the cave soil, 1 g of sample was serially diluted in normal saline and then was spread on Nutrient agar plates aseptically, and plates were incubated for 24 hrs, aerobically at 35°C. Viable cell count was calculated as CFU/ml. The isolate KC2-MRL was incubated at 25, 35 and 45°C and the growth (O.D at 600 nm). A growth curve was constructed by taking values of cell concentration on y-axis versus time along x-axis. Using a standard formula, growth rate and generation time was calculated from the graph. Screening and isolation of antibacterial compound producing bacteria Nutrient agar medium was used for isolation of antibiotic producing bacteria. Lawns of susceptible test organisms i.e. Micrococcus luteus (ATCC 10240), Klebsiella sp., Pseudomonas sp. and Staphylococcus aureus (ATCC 6538), were made on nutrient agar plates (Gauthier, 1976), which were then sprinkled with 20-25 particles of soil. All the plates were gently shaken so that the soil particles spread uniformly. Plates were then incubated at 35°C for 24 hours, lid side up, so that the soil particles do not fall off the agar. After 24 hours of incubation, plates were checked for antibacterial activity by
the formation of clear zone of inhibition around bacterial (KC2-MRL) colony. Zone producing isolates were purified and stored at 4°C. Morphological and biochemical characterization of bacterial isolates Colony morphology, Gram’s staining and biochemical tests (citrate utilization, oxidase and catalase production, nitrate and sulfate reduction, H2S production and carbohydrate fermentation) were performed according to Bergey’s Manual of Determinative Bacteriology (Holt et al., 2012). Molecular identification of the selected isolates The DNA extraction from bacteria was done by spinning 1 mL of culture at 10,000 rpm for about 3 min, the cells were pelleted out and rinsed twice in 400 µL TE buffer after removing the supernatant. Then the cells were centrifuged at 10,000 rpm for 3 min, the pellets were resuspended in 200 µL TE buffer. Then 100 µL Tris-saturated phenols of pH 8.0 were added to these tubes, followed by a vortex-mixing step of 60 sec, to lyse the cells. To remove the aqueous phase from organic phase, the samples were centrifuged at 13,000 rpm at 4°C for 5 minutes. Then 160 μL of upper aqueous phase was taken in a 1.5 mL Eppendorf. About 40 µL of TE buffer was added to make 200 µL and mixed with 100 µL of chloroform: isoamyl alcohol (24:1) and centrifuged for 5 min at 13,000 rpm at 4°C. Chloroform: isoamyl alcohol (24:1) extraction was used for the purification of lysate, when there was no longer a white interface, and the same method was repeated twice or thrice (Aitken, 2012). Purified DNA was present in the aqueous phase and was stored at -20°C for further use. The purified DNA was analyzed through agarose gel 1.5 g in 1x TBE, and staining with ethidium bromide. Phylogenetic analysis Phylogenetic analysis was performed through ClustalW program implemented in MEGA4.0 (Thompson et al., 1994). The similar sequences were downloaded from NCBI. All sequences were aligned and the phylogenetic tree was constructed using Neighbor Joining method in MEGA4.0 bootstrap analysis 1000 replicate, was performed for the significance of the generated tree.
Production of antibacterial compounds by B. licheniformis KC2-MRL Inoculum of B. licheniformis KC2-MRLwas selected after screening, on the basis of larger zone of inhibition against test strains,was prepared in nutrient broth. About 50 ml of nutrient broth was prepared in 250 ml flask and autoclaved and incubated at 35°C overnight to check the sterility and inoculated with the producer strain, and incubated
at 35°C for 24 hours in orbital shaker at 150 rpm. Sterilized nutrient broth (50 ml) was taken in 100 ml flasks and pH was adjusted to 5 (pH of sampling site was 5). Approximately, 10% inoculum was added in each flask and incubated at 35°C in orbital shaker at 150 rpm. After every 24 hrs, samples were collected (centrifuged at 10,000 rpm for 16 minutes), for a total of 96 hrs to obtain cell free supernatant (CFS). The CFS was checked for antibacterial activity by agar well diffusion assay. Agar well diffusion assay The production of antimicrobial metabolites by B. licheniformis KC2-MRL, was checked by agar well diffusion method (Sen et al., 1995). About 80 l of CFS was added in the wells and the plates were incubated at 35°C for 24 hours. After 24 hrs, the zones of inhibition were observed and the diameter of the zone of inhibition (mm) was measured. Effect of medium, pH, temperature and incubation on the antibacterial activity Different media used for the production of antibacterial compounds by B. licheniformis KC2-MRL, including Trypticase soya broth (TSB), nutrient broth (NB) and Luria Bertani (LB) broth. Inoculum (10%) was added and incubated at 37°C and at 150 rpm. The cell growth was measured by optical density at 600 nm and antimicrobial activity was checked by agar well diffusion assay. To check the effect of time of incubation on the antimicrobial activity, the strain was incubated at 37°C in orbital shaker at 150 rpm and samples were drawn after every 24 hours from 0 to 96 hours. The antimicrobial activity of all the collected CFS was checked against S. aureus, M. luteus, Klebsiella sp. and E. coli. Effect of temperature (15, 25, 35 and 45ºC) on optimum antibacterial activity was studied by inoculating B. licheniformis KC2-MRL in Nutrient broth and incubated at 15, 25, 35 and 45°C at 150 rpm. Samples were drawn 24 hourly from 0- 96 hrs. Centrifuged and CFS were used for further analysis using S. aureus, M. luteus, Klebsiella sp. and Pseudomonas sp. as test strains. Effect of pH (5, 6, 7 and 8) on the production of antibiotics was studied by inoculating B. licheniformis KC2-MRL in the growth medium having different pH. Samples were drawn from 0 to 96 hours after every 24 hours, centrifuged and cell free supernatants were used for further analysis. Antibiotic susceptibility test
Antibiotic susceptibility test was performed to check the sensitivity of the selected strain against various broad spectrum antibiotics, to check for the intrinsic ability of microorganisms to resist antibiotics. Partial purification and FTIR of the antibacterial compound Cell free supernatant of B. licheniformis KC2-MRL culture grown under optimized conditions was used for the precipitation of antibacterial compounds using increasing concentrations (10- 80%) of ammonium sulfate. The pellet was kept at - 20°C in 10 ml of 0.1M phosphate buffer, pH 7. FTIR was performed to identify unknown compounds. Spectrum of the antibacterial compound, produced by Bacillus licheniformis KC2-MRL was compared with that of bacitracin as a control. Samples were scanned from 4000-400 cm-1 at resolution of 6.0 cm-1.
RESULTS Mineralogical Analysis Experimental X-ray pattern of smast-5 and smast-7 along with the ICSD (Inorganic Crystal Structure Database) reference code data of different crystals is shown in Fig. 3.1.2a and 3.1.2b. In Fig. 3.1.2a, the reflections of two prominent peaks at 2θ 26.624 and 29.420 were observed. The observed X-ray patterns match with the ICSD Reference codes 03-065-0466 Quartz and 01-086-1385 Muscovite-2M1. The observed fractions refer to Quartz and Muscovite- 2M1 crystalline fractions. Along with these peaks, some other weak peaks also matched with reference peaks of 01-075-8291 chlorite-ll-4,01-080-1108 Biotite, 01-075-1656 Dolomite, 01-077-0022 Vermiculite- 2M, and 01-075-8291 clinochlore-llb-4. Fig. 3.1.2b indicates three prominent peaks at 2θ 26.661, 29.442 and 30.984. The observed fractions matched with ICSD Reference codes 01-087-2096 Quartz, 01-072-4582 Calcite and 01-076-6603 Vermiculite. Silicate minerals found in smast were illite, muscovite, vermiculite, chlorite, clinochlore and quartz. The chemical composition of the minerals is given in (Table 3.1.1).
Fig. 3.1.2. XRD patterns of Kashmir smast (a) from the floor and (b) from the wall along-with the matched peaks of the mineral ICSD (Inorganic Crystal Structure Database)
Table 3.1.1. List of minerals obtained from smast-5 floor and smast-7 wall of Kashmir smast samples
Table 3.1.2. Concentration of elements in sample collected from cave floor and outside cave soil (control)
Metals (mg/kg) Soil samples Ni Cr Co Cu Zn Ca Mg Pb
Cave soil 0.965 0.571 0.266 1.824 12.7311 332.938 1.2576 1.31
Control soil 10.4 8.74 0.810 4.7 36.41 121.65 1.023 8.14
Weight percent mineral phases were used to estimate the SIROQUANT (Fig. 3.1.3) considering 100% crystalline compound to calculate the quantitative analysis. Fig. 3.1.3(a) shows that the vermiculite, illite and chlorite were the most abundant minerals in smast-5. Similarly, Fig. 3.1.3(b) shows that the vermiculite-2M1, muscovite, clinochlore-llb are the most abundant minerals in smast-7. The FTIR absorption peaks from smast were observed to determine the major and minor constituent minerals present in the sample smast-7 (Fig. 3.1.4). The samples analyzed were mixtures of the minerals such as, silicon oxide, calcite, quartz, muscovite,
clinochlore, nimite, biotite and vermiculite. Various peaks appeared indicating the presence of a variety of minerals. Mass loss steps were observed from Fig. 5 at 77, 200 and 280, 400 and 790°C with mass losses of 10.23, 21.55, 5.20 and 7.58% recorded due to carbonates. SEM observations (Fig. 3.1.6) suggest that smast (cave) clay particles show poorly crystallized clasts with angula, irregular outlines, swirly texture with face-to-face arrangement of clay grains. Si, Al, Fe were found enriched within the samples.
Fig. 3.1.3. Quantitative analysis of minerals, A. Wall soil sample smast-7, B. Floor soil sample smast-5
Fig. 3.1.4. Infrared spectra of Smast-7 wall
Soil Analysis
Atomic absorption spectroscopy was performed to determine the concentration of elements in the cave soil sample. Ca was 332.938 mg/kg as compared to 121.65 mg/kg in control soil, Mg was 1.2576 mg/kg in cave soil and 1.023 mg/kg in control soil and that of Ni, Cr, Co, Cu, Zn and Pb were much lower than those found in the control soil (Table 3.1.2).
Fig. 3.1.5. TGA (Thermogravimetric Analysis) plots of Kashmir smast (sample-5 Floor and sample-7 wall)
Microbiology Results Colony Forming Unit (CFU/ml) Numbers of viable cells per ml were calculated in floor brown soil sample collected from Kashmir cave. The bacterial counts (CFU/ml) were calculated as 5.25 x 104/ml.
Screening of bacterial isolates for the antibacterial activity Initial screening resulted in isolation of phenotypically different 4 bacterial strains showing antimicrobial activity (Fig. 3.1.7) against four test organisms. Out of 4, the strain B. licheniformis KC2-MRL showed maximum zone of inhibition (28 mm against Micrococcus, 20 mm against E. coli, 14 mm against Staphylococcus aureus and 15 mm
against Klebsiella). Therefore, it was selected for further analysis on the basis of greatest zone of inhibition.
Fig. 3.1.6. FE-SEM micrograph & EDS spectra of (a) smast-7 wall and (b) smast- 5 floor
Identification of antibiotic producing isolates The 16S rRNA gene sequences of the antibiotic producing cave bacteria have been submitted to NCBI GenBank. The isolates KC1-MRL, KC2-MRL, KC3-MRL and KC4-MRL were identified as Serratia sp. KC1-MRL (Accession No. KC128829.1), Bacillus licheniformis KC2-MRL (Accession No. KC128830.1), Bacillus sp. KC3- MRL (Accession No. KC128831.1) and Stenotrophomonas sp. KC4-MRL (Accession No. KC128832.1) (Fig 3.1.8a and 3.1.8b). Among these isolates the strain KC2-MRL, identified as Bacillus licheniformis, showed maximum zone of inhibition (28 mm) against Gram positive Micrococcus luteus.
Growth curve
At 25˚C and 35˚C Bacillus licheniformis KC2-MRL confirmed late growth as compared to 45˚C with an elongated lag phase and a rise in growth. While, at 45˚C rise in growth was detected.
Fig.3.1.7. Nutrient agar plate showing the zones of inhibition against the clinical isolates
Selection of medium Maximum antimicrobial activity was found in Nutrient Broth (NB) medium. The best antibacterial activity was observed in case of NB after 24 hours of incubation, with zone of inhibition of 28 mm against M. luteus, 20 mm against S. aureus, 11 mm against Klebsiella and 8 mm against E. coli. With the passage of time, decrease in antimicrobial activity was noted with the increase in growth. The antibacterial activity decreased with passage of time in all media except the NB, i.e. after 48 hrs, 42, 28, 23 and 16 mm zone of inhibition was observed against M. luteus, S. aureus, Klebsiella and E. coli, respectively. External Factors (incubation time, pH and temperature) Best antimicrobial activity (42 mm) of B. licheniformis KC2-MRL was observed against M. luteus, 28 mm against S. aureus, 23 mm against Klebsiella, 16 mm against E. coli after 48 hours of incubation, while there was a decrease in the size of zone after 48 hours of incubation (Fig. 3.1.9). With passage of time, gradual decrease in antimicrobial activity of B. licheniformis KC2-MRL was observed against M. luteus, S. aureus, Klebsiella and E. coli (Fig. 3.1.9).
Uncul bact-USA(FJ849582) Steno.maltophilia-Blg(AY040357) Uncul bact-USA(FJ849582) Uncul bact-Finland(FM873444) Steno.maltophilia-Blg(AY040357) Uncul bact-Finland(FM873497) Uncul bact-USA(FJ849359) Uncul bact-Finland(FM873444) Uncul bact-USA(FJ849526) Uncul bact-USA(FJ849584) Uncul bact-Finland(FM873497) Uncultured bacterium-USA(FJ849590) Uncl bact-USA(FJ849628) Uncul bact-USA(FJ849359) B.licheniformis-Jpn(AB525389) Serratia.sp-Ch(JN859196) Uncul bact-USA(FJ849526) B.licheniformis-Spn(AY479984) Uncul bact-USA(FJ849584) B.licheniformis-Irn(FN678352) B.licheniformis-Mex(HQ634209) Uncultured bacterium-USA(FJ849590) Bacillus.sp-Chn(EF026995) Uncul bact-Spn(HQ218495) Uncl bact-USA(FJ849628) Serratia.sp-Fr(GQ416051) Serratia.sp-Fr(GQ416052) B.licheniformis-Jpn(AB525389) Serratia.sp-Russ(JF327457) Bacillus.sp-Bglm(HE586339) Serratia.sp-Ch(JN859196) S.proteamaculans-USA(HQ219941) S.proteamaculans-USA(HQ219942) B.licheniformis-Spn(AY479984) S.rhizophila-Chn(GQ359325) B.licheniformis-Irn(FN678352) Uncul bact-UK(FJ184322) S.rhizophila-Chn(GU391467) B.licheniformis-Mex(HQ634209) S.liquefaciens-Ch(HQ334999) Bacillus.sp-Brzl(JF309228) Bacillus.sp-Chn(EF026995) Serratia.sp-USA(JN423857) S.liquefaciens-USA(NR 042062) Uncul bact-Spn(HQ218495) S.rhizophila-Chn(FJ529915) B.licheniformis-Fr(FN666245) Serratia.sp-Fr(GQ416051) Stenotrophomonas rhizophila-Fr(JF711015) Serratia.sp-Fr(GQ416052) B.licheniformis-UK(FN397495) B.licheniformis-UK(FN397507)(2) Serratia.sp-Russ(JF327457) B.licheniformis-UK(FN397507) Serratia.sp-Ch(JF833851) Bacillus.sp-Bglm(HE586339) S.liquefaciens-Ind(JN596115) B.licheniformis-Itl(HE590856) S.proteamaculans-USA(HQ219941) Stenotrophomonas sp-Chn(AJ551165) S.liquefaciens-Ch(HQ335000) S.proteamaculans-USA(HQ219942) Serratia.sp-Blgm(JN106438) Cluster I B.licheniformis-UK(FN397509) S.rhizophila-Chn(GQ359325) SN2-Pak-2012 Uncul bact-UK(FJ184322) Stenotrophomonas sp-Chn(GU391493) B.licheniformis-Russ(AF276309) S.rhizophila-Chn(GU391467) B.licheniformis-Jpn(AB680251) Bacillus.sp-Korea(GQ407180) S.liquefaciens-Ch(HQ334999) Serratia.sp-Twn(EF153429) S.proteamaculans-Ger(NR 037112) Bacillus.sp-Brzl(JF309228) B.licheniformis-UK(FN397489) B.licheniformis-UK(FN397503) Serratia.sp-USA(JN423857) B.licheniformis-USA(EU718490) S.liquefaciens-USA(NR 042062) Serratia.sp-Atlantic Ocean(FR744821) S.liquefaciens-Ch(HQ335001) S.rhizophila-Chn(FJ529915) S.proteamaculans-Russ(JF327454) SN1.pak. 2011 B.licheniformis-Fr(FN666245) Stenotrophomonas sp-Arg(DQ109991) SN4-Pak(2012) Stenotrophomonas rhizophila-Fr(JF711015) B.licheniformis-Irn(DQ228696) B.licheniformis-UK(FN397484) B.licheniformis-UK(FN397495) B.licheniformis-UK(FN397485) B.licheniformis-UK(FN397486) B.licheniformis-UK(FN397507)(2) B.licheniformis-Ind(JN118574) B.licheniformis-UK(FN397507) B.licheniformis-Pol(JN180125) B.licheniformis-Mex(HQ634208) Serratia.sp-Ch(JF833851) B.licheniformis-USA(AF372616) S.rhizophila-Ind(FM955853) S.liquefaciens-Ind(JN596115) Serratia.sp-Moroco(JF974140) Bacillus.sp-Brzl(JF309230) B.licheniformis-Itl(HE590856) Serratia.sp-Ch(HQ334998) B.licheniformis-UK(FN397487) Stenotrophomonas sp-Chn(AJ551165) Bacillus.sp-Fr(EF471917) B.licheniformis-Ind(JF700488) S.liquefaciens-Ch(HQ335000) B.licheniformis-Ind(JF700489) Serratia.sp-Blgm(JN106438) B.licheniformis-Ind(JF414759) S.proteamaculans-USA(HQ219940) B.licheniformis-UK(FN397509) S.grimesii-Ch(HQ242737) Serratia.sp-Ch(JN886902) SN2-Pak-2012 Uncultured.bact-Ch(JF697434) Serratia.sp-Itlay(HQ588852) Stenotrophomonas sp-Chn(GU391493) Serratia.sp-Italy(HQ588837) B.licheniformis-Russ(AF276309) Serratia.sp-Itlay(HQ588839) S.liquefaciens-NBRC12979-Jpn(AB680356) B.licheniformis-Jpn(AB680251) S.grimesii-NBRC13537-Jpn(AB680428) B.licheniformis-Chn(HM055609) Bacillus.sp-Korea(GQ407180) B.licheniformis-S.Af(EU870503) P.carotovorum-Spn(HQ326803) Serratia.sp-Twn(EF153429) B.licheniformis-Spn(AF397062) B.licheniformis-Jpn(AB680253) S.proteamaculans-Ger(NR 037112) B.licheniformis-Jpn(AB680252) B.licheniformis-Chn(DQ351932) B.licheniformis-UK(FN397489) Bacillus.sp-Spn(FR823409) B.licheniformis-UK(FN397503) Bacillus.sp-Jpn(AB425348) SN3-Pak(2012) B.licheniformis-USA(EU718490) Bacterium.FJAT-Chn(JN411103) B.licheniformis-Hol(GQ340506) Serratia.sp-Atlantic Ocean(FR744821) Serratia.sp-Ch(HQ335002) Stenotrophomonas sp-Fr(HQ670711) S.liquefaciens-Ch(HQ335001) Bacillus.sp-Brzl(JF309229) Bacillus.sp-Brzl(JF309229)(2) S.proteamaculans-Russ(JF327454) Bacillus.sp-Jpn(AB188216) SN1.pak. 2011 B.licheniformis-Chn(HM006901) B.licheniformis-Chn(HM006899) Stenotrophomonas sp-Arg(DQ109991) B.licheniformis-Ind(EF059752) B.licheniformis-Twn(DQ993676) SN4-Pak(2012) Bacillus.sp-Chn(JQ068114) B.licheniformis-Chn(HM006898) B.licheniformis-Irn(DQ228696) Bacillus.sp-Ch(HE574482) S.proteamaculans-Russ(JF327473) B.licheniformis-UK(FN397484) Bacillus.sp-Ch(JF772468) Uncultured Bacillus.sp-Chn(JN377797) B.licheniformis-UK(FN397485) Uncult.bacillus-Chn(JN377799) B.licheniformis-UK(FN397486) Bacillus.sp-Fr(EU362149) B.licheniformis-Ind(JN118574) B.licheniformis-Pol(JN180125) B.licheniformis-Mex(HQ634208) B.licheniformis-USA(AF372616) S.rhizophila-Ind(FM955853) Serratia.sp-Moroco(JF974140) Bacillus.sp-Brzl(JF309230) Serratia.sp-Ch(HQ334998) B.licheniformis-UK(FN397487) Bacillus.sp-Fr(EF471917) B.licheniformis-Ind(JF700488) B.licheniformis-Ind(JF700489) B.licheniformis-Ind(JF414759) S.proteamaculans-USA(HQ219940) S.grimesii-Ch(HQ242737) Serratia.sp-Ch(JN886902) Uncultured.bact-Ch(JF697434) Serratia.sp-Itlay(HQ588852) Serratia.sp-Italy(HQ588837) Serratia.sp-Itlay(HQ588839) S.liquefaciens-NBRC12979-Jpn(AB680356) S.grimesii-NBRC13537-Jpn(AB680428) B.licheniformis-Chn(HM055609) B.licheniformis-S.Af(EU870503) P.carotovorum-Spn(HQ326803) B.licheniformis-Spn(AF397062) B.licheniformis-Jpn(AB680253) B.licheniformis-Jpn(AB680252) B.licheniformis-Chn(DQ351932) Bacillus.sp-Spn(FR823409) Bacillus.sp-Jpn(AB425348) SN3-Pak(2012) Bacterium.FJAT-Chn(JN411103) B.licheniformis-Hol(GQ340506) Serratia.sp-Ch(HQ335002) Stenotrophomonas sp-Fr(HQ670711) Bacillus.sp-Brzl(JF309229) Bacillus.sp-Brzl(JF309229)(2) Bacillus.sp-Jpn(AB188216) B.licheniformis-Chn(HM006901) B.licheniformis-Chn(HM006899) B.licheniformis-Ind(EF059752) B.licheniformis-Twn(DQ993676) Bacillus.sp-Chn(JQ068114) B.licheniformis-Chn(HM006898) Bacillus.sp-Ch(HE574482) S.proteamaculans-Russ(JF327473) Bacillus.sp-Ch(JF772468) Uncultured Bacillus.sp-Chn(JN377797) Uncult.bacillus-Chn(JN377799) Bacillus.sp-Fr(EU362149) Uncul bact-USA(FJ849582) Steno.maltophilia-Blg(AY040357) Uncul bact-Finland(FM873444) Uncul bact-Finland(FM873497) Uncul bact-USA(FJ849359) Uncul bact-USA(FJ849526) Uncul bact-USA(FJ849584) Uncultured bacterium-USA(FJ849590) Uncl bact-USA(FJ849628) B.licheniformis-Jpn(AB525389) Serratia.sp-Ch(JN859196) B.licheniformis-Spn(AY479984) B.licheniformis-Irn(FN678352) B.licheniformis-Mex(HQ634209) Bacillus.sp-Chn(EF026995) Uncul bact-Spn(HQ218495) Serratia.sp-Fr(GQ416051) Serratia.sp-Fr(GQ416052) Serratia.sp-Russ(JF327457) Bacillus.sp-Bglm(HE586339) S.proteamaculans-USA(HQ219941) S.proteamaculans-USA(HQ219942) S.rhizophila-Chn(GQ359325) Uncul bact-UK(FJ184322) S.rhizophila-Chn(GU391467) S.liquefaciens-Ch(HQ334999) Bacillus.sp-Brzl(JF309228) Serratia.sp-USA(JN423857) S.liquefaciens-USA(NR 042062) S.rhizophila-Chn(FJ529915) B.licheniformis-Fr(FN666245) Stenotrophomonas rhizophila-Fr(JF711015) B.licheniformis-UK(FN397495) B.licheniformis-UK(FN397507)(2) B.licheniformis-UK(FN397507) Serratia.sp-Ch(JF833851) S.liquefaciens-Ind(JN596115) B.licheniformis-Itl(HE590856) Stenotrophomonas sp-Chn(AJ551165) S.liquefaciens-Ch(HQ335000) Serratia.sp-Blgm(JN106438) B.licheniformis-UK(FN397509) SN2-Pak-2012 Stenotrophomonas sp-Chn(GU391493) B.licheniformis-Russ(AF276309) B.licheniformis-Jpn(AB680251) Bacillus.sp-Korea(GQ407180) Serratia.sp-Twn(EF153429) S.proteamaculans-Ger(NR 037112) B.licheniformis-UK(FN397489) B.licheniformis-UK(FN397503) B.licheniformis-USA(EU718490) Serratia.sp-Atlantic Ocean(FR744821) S.liquefaciens-Ch(HQ335001) S.proteamaculans-Russ(JF327454) SN1.pak. 2011 Stenotrophomonas sp-Arg(DQ109991) SN4-Pak(2012) B.licheniformis-Irn(DQ228696) B.licheniformis-UK(FN397484) B.licheniformis-UK(FN397485)
B.licheniformis-UK(FN397486) Uncul bact-USA(FJ849582) Steno.maltophilia-Blg(AY040357) B.licheniformis-Ind(JN118574) Uncul bact-Finland(FM873444) Uncul bact-Finland(FM873497) B.licheniformis-Pol(JN180125) Uncul bact-USA(FJ849359) Uncul bact-USA(FJ849526) B.licheniformis-Mex(HQ634208) Uncul bact-USA(FJ849584) Uncultured bacterium-USA(FJ849590) B.licheniformis-USA(AF372616) Uncl bact-USA(FJ849628) B.licheniformis-Jpn(AB525389) Serratia.sp-Ch(JN859196) S.rhizophila-Ind(FM955853) B.licheniformis-Spn(AY479984) B.licheniformis-Irn(FN678352) Serratia.sp-Moroco(JF974140) B.licheniformis-Mex(HQ634209) Bacillus.sp-Chn(EF026995) Bacillus.sp-Brzl(JF309230) Uncul bact-Spn(HQ218495) Serratia.sp-Fr(GQ416051) Serratia.sp-Ch(HQ334998) Serratia.sp-Fr(GQ416052) Serratia.sp-Russ(JF327457) B.licheniformis-UK(FN397487) Bacillus.sp-Bglm(HE586339) S.proteamaculans-USA(HQ219941) S.proteamaculans-USA(HQ219942) Bacillus.sp-Fr(EF471917) S.rhizophila-Chn(GQ359325) Uncul bact-UK(FJ184322) B.licheniformis-Ind(JF700488) Cluster II S.rhizophila-Chn(GU391467) S.liquefaciens-Ch(HQ334999) B.licheniformis-Ind(JF700489) Bacillus.sp-Brzl(JF309228) Serratia.sp-USA(JN423857) B.licheniformis-Ind(JF414759) S.liquefaciens-USA(NR 042062) S.rhizophila-Chn(FJ529915) S.proteamaculans-USA(HQ219940) B.licheniformis-Fr(FN666245) Stenotrophomonas rhizophila-Fr(JF711015) B.licheniformis-UK(FN397495) S.grimesii-Ch(HQ242737) B.licheniformis-UK(FN397507)(2) B.licheniformis-UK(FN397507) Serratia.sp-Ch(JN886902) Serratia.sp-Ch(JF833851) S.liquefaciens-Ind(JN596115) Uncultured.bact-Ch(JF697434) B.licheniformis-Itl(HE590856) Stenotrophomonas sp-Chn(AJ551165) Serratia.sp-Itlay(HQ588852) S.liquefaciens-Ch(HQ335000) Serratia.sp-Blgm(JN106438) Serratia.sp-Italy(HQ588837) B.licheniformis-UK(FN397509) SN2-Pak-2012 Serratia.sp-Itlay(HQ588839) Stenotrophomonas sp-Chn(GU391493) B.licheniformis-Russ(AF276309) B.licheniformis-Jpn(AB680251) S.liquefaciens-NBRC12979-Jpn(AB680356) Bacillus.sp-Korea(GQ407180) Serratia.sp-Twn(EF153429) S.grimesii-NBRC13537-Jpn(AB680428) S.proteamaculans-Ger(NR 037112) B.licheniformis-UK(FN397489) B.licheniformis-Chn(HM055609) B.licheniformis-UK(FN397503) B.licheniformis-USA(EU718490) B.licheniformis-S.Af(EU870503) Serratia.sp-Atlantic Ocean(FR744821) S.liquefaciens-Ch(HQ335001) P.carotovorum-Spn(HQ326803) S.proteamaculans-Russ(JF327454) SN1.pak. 2011 Stenotrophomonas sp-Arg(DQ109991) B.licheniformis-Spn(AF397062) SN4-Pak(2012) B.licheniformis-Irn(DQ228696) B.licheniformis-Jpn(AB680253) B.licheniformis-UK(FN397484) B.licheniformis-UK(FN397485) B.licheniformis-Jpn(AB680252) B.licheniformis-UK(FN397486) B.licheniformis-Ind(JN118574) B.licheniformis-Chn(DQ351932) B.licheniformis-Pol(JN180125) B.licheniformis-Mex(HQ634208) Bacillus.sp-Spn(FR823409) B.licheniformis-USA(AF372616) S.rhizophila-Ind(FM955853) Serratia.sp-Moroco(JF974140) Bacillus.sp-Jpn(AB425348) Bacillus.sp-Brzl(JF309230) Serratia.sp-Ch(HQ334998) SN3-Pak(2012) B.licheniformis-UK(FN397487) Bacillus.sp-Fr(EF471917) Bacterium.FJAT-Chn(JN411103) B.licheniformis-Ind(JF700488) B.licheniformis-Ind(JF700489) B.licheniformis-Hol(GQ340506) B.licheniformis-Ind(JF414759) S.proteamaculans-USA(HQ219940) Serratia.sp-Ch(HQ335002) S.grimesii-Ch(HQ242737) Serratia.sp-Ch(JN886902) Stenotrophomonas sp-Fr(HQ670711) Uncultured.bact-Ch(JF697434) Serratia.sp-Itlay(HQ588852) Serratia.sp-Italy(HQ588837) Bacillus.sp-Brzl(JF309229) Serratia.sp-Itlay(HQ588839) S.liquefaciens-NBRC12979-Jpn(AB680356) Bacillus.sp-Brzl(JF309229)(2) S.grimesii-NBRC13537-Jpn(AB680428) B.licheniformis-Chn(HM055609) Bacillus.sp-Jpn(AB188216) B.licheniformis-S.Af(EU870503) P.carotovorum-Spn(HQ326803) B.licheniformis-Chn(HM006901) B.licheniformis-Spn(AF397062) B.licheniformis-Jpn(AB680253) B.licheniformis-Chn(HM006899) B.licheniformis-Jpn(AB680252) B.licheniformis-Chn(DQ351932) Bacillus.sp-Spn(FR823409) B.licheniformis-Ind(EF059752) Bacillus.sp-Jpn(AB425348) SN3-Pak(2012) B.licheniformis-Twn(DQ993676) Bacterium.FJAT-Chn(JN411103) B.licheniformis-Hol(GQ340506) Bacillus.sp-Chn(JQ068114) Serratia.sp-Ch(HQ335002) Stenotrophomonas sp-Fr(HQ670711) B.licheniformis-Chn(HM006898) Bacillus.sp-Brzl(JF309229) Bacillus.sp-Brzl(JF309229)(2) Bacillus.sp-Ch(HE574482) Bacillus.sp-Jpn(AB188216) B.licheniformis-Chn(HM006901) B.licheniformis-Chn(HM006899) S.proteamaculans-Russ(JF327473) B.licheniformis-Ind(EF059752) B.licheniformis-Twn(DQ993676) Bacillus.sp-Ch(JF772468) Bacillus.sp-Chn(JQ068114) B.licheniformis-Chn(HM006898) Uncultured Bacillus.sp-Chn(JN377797) Bacillus.sp-Ch(HE574482) S.proteamaculans-Russ(JF327473) Uncult.bacillus-Chn(JN377799) Bacillus.sp-Ch(JF772468) Uncultured Bacillus.sp-Chn(JN377797) Bacillus.sp-Fr(EU362149) Uncult.bacillus-Chn(JN377799) Bacillus.sp-Fr(EU362149)
Fig. 3.1.8. Phylogenetic tree of all four species with related sequences in NCBI
The maximum antibacterial activity i.e. 22 mm and 28 mm was observed against M. luteus and S. aureus, respectively, 17 mm against E. coli and no activity against
° Klebsiella, at 35 C after 48 hrs of incubation with growth OD600 2.25. The activity in terms of zone of inhibition decreased with increase in temperature and low activity was
° observed at 45 C (OD600 0.306). After 72 hours of incubation, the zone of 9 mm was observed against M. luteus, 10 mm against S. aureus, 6 mm against Klebsiella and no activity against E. coli. The diameter of zone of inhibition gradually decreased with time (Fig. 3.1.9).
Fig. 3.1.9. Effect of time of incubation, pH and temperature on the growth and antimicrobial activity of B. licheniformis KC2-MRL against M. luteus, S. aureus, Klebsiella and E. coli
Effect of pH (5, 6, 7 and 8) on the production of antibiotics was studied. Activity in terms of zone of inhibition (mm) was measured against test organisms i.e. M. luteus, Klebsiella, E. coli and S. aureus. At pH 5, the best activities as 23 mm against S. aureus, followed by M. luteus, Klebsiella and E. coli, were observed after 24 hrs of incubation. The second best activity was observed at pH 6, and the gradual decrease in activity was observed with passage of time (Fig. 3.1.9). Temperature and pH Stability
To check the stability of antimicrobial compounds at different temperatures, the cell free supernatant (CFS) was treated at 15, 25, 35 and 45°C for 1 hour i.e. the antibacterial activity (26 mm) was observed until 40°C but the activity decreased at a temperature above 40°C and was totally lost with further rise in temperature. The antimicrobial compound produced by B. licheniformis KC2-MRL was stable at range pH 5-8. The highest activity was observed at pH 5 and 6, whereas, activity decreased at pH 7 and 8. B. licheniformis KC2-MRL produced optimum activity at acidic pH 5-6 after 24 hrs of incubation. The antimicrobial activity was stable up to 45°C. It was found that antimicrobial compounds were stable at range of 5 -7. Antibiotic susceptibility test Vancomycin, Nalidixic acid, Cefotoxime, Ampicillin, Amoxicillin, Imipenem, Methicillin, Cefoten and Levofloxacin were tested to check the susceptibility of Bacillus licheniformis KC2-MRL. The organism was more susceptible to Levofloxacin by producing 40 mm zone of inhibition (Fig. 3.1.10).
45
40 VA30 35 NA30 30 CTX30 25 Amp25
20 Amc30 IPM10 15 Met5 10 Zone of Inhibition (mm) Inhibition of Zone CN10 5 LEV(5) 0 KC1 KC2 KC3 KC4
Fig. 3.1.10. Zone of inhibition of our four antibiotic producing strains (Serratia sp. KC1-MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3-MRL and Stenotrophomonas sp. KC4-MRL) against selected antibiotics in mm
Fourier Transform Infrared Spectroscopic analysis of antibacterial compound produced by B. licheniformis KC2-MRL We used a solution of bacitracin as a standard. FTIR spectrum of B. licheniformis KC2-MRL’s precipitated protein was compared with the spectrum of control. FTIR spectrum of bacitracin showed the absorption bands at 3295.63, 3016.9, 2133.64, and 1635 cm-1 which were corresponding to NH, CH, C-C, and C=C groups. Similarly, in case of B. licheniformis KC2-MRL protein the absorption bands appeared at 3271.98, 3016.90, 2120.12, 1635.20 and 1076.22 cm-1 which were attributing to the NH, CH, C=C and C-N (Fig. 3.1.11).
Fig. 3.1.11. Comparison of FTIR spectra of control (Bacitracin) and the antibacterial compound produced by B. licheniformis KC2-MRL
DISCUSSION Solution caves are formed in carbonate and sulfate rocks such as limestone, dolomite, marble, and gypsum by the action of slowly moving ground water that dissolves the rock to form tunnels, irregular passages, and even large caverns along joints and bedding planes (Davies and Morgan, 2000). Caves usually have extremely starved conditions, but still they contain diverse and often unique microbial communities (Barton, 2007). Caves on other worlds such as Mars may provide protected sites for extraterrestrial life forms (Nelson, 1996). Subsurface of Earth is considered as the the best possible site to look for microbial life and the characteristic lithologies that indicates the remnants of life. The subterrain, where surface conditions are hostile, like Mars and subsurface offers habitat for signs of lifeforms (Boston et al., 2001). Microbial analysis of caves showed Bacillus as the most commonly detected microbial genus (Adetutu et al., 2012). It is important to understand how the ecosystems are operating and accommodating microbial diversity. The rock composition and mineralogy can be helpful to understand the geomicrobiology and potential metabolic capabilities of the microorganisms to use ions within the rock as nutrients and for chemolithotrophic energy production. Cave sediments can therefore act as reservoirs of microorganisms (Adetutu et al., 2012). The use of these ions may be supported by the formation of a corrosion residue, through microbial scavenging activities (Barton, 2007). Cave microorganisms also have potential to produce unique antibiotics and cancer treatment drugs (Onaga, 2001). Minerals have profound effect on the production of antibiotics by microorganisms. Basak and Majumdar (1975) reported that kanamycin production by Streptomyces kanamyceticus ATCC 12853 required magnesium sulfate and potassium phosphate (0.4 and 1.0 g/L, respectively), Fe and Zn (0.25 and 0.575 μg/ml, respectively), Mn and Ca did not have any effect, whereas, Cu, Co, Ni, and V have inhibitory effect on growth and production of kanamycin. Divalent ions as Mn2+, Cu2+, Fe2+ stimulated AK-111-81 antibiotic biosynthesis by Streptomyces hygroscopicus 111-81 (Gesheva et al., 2005). The divalent metal ions (Mg, Fe and Mn) sodium dihydrogen phosphate were found essential for bacitracin production by
Bacillus licheniformis, whereas Na2SO4 and CaCl2 decreased the bacitracin yield (Yousaf, 1997). The sample from which B. licheniformis KC2-MRL was isolated, was reddish- brown in colour. Brown soils are usually low in organic matter. The red- brownish red soil is heavy and clay-rich (silty-clay to clayey) soil, strongly reddish, developed on limestone
or dolomite. Terra rossa derives is usually derived from the insoluble residue of the underlying limestone. Following dissolution of calcium carbonate by rain, clay contained in limestone sediments with other insoluble substances or rock fragments, forming discontinuous residual layers variable in depth. Under oxidizing conditions iron oxides appear, which produces the characteristic red color. According to this theory, Terra rossa is usually derived a polygenetic relict soil, formed during the Tertiary and subjected to hot and humid periods during the Quaternary (Jordán, 2014). XRD-analysis of the smast confirmed the presence of clay minerals, carbonates, silicates (Hill, 1999). Minerals are produced as a result of intense chemical weathering on land under possibly tropical conditions, where abundant rainfall favored ionic transfer and pedogenic development (Millot, 1970). Carbonates found from Kashmir smast are predominantly calcite and traces of dolomite (Vogel and Mylroie, 1990; Schwabe and Carew, 1993). Mostly, illite is found in fault zones and also occurs as clay floor deposits (Hill, 1999). Illite is commonly present as little-altered disintegrated particles (Weaver, 1989). Pedogenic clay minerals are derived from moderate chemical weathering, which generally develop in poorly drained tropical to subtropical areas of low relief, marked by flooding during humid seasons and subsequent concentration of solutions in the soil during dry seasons. Al, Fe and Si are transported by mean of water saturation during wet seasons, concentration for mineral growth takes place during in dry seasons (Chamley, 1989). During pedogenesis, chlorite transforms into kaolinite, and in intense weathering laterite soils chlorite would be completely eliminated (Vicente and Elsass, 1997). The clay mineral accumulation of illite, kaolinite, chlorite, dolomite and muscovite in smast are probably indicative of changes in degree of weathering, and thus reflect the changes in climatic conditions. The degree of weathering related to the presence of SiO, Al2O3 and show a similar pattern to clay indices (Tardy and Nahon, 1985; Zhao and Yang, 1995). The mineral assemblages investigated in smast are diverse. The quantitative mineral analysis technique SIROQUANT described mineral compositions of rocks, including clay mineral content. Thermal analysis offers an important technique for the determination of thermal stability of minerals and roughly estimating organic content of the samples. Importantly, the decomposition steps can be obtained and mechanism of decomposition of the mineral is determined. Generally, the theoretical mass loss of water is 10.46%, the structural disorganization upon thermal treatment may occur in response to the loss of hydration water, which could provoke
collapse of the crystalline structure (Doak and Gallagher, 1965). The two overlapping mass loss steps at 263 and 280°C are attributed to the hydroxyl group (Palmer and Frost, 2010). The higher mass loss at 280oC is believed to be due to the loss of both OH and ° CO3. The broad mass loss at 485 C is ascribed to the loss of carbonate as carbon dioxide ° (CO2) (Frost and Hales, 2009). The higher temperature mass loss at 828 C is attributed to the Mg. Clay particles were observed having poorly crystallized clasts with angula, irregular outlines, swirly texture with face-to-face arrangement of clay grains, as also reported by Manju and Nair (2001) in Madayi kaolin deposit, North Kerala, India. Generally, intensive weathering of clay flakes show ragged edges, exhibit a rounded outline, or bay-shaped edges, and poor lateral dimension, with a particularly small platy thickness. Analysis shows that Si, Al, Fe were enriched within the samples, which probably are the reflection of trapped minerals such as Quartz, feldspar, clay minerals and Fe-oxide (Jeong and Kim, 2003). FTIR analysis showed the peaks at 885 cm-1, 746 cm-1 and 715 cm-1 appeared because of presence of dolomite (White, 1964; Marel and Beutelspacher, 1976). A wide band
-1 around 1020 cm is assigned to quartz SiO2 ( Russell, 1987; Ravisankar et al., 2012) the peak at 1646 cm-1 is attributed to the bending vibration modes of water (Manoharan and Venkatachalapathy, 2007). Peaks in the region of 2800-3000 cm-1 are ascribed to the C-C stretching which is present in the form of organic matter in the mineral contribution (Maritan and Mazzoli, 2005), and may be due to P-OH stretching bond around 2845 cm-1 and 2935 cm-1. The sharp peak at 2513 cm-1 is due to the presence of silicate minerals like (quartz, nimite, musciovite, vermiculite) (Vedder, 1964). The appearance of broad band in the region of 3000 cm-1 to 3700 cm-1 is attributed to the structural water present in the mineral (vermiculite) and due to the moisture present in the sample (Zadrapa and Zykova, 2010). The hydroxyl and water-stretching region near 3200 cm-1 for most hydrated carbonates usually consists of one or two broad band shifted somewhat to lower frequencies due to hydrogen bonding (Nakamoto, 2008; Schrader, 2008), but the appearance of the broad band is due to the interpretation OH and H2O group in the mineral in which some minerals were participating in hydrogen bonding and some were not involved, i.e. non- hydrogen bonded Al-OH units (White, 1964; Marel and Beutelspacher, 1976). Atomic absorption spectroscopy was performed to determine the concentrations of elements; calcium, magnesium, chromium, cobalt, nickel, zinc, copper and lead incave
floor soil sample and it was found that the soil contained very high amount of calcium as compared to the outside soil which was used as a control taken from above the cave. Microbiology Screening of bacterial isolates for the antibacterial activity The biocapacity of bacteria inhabiting karstic caves to produce valuable biologically active compounds is still not investigated much (Tomova et al., 2013). Soil is a natural reservoir for microorganisms and their antimicrobial products (Dancer 2004). The four selected strains isolated from cave soil were screened for the production of antibiotics by using agar well diffusion assay against Staphylococcus aureus, Klebsiella, E. coli and Micrococcus luteus. B. licheniformis KC2-MRL was selected for further analysis on the basis of the greatest zone of inhibition. In the present study, B. licheniformis KC2-MRL showed the best antimicrobial activity against M. luteus, followed by S. aureus, Klebsiella and E. coli after 48 hrs of incubation. Identification Studies show that caves are inhabited by different types of microorganisms having unique characteristics. Cave ecosystem has a deficiency of nutrients that is why microorganisms present in the cave mostly compete for the nutrients and fight for survival. Due to this ability of microbes, these microbes have the potential to produce antibiotics against other microbes. There are nine different groups of bacteria reported to be present in the caves i.e. Proteobacteria, Acidobacteria, Planctomycetes, Chloroflexi, Bacteroidetes, Gemmatimonadetes, Nitrospirae, Actinobacteria and Firmicutes (Zhou et al., 2007; Porttillo et al., 2008). Proteobacteria are the dominant bacteria in cave which is about 45% (Zhou et al., 2007). The 16S rRNA gene sequences of the antibiotic producing cave bacteria have been submitted to NCBI GenBank. The isolates KC1-MRL, KC2-MRL, KC3-MRL and KC4-MRL were identified as Serratia sp. KC1-MRL (Accession No. KC128829.1), Bacillus licheniformis KC2-MRL (Accession No. KC128830.1), Bacillus sp. KC3-MRL (Accession No. KC128831.1) and Stenotrophomonas sp. KC4-MRL (Accession No. KC128832.1). In Magura Cave, Bulgaria, Tomonova et al. (2013) reported that Gram-positive bacteria were represented by the genera Bacillus, Arthrobacter and Micrococcus. Soil bacterial species, i.e. Bacillus, Streptomyces and Pseudomonas synthesize a high proportion of agriculturally and medically important antibiotics (Yoshiko et al., 1998; Sharga et al., 2004). Peptide antibiotics are the major group of antibiotics (Pinchuk et al., 2002). Antibiotic producing microorganisms can be found in different habitats but
the majority are common inhabitants of soil. Caves contain abundant Actinobacteria, which are valuable sources of novel antibiotics, replacing currently ineffective antibiotics (Montano and Henderson, 2012). Molecular analysis of a sample from Kashmir cave showed the presence of different bacterial strains. Isolated strains were screened for the production of antimicrobial compounds by using agar well diffusion assay. Ducluzeau et al. (1978) isolated Bacillus licheniformis that was active against Clostridium perfringens or Lactobacillus sp. Muhammad et al. (2009) also observed that Bacillus metabolites showed activity against M. luteus and S. aureus. Bacitracin is a major polypeptide antibiotic produced by Bacillus licheniformis and Bacillus subtilis, using M. luteus as a test organism (Vieira et al., 2011). B. licheniformis isolated from marine sediments showed best antimicrobial activity against pathogenic test strains; S. aureus, E. coli and P. aeruginosa (Eldewany et al., 2011). The antibiotic production depends upon the composition of the medium, which is required for cell biomass and for its maintenance (Stanbury et al., 1995). Maximum activity was found when organism was grown in Nutrient broth. Similarly, Vieira et al. (2011) used Nutrient broth for the growth of Bacillus licheniformis and incubated at 46ºC in a shaking incubator at 150 rpm. Whereas, Al- Janabi and Hussein (2006), Yilmaz et al. (2006) and Al-Ajlani and Hasnain. (2010) also reported maximum production of antimicrobial compound by Bacillus sp. in NB medium under varying temperatures. Effect of time of incubation, pH and temperature on antimicrobial activity External factors can also affect the growth of microorganisms and the production of antibiotics (Marwick et al., 1999), it has been reported that environmentalfactors such as temperature, pH and incubation influence on antibiotic production (Iwai et al., 1973). In our study, optimum temperature for antimicrobial compound production was observed at 30-35°C. Berdy (1974) and Al-Gelawi et al. (2007) observed production of bacitracin and other antibiotics by B. licheniformis (Zarei, 2012) at 37°C and also at 30°C (Eldewany et al., 2011) as well. We found that our selected organism showed optimum activity at pH 5-6. Perlman and Flickinger (1979) reported pH 6.5 for the optimum production of antibiotics by B. licheniformis. Al-Gelawi et al. (2007) found maximum bacitracin production rate (192 units’ mL-1) at pH 7.5. The similar study was conducted in which antimicrobial activity was best at the wide pH range of 6-8 by Bacillus sp. (Gulahmadov et al., 2006). Newly emergent infectious diseases, re-emerging diseases and multidrug-resistant bacteria
mean that there is a persistent need to produce novel antimicrobial compounds (Uzair et al., 2009). We performed the antibiotic susceptibility test, in which the B. licheniformis KC2-MRL was found resistant to cefotaxime (CTX) and was more susceptible to levofloxacin by producing 40 mm zone of inhibition. B. thuringiensis RSKK 380 was reported to be unaffected by cephazolin, cefoxitin and cefamandole (Yimaz et al., 2006). Temperature and pH stability Our results show that the antibacterial activity was stable up to 45°C. Similar study by He et al. (2006), reported B. licheniformis to be stable at 25°C for 6 hrs and inactivated above 40°C. However, in some cases the antimicrobial compounds retained its activity even after autoclaving the sample at 121°C (Fontoura et al., 2009; Tabbene et al., 2009; Uzair et al., 2009; Ebrahimipour et al., 2010). At the same time, sensitivity to different pH was also evaluated in the present study and the antimicrobial compound was found to be stable at pH 5-7. The similar study, in which antimicrobial activity was found to be stable at pH range of 7 was reported by He et al. (2006). The stability of antibacterial activity over a range of pH 7 and after heat treatment might be useful in several industrial applications (Tabbene et al., 2009). Our study showed best activity against M. luteus, S. aureus and E. coli after 48 hours of incubation. A similar study by Aslim et al. (2002), who showed the maximum zone of inhibition after 24-48 hrs. Bacillus licheniformis KC2-MRL was further tested for antibiotic sensitivity by using different antibiotics i.e. vancomycin, nalidixic acid, cefotoxime, ampicillin, amoxicillin, imipenem, methicillin, cefoten and levofloxacin. It was found that the selected strain was more susceptible to levofloxacin by producing 40 mm zone of inhibition (Fig. 9). Sirtori et al. (2006) reported clear absorption peaks at 3,500, 2,925, 1,639, and 1,546 cm-1 corresponding to the O-H, C-H, C-N and angular deformation of the N-H band. Kong and Shaoning (2007) also detected bands at peaks of 3100, 1600-1690, 1480- 1575 and 1229-1301 cm-1 which are assigned to N-H, C=O, CN and NH. Kumar et al. (2010) reported absorption bands at 1670, 1539, 1418 and 1488 cm-1 attributing to N- H, C=O, O-H and CO. Our study explored the ability of cave microorganisms to produce antibiotics and characterization of the producer strain. Due to the internal acidic environment and high calcium concentration in cave, B. licheniformis KC2-MRL grew better under acidic
conditions at temperatures higher than that in the cave. From our study it can be concluded that caves of Pakistan have never been explored for the presence of bacteria with regards to diversity or having ability to produce novel antimicrobial metabolites. These metabolites as well as other caves can be further investigated to find some bioactive compound with unique characteristic.
BIODEGRADATION OF POLYETHYLENE BY BACTERIAL STRAINS
ISOLATED FROM KASHMIR CAVE, BUNER, PAKISTAN
Abstract:
Low density polyethylene (LDPE) is used for making common shopping bags and plastic sheets and is a significant source of environmental pollution. The present study was aimed at testing the ability of bacterial strains identified as Serratia sp. KC1-MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3-MRL and Stenotrophomonas sp. KC4-
MRL isolated from a limestone cave to degrade polyethylene. These strains were isolated from soil of Kashmir Smast, a limestone cave in Buner, Pakistan. These strains showed antibacterial activity against Micrococcus luteus, Klebsiella sp., Pseudomonas sp., and
Staphylococcus aureus. The pieces of LDPE plastic were incubated along with bacterial strains for a period of one month and then analyzed. Degradation was observed in terms of growth of microorganisms used in consortia, chemical changes in the composition of LDPE by fourier-transform infrared spectroscopy, and changes in physical structure of LDPE by scanning electron microscopy. Maximum growth (107×105 CFU/ml) at 28 °C and subsequent change in chemical and physical properties of plastic were observed in the presence of calcium and glucose. The cave-soil sample had a very high concentration of calcium. The microscopy showed adherence of bacteria with lots of mechanical damage and erosion on the surface of plastic films incubated with bacterial consortia. The spectroscopy showed breakdown and formation of many compounds, as evident by the appearance and disappearance of peaks in LDPE treated with bacterial consortia as compared to the untreated control. We conclude that antibiotic-producing cave bacteria were able to bring about physical and chemical changes in LDPE pieces and degradation of LDPE was enhanced in media augmented with calcium.
INTRODUCTION
Plastics are polymers of carbon, oxygen, and hydrogen and that are synthetically derived from petrochemicals and suitable for a wide range of usage. Since plastics are artificially manufactured they are xenobiotic compounds, and they resist degradation
(Kawai, 2010).
Polyethylene is one of the most commonly used commercial plastics, found in various products ranging from simple plastic bags to artificial limbs (Orhan and Büyükgüngör,
2000; Shimao, 2001). Thermal and mechanical stability and their morphologies make polymeric substances one of the most popular commodity of the modern world (Rivard et al., 1995). Plastic waste is an environmental hazard. Plastic debris poses a direct threat to wildlife. The main dangers associated with plastic objects for most species are related to entanglement and ingestion. Juvenile animals in particular often become entangled in plastic debris, which can result in serious injury as the animal grows, not to mention restriction of movement, preventing animals from properly feeding and, in the case of mammals, breathing (Webb et al., 2012). Due to plastic’s resilience against degradation and its proliferation in industry, the issue of plastic pollution has evolved to become a threat to global ecology.
Management of plastic waste is an ever-increasing problem, and none of the current techniques of solid-waste management completely alleviate all the concerns related to these recalcitrant polymers (Nkwachukwu et al., 2013). One way to deal with these polymers could be to alter the manufacturing process, and new formulations should be developed with special considerations on mechanism for their biodegradation. These alterations could include looking into factors that can aid in biodegradation like pH, temperature, chemical structure, polymeric morphology, presence or absence of certain
additives, and, most importantly the type of organisms that can be involved (Gu and
Gu, 2005).
Degradation of plastics is carried out by organisms that are chemoheterotrophs. Many studies have shown the presence of such bacteria in caves. Bacteria also have the ability to utilize hydrocarbons as a source of energy. Studies have shown that a variety of culturable chemoheterotrophs are present in micro-habitats of caves and catacombs (De
Leo et al., 2012). The microorganisms living under stressful or low-nutrient habitats can develop the ability to use any available nutrient to survive. Studies on a bacterial strain belonging to the Arthrobacter genus from alpine ice showed biodegradation of phenol under various environmental conditions (Margesin et al., 2004). Bacteria present in caves are capable of carrying out a variety of biodegradative and biodeteriorative processes. Extensive studies have reported biodeteriorative effects of microorganisms in cave environment (Cuezva et al., 2012; Schabereiter-Gurtner et al.,
2002).
Despite much study, the knowledge is very limited about microbial life in diverse and extreme habitats like caves. There exists much potential for isolating and studying microbes in caves that have unique and unexplored characteristics of potential commercial applicability. These studies can also be helpful in investigating evolutionary relationships of microorganisms in cave environment. Most caves are characterized as having very low nutrient availability, constant low temperature, and high humidity. Caves can either be terrestrial or aquatic. Some may be rich in specific natural minerals or have exposure to nutrient sources, and therefore different caves will have different types of microorganisms inhabiting various ecological niches (Zada et al., 2016). Fauna, environmental factors, temperature, humidity, organic matter and other environmental factors influence activities such as nutrient cycling, and
geomicrobiological activities including the formation or alteration of cave structures
(Adetutu and Ball, 2014). Cave organisms have evolved some extraordinary abilities to survive and live in this inhospitable environment.
Polyethylene makes a significant contribution in solid waste in developed countries.
Management of this waste can be carried out by chemical, physical, and biological methods. Various natural microflora of soil, including bacteria and fungi, are reported to degrade low-density polyethylene (LDPE) under various physical and chemical environments. This study determines the degradation capacity of four bacterial strains isolated from Kashmir Smast, Khyber Pakhtoon Khwa, Pakistan. The bacteria isolated were previously identified and tested positive for antibiotic production in the
Microbiology Research Laboratory (MRL), Department of Microbiology, Quaid-e-
Azam University, Islamabad. Since antibiotic production and LDPE degradation takes place under stressed environmental conditions, it was hypothesized that bacteria positive for the first character may also be positive for the other character. For this purpose, commercially available LDPE from a shopping bag was used. Bacterial isolates Serratia sp. KC1-MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3-
MRL, and Stenotrophomonas sp. KC4-MRL were used in consortia. The study was carried out in six different medium compositions or modifications, and their individual effects were analyzed by determining total viable cells, fourier-transform infrared spectroscopy, and scanning electron microscopy. Isolated bacteria were inoculated in mineral salt medium, and sterilized LDPE pieces were added in the flasks.
MATERIALS AND METHODS
SAMPLING SITE AND SAMPLE COLLECTION
Soil samples were collected from Kashmir Smast (cave), Nanser, Buner, Khyber
Pakhtunkhwa (GPS coordinates 34°25′42.12″N 72°13′10.82″E) (Fig. 3.2.1). The
Kashmir Smast is a series of natural limestone caves probably of marine origin (Khan,
2013). These caves are located in the Babozai Mountains between Mardan and Buner in northern Pakistan. The only source of water was drip water. Two soil samples were collected from wall and ground surfaces of the cave in sterile Falcon tubes under aseptic conditions. Samples were collected from the dark end of the cave about 188 m from the entrance. This cave is located far away from human access, so human intervention is negligible (Zada et al., 2016). The samples were then brought to the laboratory in an icebox and stored at 4 °C for further processing. The pH and temperature of soil was recorded as 7.2 and 25 °C.
SOIL ANALYSIS BY ATOMIC ABSORPTION
For the quantitative analysis of elements in the soil sample, atomic-absorption spectrophotometry was performed with a AA240FS Fast Sequential Atomic Absorption
Spectrophotometer. Soil digestion was performed to prepare samples for analysis. One gram each of soil from the cave floor and control soil from outside the cave were ground separately and were then mixed in 15 mL aqua regia, heated at 15 °C, and left overnight.
Then 5 mL of HClO4 was added and again heated at 150 °C. The solution almost became dry before brown fumes were produced. Whatman filter paper (No. 42) was used for filtration, and the volume was made up to 50 mL using double-distilled water (Kelly et al.,
2008).
Fig. 3.2.1. Kashmir Smast (cave), Nanser, Buner, Khyber Pakhtunkhwa
SCREENING AND ISOLATION OF LDPE-DEGRADING BACTERIA
Previously identified strains of Serratia sp. KC1-MRL, Bacillus licheniformis KC2-
MRL, Bacillus sp. KC3-MRL, and Stenotrophomonas sp. KCMRL isolated from the cave were used in consortium to carry out biodegradation of polyethylene. Nutrient agar medium was used for isolation of bacterial strains from cave soil. The strains were isolated using standard serial dilution methods and subsequent growth on Nutrient agar plates for 48h at 37 °C. All the isolated strains were screened for polyethylene degradation. For this purpose the strains were incubated for two weeks in 120 mL of
−1 mineral salt medium (g L ): [KH2PO4, 2.0; K2HPO4, 7.0; MgSO4⋅7H2O, 0.1;
ZnSO4⋅7H2O, 0.001; FeSO4⋅7H2O, 0.01; MnSO4⋅6H2O, 0.002; NH4NO3, 1.0;
CuSO4⋅7H2O, 0.0001; pH 7.2] at 37 °C with pieces of polyethylene (1 by 1 cm) (Anwar et al., 2009). Polyethylene used was pretreated by exposing it to UV light for three minutes. At the end of two weeks of incubation, viable cells were counted as CFU mL−1 by serial dilution. Four bacterial strains were found active in terms of growth in the medium. Bacterial colonies were further purified and enriched on nutrient agar plates.
PREPARATION OF INOCULUM
About 10 mL−1 of nutrient broth was inoculated with two or three loops of the pure culture of isolated strains. Bacterial growth was evaluated at 25 °C, 37 °C and 40 °C.
Maximum growth was observed at 37 °C (O.D. at 600 nm). Consortia were developed by taking inocula from each test tube into a separate flask with 100ml of nutrient broth.
Five percent of this prepared consortia was used as inoculum for further biodegradation experiments.
MEDIUM PREPARATION AND INCUBATION
Different metabolites were used in combinations to study their effects on biodegradation of polyethylene by the cave-bacteria consortia. Glucose, yeast extract, and calcium were used as co- metabolites. About 1% w/v of glucose and yeast extract were used, whereas the concentration of calcium in the medium was maintained at
0.03% to match the natural concentration of calcium of the environment where the soil was taken.
In total six combinations of these metabolites in mineral salt medium with polyethylene pieces and bacterial consortia were incubated at 150 rpm at pH 7.2 and temperature 37
°C for four weeks. A negative control was set by incubating polyethylene pieces in mineral salt medium with no bacterial inoculum.
BIODEGRADATION ANALYSIS
Biodegradation of polyethylene was analyzed by determining CFU mL−1, fourier- transform infrared spectra, and scanning electron microscope images. CFU mL−1 was determined after every week, whereas FTIR and SEM analysis were performed after one month of incubation. The viable cell count was done for bacterial growth determination through serial dilution and calculating CFU mL−1. Test LDPE samples were compared with the untreated control samples. FTIR (Jasco FT/ IR – 620) analysis was performed to check the degradation of LDPE pieces after being mixed with the growing bacterial consortia in liquid medium. This analysis detects any change in the functional groups. Spectrum was recorded at 500-4000 wave-numbers cm–1 for all the
LDPE samples. Surface morphology of LDPE pieces was observed by SEM (JSM 5910
Joel, Japan) to look for any change in structure or surface of LDPE piece after treating with microbial consortia. After rinsing of the LDPE pieces with autoclaved distilled water, LDPE pieces were mounted on the copper stubs with gold paint. Gold coating was carried out under vacuum by evaporation to make the samples conducting.
RESULTS
SOIL ANALYSIS
Atomic absorption spectroscopy was performed to determine the concentration of elements in the cave soil sample (Zada et al., 2016). Ca was 332.938 mg kg–1 as compared to 121.65 mg kg–1 in control soil, Mg was 1.2576 mg kg–1 in cave soil and
1.023 mg kg–1 in control soil. Ni 0.965 mg kg–1 in cave soil and 10.4 mg kg–1 in control soil, Cr 0.571 mg kg–1 in cave soil and 8.74 mg kg–1 in control soil, Co 0.266 mg kg–1 in cave soil and 0.810 mg kg–1 in control soil, Cu 1.824 mg kg–1 in cave soil and 4.7 mg kg–1 in control soil, Zn 12.7311 mg kg–1 in cave soil and 36.41 mg kg–1 in control soil,
and Pb 1.31 mg kg–1 in cave soil and 8.14 mg kg–1 in control soil were much lower than those found in the control soil (Zada et al., 2016).
VIABLE CELL COUNT
The concentration of viable cells in CFU/ml was determined at time zero, before initial incubation, and then after every week for a period of one month (Table 1). Since polyethylene in the medium was the sole carbon source, CFU mL–1 is directly proportional to the ability of organisms to degrade polyethylene and use it as a carbon source. There was a consistent decrease in CFU mL–1 after three and four weeks of incubation. The soil from where the bacteria were isolated contained exceptionally high concentration of calcium. Considering this high amount of calcium in the native habitat of the organism, extra calcium salt was added in the medium so that the organisms experience minimum deviation from their natural environment. Medium augmented with extra calcium showed increasing values of viable cell count in the first two weeks of incubation. Increase in the values of viable cell count was also observed in the first two weeks of incubation when the medium is augmented with glucose; these higher values of CFU mL–1 in first two weeks were because the bacteria were provided with glucose that acted as growth activator. Additional amounts of calcium proved to be helpful for better growth of bacterial colonies.
Table 3.2.1. Viable Cell Count of bacterial consortium in different media compositions incubated at 37 °C.
Time MSM + PE + MSM + PE + MSM + MSM + MSM + MSM + (days) bacterial calcium Glucose + PE glucose + glucose + calcium consortium carbonate + + bacterial calcium calcium carbonate + bacterial consortium carbonate + carbonate + yeast extract+ consortium PE + bacterial yeast extract + PE + bacterial consortium PE + bacterial consortium consortium CFU/ml (x105) CFU/ml (x105) CFU/ml (x105) CFU/ml (x105) CFU/ml (x105) CFU/ml (x105)
0 280 246 241 220 256 255 7 256 296 292 265 284 295
14 192 221 210 225 237 236 21 110 125 110 164 126 119
28 23 103 68 107 95 92
FOURIER TRANSFORM INFRA-RED SPECTROSCOPY
FTIR was carried out on LDPE films after incubation with bacterial consortia for four weeks. The peaks formed were compared with the control (Fig. 3.2.2).
LDPE in mineral salt medium containing bacterial consortia: Absorbance peaks formed at 2915 cm–1 and 2848 cm –1 suggest presence of C–H bonds. A peak of variable strength at 1472 cm–1 and
1462 cm–1 shows formation of C=C bonds. A strong peak at 1035 cm–1 shows formation of stretch of C–O bonds. Absorbance peaks formed at 615 cm–1, 718 cm–1 and 730 cm–1 show presence of
=C–H bending bonds.
LDPE in MSM containing calcium carbonate and bacterial consortia: New peaks were formed at
1645 cm–1 that represent formation of C=C, and a peak formed at 3442 cm–1 shows stretching of
O-H bonds.
LDPE in MSM containing glucose and bacterial consortia: Formation of new peak at 2978 cm–1 shows stretching of C–H bonds and peak at 3638 cm–1 represents of stretching of O–H bonds.
LDPE in MSM containing calcium carbonate, glucose, and bacterial consortia: Maximum variety of peaks was observed in this film. Formation of peaks at 3251 cm–1 and 3032 cm–1 shows formation of O–H bonds. A peak at 2915 cm–1 and 2848 cm–1 represents formation of stretch of C–
H bonds in the polyethylene. Peak formation at 2233 cm–1, 2178 cm–1, 2167 cm–1, 2103 cm–1, and
2013 cm–1 shows that new Nitrile bonds of –C ≡ N are formed. Absorbance peaks at 1462 cm–1 and 1472 cm–1 show bending of –C-H- bonds. Peak at 1084 cm–1 shows formation of stretch of C–
O functional group. Absorbance peaks formed at 615 cm–1, 718 cm–1, and 730 cm–1 shows presence of =C-H bending bonds.
A: Control (untreate d) sample
B: No additives
C: Added Calcium Carbonat e
D: Added Glucose
E: Added Calcium Carbonat e and Glucose
F: Added Calcium Carbonat e, Glucose and Yeast extract
G: Added Calcium Carbonat e and Yeast extract
Figure 3.2.2. Fourier-transform infrared spectra from control and different media after incubation at 37 °C for one month
LDPE in MSM containing calcium carbonate, glucose, yeast extract, and bacterial consortia: In comparison with control, new peak was formed in this medium combination at 1033 cm–1 represents formation of C–O bond.
LDPE in MSM containing calcium carbonate, yeast extract and bacterial consortia: In comparison with control, anew peak was formed in this medium combination at 1033 cm–1 that represents formation of C-O bond.
SCANNING ELECTRON MICROSCOPY
SEM showed adherence of bacteria that caused mechanical damage and erosion on the surface of plastic films incubated with bacterial consortia as compared to the untreated control (Fig. 3.2.3). More changes in surface topology and attachment of cells, despite the washing, were observed on the LDPE piece incubated in the presence of glucose and calcium.
A: Control (untreated) sample
.
B: No additives
C: Added Calcium Carbonate
D: Added Glucose
E: Added Calcium Carbonate and Glucose
F: Added Calcium Carbonate, Glucose and Yeast extract
G: Added Calcium Carbonate and Yeast extract
Figure. 3.2.3. A-G. Scanning electron microscopy of low-density polyethylene samples under specified treatment after incubation with bacterial consortia at 37 °C for one month.
DISCUSSION
There is an increasing interest in investigating biodegradation of non-degradable plastics using efficient microorganisms (Bonhomme et al., 2003; Boonchan et al., 2000; Lee et al., 1991). In our present study, bacterial isolates were obtained from the soil of Kashmir Smast, which is a limestone cave in Khyber Pakhtoonkhuwa province, Pakistan. The four isolates were identified as Serratia sp. KC1-MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3-MRL, and Stenotrophomonas sp. KC4-MRL. These strains were used in consortia to check for the ability of these microbes to degrade polyethylene. Since both antibiotic production and polyethylene degradation occur under stressed environmental conditions, the hypothesis of the current study was that bacteria having the first property are more likely to give positive result for the other. Studies on Brevibacillus borstelensis 707 showed an increase in potential of polyethylene biodegradation when grown on nitrogen-limit stressed cultures (Hadad et al., 2005). It was also observed that not only nitrogen deprivation, based on the amount of KNO3 in medium, but also carbon limitation in a mannitol- free medium alone enhanced degradation which was also enhanced when used in combination. It was observed that mannitol-free medium supplemented with nitrogen source showed maximum biodegradation in 30 days of incubation. In the present study, it was found that bacterial consortia showed higher viable cells measured as CFU mL–1 when the medium was supplemented with the nitrogen source.
Loss of tensile strength of plastic after incubation with Pseudomonas stutzeri suggests that the bacterium is capable of degrading the polymer (Sharma and Sharma, 2004). When bacteria are grown in different media compositions along with a polymer, maximum turbidity is observed on forty-fifth day (Ciferri, 1999). The increase in growth rate in glucose as well as in minimal medium suggests that the bacteria were completely depending upon polyethylene film for its source of carbon in the absence of glucose. CFU mL–1 increased from 101 to 103 from day 15 to day 30
(Table 1). These results suggest that the bacteria growing in Lascaux Cave, France are capable of using plastics and other resins such as glue as their sole source of carbon. Higher values of CFU/ml in the first two weeks were observed in all those media combinations in which 1% glucose w/v was added at the beginning. When the nutrients depleted, bacteria had polyethylene as the only carbon source. Addition of easily available substrate like glucose in MSM medium increases bacterial growth in initial stages of pesticide degradation (Cycoń et al., 2011). A similar effect on growth was observed when MSM media is augmented with yeast extract. The increased CFU mL–
1 was observed in first two weeks of the experiment and then the growth decreased, which indicates depletion of glucose in the medium.
In our study, fourier-transform infrared spectral analysis was carried out to check chemical degradation of polyethylene. Low- and high-density polyethylene are made of the elements carbon and hydrogen forming chains of repeating – CH2 – units (Rajandas et al., 2012). In the process of biodegradation of LDPE, enzymes catalyze a specific series of biochemical reactions that lead to various kinds of chemical conversions, such as oxidation, reduction, hydrolysis, esterification, and molecular inner conversion (Harshvardhan and Jha, 2013). Keto and ester carbonyls have been reported as major products in the presence of oxidoreductase (Karlsson and Albertsson, 1998).
Analysis of FTIR showed new peaks when LDPE is treated with bacterial consortia, indicating polymer breakdown and formation of new functional groups. The results of scanning electron microscopy have shown that all those media to which calcium was added showed strong biofilm development and hence increased biodegradation. It is also evident in several studies that bacteria release various surface-active substances extracellularly, which increase the bioavailability to the polymer. Studies on Pseudomonas sp. AKS2 showed that the strain was capable of degradation 5
± 1 % of initial LDPE in 45 days (Tribedi and Sil, 2013). The degradation by Pseudomonas
depends on its capability to colonize the surface of the polymer and degrade it. Addition of calcium increases biofilm development of Xylella fastidiosa under in vitro conditions (Cruz et al., 2012).
The efficient increase in formation of biofilm was observed when at least 1.0 mM CaCl2 was added in the medium. There was no effect of Ca on attachment when bacteria were treated with tetracycline, indicating that Ca has a regulatory role in colonization or attachment of the cells. In another study, Ehret and Böl (2013) showed that Ca ions crosslink alginates, which is the key constituent of the extracellular polymeric material produced by the mucoid P. aeruginosa strain to produce biofilms. Ciferri reported a list of bacteria responsible for degradation of paints (Ciferri,
1999).
In the present study, SEM showed discoloration, spots, erosion, and cracking on the surface of polyethylene film. Modification on the surface of polyethylene after bacterial treatment was also reported by (Matsunaga and Whitney, 2000). Formation of pits and erosion on the surface of LDPE when observed through electron microscopy when incubated with Fusarium sp. indicating adherence and degradation of LDPE (Hasan et al., 2007).
Bacteria capable of adhering to plastic surfaces, growing, and possibly degrading it by oxidation are commonly present in soil. Microorganisms that can adhere to the surface of pre-oxidized PE are also commonly present in soil. If the pre-oxidant technology is commonly employed in the PE manufacturing process, one can expect that these plastics will be able to degrade in waste-disposal sites. In the current study, structural and surface changes in PE in the form of depression, pits, and erosions were visible in SEM. Physical erosion of the surface of polyethylene observed through
SEM by fungi has been reported by Bonhomme et al. (2003). Polymer treated with microorganism loses its physical strength and disintegrates on applying mild pressure. Wide-spread pits and holes in polycaprolactone surface are reported by Shaw et al. (2015) after ten days of incubation with
thermophilic bacterium Ralstonia sp. strain MRL-TL. SEM has shown that biofilms develop on the surface of polyethylene with time by PE-degrading bacteria. It is known that formation of biofilm on the surface of plastics favors adhesion of bacteria on the surface and helps them survive under low-nutrient conditions and to use polyethylene as their source of carbon (Linos et al., 2000).
CONCLUSIONS
Our study indicates that antibiotic-producing bacteria in consortia isolated from a limestone cave could degrade the synthetic polymer polyethylene. Maximum biodegradation was observed when the medium was augmented with calcium salt, indicating higher degradation potential of bacterial consortia when in a medium close to the natural chemical composition of their native environment.
Spectroscopy and microscopy results showed certain changes in low-density polyethylene test samples as compared to a control, indicating microbial breakdown of LDPE. Further research is needed to understand the mechanism of degradation of LDPE at a molecular level. All the bacterial strains were found to viable at the end of the experiment.
ACKNOWLEDGEMENTS
We are thankful to Quaid-i-Azam University, Islamabad, for providing funds to accomplish this research.
Lysinibacillus sphaericus AB-8 isolated from a limestone cave (Kashmir Smast) Pirsai Mardan, Pakistan, and its ability to produce industrially important enzymes Abstract In the present study, a bacterial isolate, KC5- MRL isolated from Kashmir Smast (cave), a limestone cave in Pirsai Mardan, KPK, Pakistan, was found to produce three industrially important enzymes; lipase, protease and amylase. The bacterial isolate was identified as Lysinibacillus sphaericus KC5-MRL (Accession No. KF010827). Optimum pH for the growth of Lysinibacillus sphaericus KC5- MRL, was around 7 and grew best at 35ºC. The optimum activity of lipase was observed at 30°C after 24 hr of incubation and pH 5 (42.23 U/ml). Maximum lypolytic activity (181.93 U/ml) was observed when 8% inoculum was used. Amylolytic activity of Lysinibacillus sphaericus KC5-MRL was optimum (15 U/ml) after 24 hr of incubation at 30°C. Proteolytic activity of L. sphaericus KC5-MRL was found to be 59 U/ml, after 48 hr at 30°C. Highest stability (42%) of lipase was observed at pH 10. pH stability of amylase showed highest activity at pH 7 i.e. 99.4%, whereas, protease stability was highest at pH 8. L. sphaericus KC5-MRL lipase, protease and amylase were stable at 35ºC and with residual activity as 118%, 104% and 107%, respectively. Triton X-100 and sodium dodecyl sulfate (SDS) stimulated the lipase and protease
++ + ++ activities, whereas, Triton X-100 and T-80 stimulated amylolytic activity. Mg , NH4 and Ca stimulated the lipase activity and Zn++ showed highest inhibitory effect on lipase activity. Hg+, ++ ++ + + ++ Mg , Zn and NH4 reduced amylase activity, whereas, Na and Ca showed stimulatory effect. + ++ ++ + + + Hg , Zn , Ca and NH4 reduced protease activity but Na and Mg stimulated protease activity. Chloroform, formaldehyde, methanol and benzene stimulated amylase activity. Nitobenzene, methanol, benzene and acetone stimulated protease activity. Ethylenediamine tetraacetic acid (EDTA) showed stimulatory effect on lipase. EDTA, Trisodium citrate TSC, mercaptoethanol, Phenyl acetaldehyde (PAA) and PMSF reduced amylase activity. All the modulators reduced protease activity except TSC and PMSF which stimulated its activity. The study concludes that these enzymes can be used for different purposes in various industries such as food and detergent. Keywords: Lysinibacillus sphaericus KC5- MRL, Kashmir Smast, lipase, limestone, Pakistan
INTRODUCTION
Caves are known to have very poor nutrient ecology and have constant low temperature and high humidity. Because of this environment, the cave organisms have some extraordinary properties to survive and live in such hard and crucial condition (Northup et al., 2009; Engel et al., 2004; Simmons et al., 2008). Cave ecology is rich in different types of microorganisms having outstanding characteristics. The most abundant organisms observed in caves are filamentous and belong to Actinobacteria group, followed by coccoid and bacilli forms (Cuezva et al., 2009). A few pathogenic microorganisms have been reported from the Altamira cave (Jurado et al., 2006). Luong et al. (2010) described for the first time, A. altamirensis recovered from human medical constituents. E. coli and S. aureus which are disease causing bacteria have also been isolated from caves (Lavoie et al., 2005), species of Pseudomonas, Sphingomonas and Alcaligenes sp. (Ikner et al., 2007) and Inquilinu sp. (Laiz et al. 1999). Nutrient recycling in the cave totally depends upon the bacterial and fungal activities (Prabhavathi et al., 2012).
Caves have fewer amounts of nutrients, less light or no light ecosystem, which have relatively low temperature and high humidity. From different caves, around the world, different types of microbial species have been isolated (Tomova et al., 2013).
Most of the caves are valuable, as it contains some historical paintings which are affected by progressive microbial colonization and biodeterioration and how to handle this worldwide problem (Allemand and Bahn, 2005; Cañaveras et al., 2001; Fox, 2008). Most of the caves which have been studied are in Spain, Italy, France, Romania and USA. Altamira Cave in Spain and Lascaux Cave of France is the most microbiologically studied cave (Bastian et al., 2009; Schabereiter et al., 2002a).
Significant growth in biospeleological research has been observed in the last two decades (Urzì et al., 2010). Caves are considered as extreme environments providing niches for highly specialized microorganisms (Schabereiter et al., 2002b). Enzymes which are isolated from cave microbes are active at low temperature. Enzymes of psychrotrophs have great biotechnological importance (Tomova et al., 2013). These microbes survive in such environment because of production of enzymes which function in such extreme conditions (Burg, 2003). Our knowledge about cave
microbial diversity is limited, so there is great potential to find novel microorganisms in caves (Engel, 2010).
Caves in Pakistan are less studied area especially with regards to industrially important enzymes from cave bacteria have not been reported or isolated so for. Therefore, in our study we focused, on the isolation of bacteria from Kashmir cave, and their potential to produce industrially important enzymes having some unique properties.
MATERIALS AND METHODS
The present study was carried out in the Microbiology Research Laboratory (MRL), Department of Microbiology, Quaid-i-Azam University, Islamabad, Pakistan. Production of lipase, amylase and protease by Lysinibacillus sphaericus KC5-MRL, isolated from a limestone cave (Kashmir Smast Pirsai Mardan, Pakistan) (Fig. 1) was investigated. The cave (GPS co-ordinates 34° 25' 48.24" N 72° 13' 43.18" E) is 188 m long, 30 m high and 25 m wide, is located in Pirsai Mardan, Pakistan (Fig. 3.3.1). Soil samples were collected in sterile bags under aseptic conditions.
Fig. 3.3.1. Limestone cave (Kashmir Smast Pirsai Mardan, Pakistan)
Qualitative tests for three enzymes
Protease, Lipase and Amylase
Nutrient agar plates containing (1%) casein as substrate were used to check the proteolytic activity which was observed in the form of clear zones of hydrolysis around the colonies. For the screening of lipase, Tween 80 (1%) was used as lipid substrate. After incubation at 37°C for 24 h, appearance of clear zones was observed (Hasan and Hameed, 2001). Starch was used as substrate to detect the production of amylolytic enzyme. Inoculated plates were incubated at 37°C for 24 h and clear zones of hydrolysis were observed.
Molecular characterization
A bacterial isolate having the ability to produce the three enzymes (lipase, protease and amylase) was identified on the basis of 16S rRNA gene sequencing.
Sequence Analysis
The sequencing of 16S rRNA gene was performed through Macrogen service, Seoul Korea. Different bioinformatics tools were used to analyze the sequences i.e. BLAST and FASTA used for sequenced data observation, alignment of the sequences was performed by using MEGA 5.0 and CLUSTAL W.
Quantitative analysis of lipase, protease and amylase
About 150 ml of medium was added in four different Erlenmeyer flasks, autoclaved for 20 minutes at 121°C and 15 lbs. About 1.5 ml of olive oil was added to the above medium which was already sterilized in oven at 121°C for half an hour. Then medium was inoculated with two loopful of Lysinibacillus sphaericus KC5-MRL (KF010827) culture under sterilized conditions, then flasks were kept in shaking incubator at 37°C and 150 rpm. Samples were drawn after 0, 24, 48 and 72 h. Enzyme assay for lipase was performed by the method of (Lesuisse et al., 1993). Unit of enzyme is defined as the amount of enzyme that hydrolyzes 1 μmol substrate in 1 minute. For protease assay (Kunitz, 1965) method was used and unit of protease enzymes were defined as that amount of enzyme which releases 1 μmol tyrosine under standard conditions of assay method. For amylase
enzyme assay (Bernfeld, 1951) method was used and unit of amylase were defined as the amount of enzyme in 10 ml of filtrate, which releases 1 mg of reducing sugar from 1% starch solution in 1 hr at 37ºC and pH 7.
Shake flask fermentation for optimization of different parameters
To determine the optimum pH for lipase production, fermentation was carried out at different pH (3, 4, 5, 6, 7, 8 and 9) and optimum temperature was determined by carrying out shake flask fermentation at various temperatures (15, 25, 30, 35 and 40°C).
Size of inoculum
A 24 h old bacterial culture grown in Nutrient broth, was added at a concentration of 1 to10% to the production medium. About 150 ml of production medium was poured into four different flasks. Samples were taken after 0, 24, 48 and 72 hr and assayed for lipase activity, and centrifuged at 10,000 rpm for 30 minutes at 4°C. Precipitates were removed by filtration and supernatant was used as the crude enzyme for the estimation of enzyme activity.
Effect of temperature on enzyme activity
Activity of crude enzyme was determined after incubating for 1 hr at 30, 35 40, 50, 60, 70 and 80 and 100˚C.
Effect of pH on enzyme activity
Effect of pH on the activity of crude enzyme extract was studied at different pH. The crude enzyme was incubated for 1 h in buffers having different pH [0.02 M of acetate buffer (pH 4, 5, 6), phosphate buffer (pH 6, 7), Tris-HCl (pH 8), glycine-NaOH buffer (pH 9) and Na2HPO4/NaOH (pH 10)], and remaining enzyme activity was determined under standard conditions.
Effect of metal ions on enzyme activity
Effect of different metals [100 mM of HgCl2, NaCl, MgCl2, ZnCl2, (NH4)2SO4 and CaCl2 separately], on activity of enzymes was determined. These metals were pre-incubated with crude extract (1:1) for 1 hr at room temperature and then remaining activity was determined.
Effect of modulators on enzyme activity
Effect of different modulators on enzyme activity was determined for EDTA, Trisodium citrate, mercaptoethanol and PMSF. About 1% of these compounds were pre-incubated for 1 h at room temperature (25°C) with crude enzyme (1:1).
Effect of solvents on enzyme activity
Effect of different solvents on activity of lipase was determined after pre-incubating crude enzyme in the presence of (1% each, separately) xylene, benzene, chloroform, nitrobenzene, formaldehyde, methanol and acetone (1:1) for 1 h at room temperature.
Protein estimation
For protein estimation, Lowry’s method (Lowry, 1951) was used.
RESULTS AND DISCUSSION
Enzymes are known as nature’s catalysts (Louwier et al., 1998). So far different enzymes are isolated from different living organisms like plants, animals, fungi, yeast and bacteria (Wiseman, 1995). Throughout the world only 2% enzymes are isolated from microorganisms (Frost et al., 1987). These microorganisms are isolated from different ecosystems. The hidden potential of microorganisms in sense of valuable industrial enzymes exploration is still on wait. These enzymes have number of applications in different industries like food, medicine, cosmetics, textile, leather, detergents, oleochemical and for different diagnostic tools (Hasan et al., 2006). Caves are extreme environment with poor nutrient content. Caves still have to be explored for new horizons in Microspeleology sectors. In the world different antibiotics are isolated from different caves. From ice caves different types of industrially important enzymes have also been reported. Our present study is focused on caves bacteria to find out industrially important enzymes. Lysinibacillus sphaericus is a gram positive, rod shaped mesophilic bacterium that naturally occur in soil.
Sequence Analysis The bacterial isolate was identified as Lysinibacillus sphaericus KC5-MRL (Accession No. KF010827) (Fig. 3.3.2).
43 Lysinibacillus sphaericus(JN377786) 64 Lysinibacillus sphaericus(JN377784) Fig. 11 Lysinibacillus sphaericus(JN377785) 2 Lysinibacillus fusiformis(FJ418643) Lysinibacillus sphaericus(GU204967) 2 8 Lysinibacillus sp(HM222673) Lysinibacillus sp(AM910304) 14 Lysinibacillus sp(AB689752) 85 12 31 Lysinibacillus sp(JN695724) Lysinibacillus sp(JX566617) 20 Lysinibacillus sphaericus-(EU880531) Lysinibacillus sphaericus(CP000817) Lysinibacillus sphaericus(FJ544252) Lysinibacillus sphaericus KC5-MRL (KF010827) 3.3.2. Evolutionary relationships of taxa Lysinibacillus sphaericus KC5-MRL
Qualitative Tests for enzymes (protease, amylase and lipase) Lysinibacillus sphaericus KC5-MRL was positive for all the enzymes with largest zone of hydrolysis of substrates.
Effect of pH, temperature, size of inoculum on growth of Lysinibacillus sphaericus KC5- MRL Growth of L. sphaericus KC5-MRL was optimum at neutral pH 7, with optimum growth at 35°C and almost no growth at temperature 15 and 40°C (Fig. 3.3.3), and size of inoculum 8% after 72 h (Fig. 3.3.3).
Fig. 3.3.3. Effect of size of inoculum on the growth of Lysinibacillus sphaericus KC5-MRL and production of lipase
Effect of pH, temperature and size of inoculum for the optimum lipolytic activity
The optimum activity of L. sphaericus KC5-MRL lipase was observed at 30°C after 24 hr of incubation and pH 5 i.e. 42.23 U/ml (Fig. 3.3.5). Maximum lipolytic activity (181.93 U/ml) was observed when 8% inoculum was used (Fig. 3.3.3). Lysinibacillus sphaericus KC5-MRL showed maximum lipase production 23 U/ml at 30°C and pH 5 after 24 h of incubation. Similar specific enzyme activity was reported by different investigators for lipase (Hasan and Hameed, 2001; Ankit et al., 2011; Ozgur and Nilufer, 2012). Lipase from Pseudomonas sp., that produced maximum lipase after 24 h of incubation at 30-40°C (Kathiravan et al., 2012). Pseudomonas sp., produced maximum lipase at pH 5.5 (Kavitha and Shanthi, 2013). Variation in pH, incubation time and incubation temperature may be due to isolate specificity for different conditions. Maximum lipase activity was observed at a size of 7% inoculum. Balan et al. (2010) reported maximum lipase production when 7% inoculum of Geobacillus thermodenitrificans was used in flask fermentation.
Fig. 3.3.4. Lipolytic, amylolytic and proteolytic activity of Lysinibacillus sphaericus KC5- MRL at 30°C
Enzyme assay for amylase and protease The amylolytic activity of Lysinibacillus sphaericus KC5-MRL was optimum, 15 U/ml, after 24 h of incubation at 30°C, whereas it decreased much after 72 h i.e. 4 U/ml (Fig. 3.3.4). Proteolytic activity of L. sphaericus KC5-MRL was calculated to be 59 U/ml, after 48 h of incubation at 30°C, and decreased after 24 h i.e. 26 U/ml (Fig. 4). Amylase activity and incubation period were noted between 9-16 U/ml and 24-72hrs. Demirkan, (2010) isolated amylase from Bacillus subtilis and its mutant have the some incubation periods but our amylase units is in contradict. Our strains amylase activity is less from his isolated amylase. The protease activity and incubation period were noted between 40-75 U/ml and 24-72 hrs. Different investigators (Mayerhofer et al., 1973; Patil et al., 2011) have been reported that protease is produced during stationary phase. They isolated these proteases from Pseudomonas fluorescens P26 and Bacillus spp. Amro et al. (2009) isolated bacterial proteases from different bacteria which have maximum protease activity in the range of 34-44 U/ml. In our research the proteases have high activity from them.
Characterization of crude lipase, amylase and protease produced by L. sphaericus KC5- MRL
Stability of crude enzymes at different pH
Lipase was not stable at pH 4. Highest stability of lipase was observed at pH 10 i.e. 42%, followed by pH 6 with lipase stability as 22%. At pH 5, 7, 8 and at pH 9, lipase activity reduced to 11.4%, 24.5%, 4.5% and 9.8% respectively. pH stability results showed that highest amylolytic activity was observed at pH 7 i.e. 99.4%. Lowest amylase activity was observed at pH 4 and 5, as 9.9 and 31.4%, respectively. Protease produced by L. sphaericus KC5-MRL showed lowest stability at pH 10 i.e. 92%. Highest activity was noted at pH i.e. 115%. L. sphaericus KC5-MRL protease was found stable at all pH values (Fig. 3.3.5). Mobarak et al. (2011) reported that lipase of Pseudomonas aeruginosa KM110 showed optimum activity at pH 8 i.e. 27%. Our isolated lipase also showed 142% activity at alkaline pH 10. This was maximum activity for the pH value tested. Kojima et al. (1994) isolated bacterial lipase from Pseudomonas fluorescens AK102 which were stable at pH 4-10 and maximum stability was noted at pH 8-10. All the amylase of the four strains showed stability from 6-10 pH value. It’s showed that these amylase work best it alkaline pH. Bacillus subtilis AB-22 amylase was stimulated by pH 10. A sharp decline was noted at a pH 4 and 5.Our results are similar with Krishnan and Chandra (1983) isolated amylase from Bacillus licheniformis CUMC305 which have maximum activity and stability at pH 9. But the amylase was active at a wide range of pH from 3-10. Most of the protease in our study was noted to be stable from 4-10 pH values. Stimulation of activity observed at alkaline pH. Only one protease Bacillus subtilis AB-3 showed highest activity at pH 7 ie 127%. The remaining 3 proteases were showed maximum activity at pH 8-10. Our results are similar to that of Adinarayana et al., (2003) who isolated protease from Bacillus spp. BZI-2 and Femi-Ola and Oladokun, (2012) from Lactobacillus brevis.
Fig. 3.3.5. Stability of crude extracts of lipase, amylase and protease of Lysinibacillus sphaericus KC5-MRL at different pH
Stability of crude enzymes at different temperature
L. sphaericus KC5-MRL lipase highest (118%) stability was observed at 35°C, lowest stability (64%) was observed at 80°C. The highest temperature stability of L. sphaericus KC5-MRL amylase was observed at 35ºC i.e. 107%. Almost no amylolytic activity was recorded at 100ºC. The highest temperature stability of L. sphaericus KC5-MRL protease was observed at 35ºC i.e. 104%, whereas, lowest stability was reported at 80ºC and 100ºC i.e. 70% and 31%, respectively (Fig. 3.3.6). Lipase retained its activity at all temperatures tested. The highest activity was observed at 35°C. The enzymes were stable at 30-80°C. Wang et al. (1995) characterized a bacterial lipase of Bacillus strain A30-1 (ATCC 53841) which was stable at 75°C. Rathi et al. (2000) isolated a lipase from Pseudomonas sp., which was stable at 90°C for 3 h. In our research, lipase was stable for 1 h at 80°C. Demirkan. (2010) isolated amylase from Bacillus subtilis reported highest activity from the current study. Different investigators (Mayerhofer et al., 1973; Patil et al., 2011) have reported that protease is produced during stationary phase. They isolated these proteases from Pseudomonas fluorescens P26 and Bacillus spp. Amro et al. (2009) isolated bacterial proteases from different bacteria which have maximum protease activity in the range of 34-44 U/ml. In our study we reported 59 U/ml proteases activity. Overall, the isolated amylases in our study were stable from 30 to 80ºC for 1 h. Malhotra et al. (2000) reported α-amylase of Bacillus thermooleovorans NP54 which was active at 100ºC for 10 minutes. The enzymes they reported were stable at 40-100ºC. Our results are similar to that of Adinarayana et al. (2003). They isolated protease from Bacillus sp. BZI-2 and Femi-Ola and Oladokun (2012) isolated protease from Lactobacillus brevis. Pathak and Deshmukh. (2012) reported a protease from Bacillus licheniformis which was active in the range of 20-90ºC. Lee et al. (2000) isolated a protease from Pseudoalteromonas sp. A28, which is in accordance with our study.
Effect of detergents on lipolytic, amylolytic and proteolytic activity of Lysinibacillus sphaericus KC5-MRL Addition of Tween-80 reduced liypolytic activity up to 4%. Triton X-100 and sodium dodecyl sulfate (SDS) stimulated the lipase activity up to 4% and 3%, respectively. SDS reduced amylase activity of L. sphaericus KC5-MRL activity up to 70%. Trition X-100 and T-80 stimulated amylolytic activity as 19% and 23%, respectively. Protease activity was reduced (62.3%) by addition of T-80. Triton X-100 and T-80 stimulated protease activity up to 9% and 23%, respectively (Fig. 3.3.7). Lee et al. (1999) reported that different detergents change the activity of
Bacillus thermoleovorans ID-1 thermophilic lipase. They reported no activity of Triton X-100 but in contrast to our study, it stimulated the lipase activity and SDS also stimulated enzyme activity. Bacillus thermoleovorans ID-1 lipase activity were reduced by Tween 80. None of the surfactants tested had a pronounced inhibitory effect on amylase activities. Oliveira et al. (2010) reported amylase from Rhizobia strains. Amylase activity was not inhibited by detergents/surfactants which they used in their experiments. They reported that Triton X-100 moderately inhibited enzyme activity and SDS and Tween-80 showed stimulatory effect on the enzyme activity. Similar results were noted in the present study. Addition of surfactants in case of proteases showed some stimulatory and as well as inhibitory effect. Nascimento and Martins (2006) isolated protease from Bacillus sp., which was inhibited by the addition of surfactants like SDS and Triton X-100. This result is similar to the present study.
Fig. 3.3.6. Stability of crude extracts of lipase, amylase and protease of Lysinibacillus sphaericus KC5-MRL at different temperature
Effect of metal ions on lipase, amylase and protease activity ++ + ++ The metal ions e.g. Mg , NH4 and Ca stimulated lipase activity as; 25.8%, 31.1% and 27.54%, respectively. Ammonium ion stimulated the lipase activity up to 31.7%. Hg+, Na+ and Zn++ inhibited lipase activity as 2%, 7% and 11.31%, respectively. Zn++ showed highest inhibitory effect on lipase activity. Some metals were inhibitory while others stimulated amylase of L. sphaericus
+ ++ ++ + KC5-MRL. Hg , Mg , Zn and NH4 reduced amylase activity as 69, 11, 26 and 55%
respectively. Presence of Na+ and Ca++ showed stimulatory effect up to 11 and 6%, respectively. Some metals were inhibitory for protease of L. sphaericus KC5-MRL (KF010827). Hg+, Zn++,
++ + + + Ca and NH4 reduced protease activity 57, 46, 47 and 15% respectively. Na and Mg stimulated protease activity up to 8 and 16%, respectively (Fig. 3.3.8). Effect of different metal ions was
++ + ++ checked on the basis of residual activity of crude extract. Mg , NH4 and Ca stimulated lipase activity. Stimulation of crude extract activity have been reported by different investigators Li and Ziaobo. (2005) reported that Ca++ and Mg++ stimulate the activity of Geobacillus sp. TW1 lipase. Our results are similar to that of Mobarak et al. (2011), where Pseudomonas aeruginosa KM110 lipase was inhibited by Zn++ and Cu++ (Mobarak et al., 2011). Some metals were found inhibitory and some stimulatory for the Lysinibacillus sphaericus KC5-MRL amylase. Hailemariam et al. (2013) also reported amylase from a Bacillus sp. which was inhibited by different metals ions. Activity of amylase from different bacterial isolates was reduced by different divalent cations (Najaf 2005; Goyal et al., 2005; Ramesh et al., 1990; Koch et al., 1991; Božić et al., 2011; Mamo et al., 1999). All metals were inhibitory for the protease. Banerjee et al.
(1999) reported Brevibacillus (Bacillus) brevis to be inhibited by CuSO4, ZnCl2 and HgCl2. Usharani and Muthuraj. (2010) reported Bacillus laterosporus protease whose activity was inhibited by the presence of different metals ions. The inhibitory effect of heavy metal ions is well documented in the literature. In the present study most of the metal ions inhibited the enzyme activity.
Fig. 3.3.7. Effects of detergents on the lipolytic, amylolytic and proteolytic activity of isolate Lysinibacillus sphaericus KC5-MRL Effect of organic solvents on lipase, amylase and protease activity
Presence of nitro-benzene (N.B) reduced lipase activity up to 3.3%. Other organic solvents stimulated lipase activity. Highest stimulation was observed in case of benzene (B) 14.32%. Addition of xylene (X) showed no effect on the activity of lipase while chloroform (Ch), formaldehyde (Frm), methanol (Met) and acetone (Ace), stimulated lipase activity as 2.5%, 10.5%, 4.2% and 3%, respectively. Chloroform, formaldehyde, methanol and benzene stimulated amylase activity up to 9, 4, 3.5, 2 and 7%, respectively. Nitro- benzene, xylene and acetone reduced amylase activity up to 33, 5 and 50%, respectively. N.B, methanol, benzene and acetone stimulated L. sphaericus KC5-MRL protease activity up to 17, 17, 4 and 27%, respectively. Formaldehyde, chloroform and xylene reduced protease activity up to 10, 66 and 30%, respectively (Fig. 3.3.9). Organic solvents mostly showed some kind of stimulatory effects. Highest stimulation noted in case of benzene while nitrobenzene was inhibitory for enzyme. Our results are similar with that of Nawani et al. (2006). They reported benzene as stimulator for the lipase of thermophilic Bacillus sp. Schmidt et al. (1994) reported that different organic solvents like methanol, acetone and ethanol enhanced lipase activity isolated from B. thermocatenulatus. Prakash et al. (2009) reported two different types of bacterial amylases whose activities were inhibited by chloroform and other organic solvents while alcohol and acetone stimulated enzyme activity. Similar observation was reported in our study. Organic solvents showed stimulatory and as well as inhibitory effects in case of proteases. Rahman et al. (2006) isolated protease from Pseudomonas aeruginosa strain K whose activity was inhibited by 1-pentanol, benzene, toluene, p-xylene and n-hexane up to 33%, 35%, 30%, 48% and 36% respectively.
Fig. 3.3.8. Effect of metal ions on lipase, amylase and protease activity of isolate Lysinibacillus sphaericus KC5-MRL
Effect of modulators on lipase, amylase and protease activity Phenyl acetaldehyde (PAA) inhibited the activity of crude extract of lipase up to 17.3%. Ethylenediamine Tetraacetic acid (EDTA) stimulated lipolytic activity up to 24.4%. Mercaptoethanol (Mre) showed no effect on lipolytic activity, whereas, Tri-sodium citrate (TSC) and phenylmethylsulfonylfluoride (PMSF) both inhibited lipase activity up to 14%. All the modulators reduced amylase activity. EDTA, TSC, mercaptoethanol, PAA and PMSF reduced L. sphaericus KC5-MRL amylase activity up to 18, 84, 1, and 16%, respectively. All the modulators reduced protease activity except TSC and PMSF which stimulated protease activity up to 24 and 5%, respectively. EDTA, mercaptoethanol and PAA reduced protease activity up to 34, 37 and 26%, respectively (Fig.3.3.10). Ballschmiter et al. (2006) reported an amylase from hyperthermophilic bacteria with reduced activity in the presence of EDTA. They reported that amylase activity was slightly stimulated by the presence of mercaptoethanol which contradicts with our results. Lysinibacillus sphaericus KC5-MR showed that this protease belong to metalo proteases. Similar results also reported by Mabrouk et al. (1999). They isolated protease from Bacillus licheniformis ATCC 21415, which was inhibited by EDTA.
Fig. 3.3.9. Effects of organic solvents on lipase, amylase and protease activity of Lysinibacillus sphaericus KC5-MRL
Fig. 3.3.10. Effect of inhibitors on lipase, amylase and protease activity of Lysinibacillus sphaericus KC5-MRL
Conclusions Bacteria isolated from cave environment were able to produced enzymes (Lipase, protease, amylase). Lysinibacillus sphaericus KC5-MRL has shown varying activities of the three enzymes. Different characteristics of enzymes indicated their possible use in biotechnology and industry.
Bio-mineralization of CaCO3 by bacterial strains isolated from Kashmir Cave. Buner
Abstract Caves are underground natural compartments and hosts diverse microbial communities. Cave bacterial strains make significant contribution in the precipitation of calcium carbonate (CaCO3).
In the present study, it is shown that the CaCO3 precipitation is due to result of microbial metabolic activities. The bacterial strains were isolated and purified from Kashmir Cave Mardan Khyber
Pukhtunkhwa (K.P.K) soil. B4 medium was used in the whole research for the CaCO3 precipitation. A total of three bacterial strains showed the capability of CaCO3 precipitation on the selected medium. Viable bacterial count was done by determining colony forming unit (CFU) of samples observed bacterial load of 4.6×104 per gram of cave soil. Bacterial cells with mineralization potential were molecularly identified through 16S rRNA gene sequencing as Bacillus toyonensis, Paracoccus Limosus and Brevundimonas diminuta The most precipitates o were observed at temperature and pH of 25 C and 5. The precipitated CaCO3 was further confirmed by Scanning Electron Microscopy (SEM), X-ray powder diffraction (X-RD), and Fourier Transform Infra Red spectroscopy (FTIR) analysis.
Introduction Microbial carbonate precipitation has emerged as a promising technology for remediation and restoration of concrete structures. A number of diverse microbial species participate in the carbonate precipitation in different natural environment as in soils, in geological formation (caves), saline CaCO3 and ocean (Bharathi, 2014). Bacteria are capable of performing metabolic activities which thereby promote precipitation of calcium carbonate in the form of calcite (Bansal et al.,
2016). Over recent years, the implementation of microbially produced calcium carbonate (CaCO3) in different industrial and environmental applications has become an alternative for conventional approaches to induce CaCO3 precipitation. However, there are many factors affecting the bio- mineralization of CaCO3, which may restrict its application (Seifan, 2017). It is widely known that microorganisms contribute to the generation of a wide diversity of minerals such as carbonates, phosphates, sulfides, and silicates. Among all bio- precipitated minerals, the production of CaCO3
has drawn much attention due to its role in environmental and industrial applications. The bio- precipitation of CaCO3 can be achieved through biologically controlled mineralization (BCM) and biologically induced mineralization (BIM) (Wei et al., 2015).
There are different hypotheses for bacterial production of carbonate. The first hypothesis is ionic exchange through the bacterial cell membrane (Castainer et al., 2000). In this approach, which is considered as an active precipitation, microbial CaCO3 precipitation is induced by successful attachment of bacterial cell walls and positively charged Ca2+ ions. The production of extracellular polymeric substances (EPS) is assumed as another hypothesis in regard to CaCO3 precipitation through the trapping of Ca2+ (Kremer et al., 2008). The precipitation of carbonates is governed mainly by four factors: (1) calcium concentration, (2) carbonate concentration, (3) pH of the environment and (4) presence of nucleation sites (Hammes and Verstraete, 2002). Calcium carbonate precipitation is a biological as well as a geochemical method of producing carbonates of calcium with help of microorganisms specially those residing inside cave soil (Mortensen et al., 2011). This biological process carry out mineral precipitation, which holds the different soil particles packed in one structure and may also enhance the stiffness ability of the soil. Cave microbes can enzymatically carry out the biological reactions to crystalize calcium carbonate in soil (Fujita et al., 2000). Precipitation of calcium carbonate by ureolytic bacteria is one such mechanism where urea is hydrolysed into ammonium and bicarbonate. The Ca2+ ions subsequently 2− react with the CO3 ions, leading to the precipitation of CaCO3 at the cell surface that serves as a nucleation site. Precipitation of carbonates by carbonic anhydrase (CA) is another mechanism (Dhami et al., 2014). This enzyme has been found to have the most potential biological catalyst for hydration of CO2 leading to formation of CaCO3 in presence of calcium source (Li et al., 2010). Several genera of halophilic bacteria have been reported to precipitate carbonates in natural marine habitats, which include Halomonas, Deleya, Flavobacterium, Acinetobacterand Salinivibrio (Ferrer et al., 1988; Rivadeneyra, 1993; Rivadeneyra et al., 2006). These bacteria have the potential to grow in wide range of osmotic concentrations, which makes them very useful for studying the effect of different salt concentrations on carbonate precipitation efficacy. Soil residing bacteria can boost up calcium carbonate precipitation by promoting the alkaline conditions in cave soil (Kohnhauser et al., 2007), this is achieved by many biogeochemical processes like nitrate, sulfate, iron reduction, and break down of urea (DeJong et al., 2010). Cave bacteria ensure
dissolution and precipitation reactions that comprise carbonates, clays, silicates, manganese, iron, sulfur, and formation of speleothems (Northup and Lavoie et al., 2001).
Bio-mineralization of calcium carbonate by cave microbes is a complex dynamic process that may be affected by many factors including pH, temperature and incubation time. Therefore, the main aims of the present study were to investigate (i) the effect of temperature on bacterial growth and
CaCO3 precipitation, (ii) time of incubation in correlation to bio-mineralization of calcium carbonate and (iii) the performance of bacteria to induce CaCO3 precipitation at different ranges of pH by bacterial strains isolated from Kashmir Cave, Buner, KPK, Pakistan.
Materials and Methods
All the chemicals and reagents utilized in this study were of analytical grade and obtained from Sigma-Aldrich Chemical Co.
Sampling Soil samples were collected from the dark zone aseptically in sterile polythene zipper bags from Kashmir Smast (cave) situated at District Buner, Nanseer Buner, Khyber Pakhtunkhwa. (GPS coordinates 34o25’42.12”N 72o13’10.82”E), Pakistan. Temperature and pH of the sampling site was recorded. Samples were carefully transported on ice to Microbiology Research Laboratory (MRL), Quaid-I-Azam University, Islamabad and stored at 4±0.5°C for further study. The Kashmir caves are a series of natural limestone caves, located in the Babozai Mountains between Mardan and Buner districts in Northern Pakistan. Details given in reports by Ziad (2006) and Zada et al., (2016).
Atomic absorption analysis of sample A B Atomic absorption spectroscopy (AA240FS Fast Sequential Atomic Absorption Spectrophotometer) was carried out for quantitative analysis of elements in the soil sample. Sample was analysed in triplicate and mean value of absorbance was used to determine metal concentration. Soil digestion procedure was performed by grinding 1 g of soil sample and then it was mixed with 15 mL aqua regia and heated at 150°C and left overnight, and added C D
HClO4 and again heated at 150°C. The solution became almost dry until brown fumes came out. Whatman filter paper (No. 42) was used for filtration and the volume was raised up to 50 ml along with double distilled water (FAO/SIDA, 1983). Ordinary garden soil was run in parallel as a control.
Isolation and enumeration of bacteria Standard protocol of serial dilution was carried out for isolation and enumeration of bacteria from soil sample. About 100 μL of each dilution was transferred and spread on nutrient agar plates and incubated at 37°C for 48 hours. The plates were checked for bacterial growth and viable cell count (CFU/g) was calculated. Different colonies were marked and sub-cultured separately to obtain pure colonies and later preserved in 30% glycerol at -20°C.
Identification of bacterial isolates Isolates were presumptively characterized by morphological and microscopic analysis. All the isolates were also subjected to biochemical characterization (Oxidase, Catalase, Citrate, Urease, TSI, Casein, Amylase and Gelatinase tests) according to Bergey’s Manual of Determinative Bacteriology (Holt et al., 2012). Molecular identification of all the isolates was carried out by sequencing the 16S rRNA genes.
DNA Isolation DNA extraction of bacterial cell was carried out by centrifugation of 1 mL broth culture at 10,000 x g for 5 minutes and the cells were pelleted out and rinsed twice in 400 µL TE buffer after removal of supernatant. Cells were suspended in 560 µL of TE buffer (10 mM Tris, 1 mM EDTA; pH 8.0) followed by 30 μL of sodium dodecyl sulfate (10% wt/vol) and 3 µL of proteinase K (2% w/v) and incubated at 37°C for 60 minutes. About 100 μL of NaCl (5 M) and 80 μL CTAB/ NaCl (10% w/v CTAB, 0.7 M NaCl) was added to cell suspension and incubated at 65°C for 10 minutes in water bath. Chloroform/isoamyl alcohol (24:1) was added in same volume to the mixture and centrifuged 12,000 x g for 7 minutes to precipitate polysaccharides. Supernatant was collected and added same volume of phenol/chloroform/isoamyl alcohol (25:24:1), mixed thoroughly and
centrifuged at 12,000 x g, the supernatant was collected. Finally, isopropanol was added to the supernatant to precipitate the DNA. After centrifugation at 12,000 x g for 7 minutes the supernatant was removed and the DNA was resuspended in 80 µL TE buffer and RNase, and stored at -4oC for further study (Ausubel et al., 1995). The purified DNA was analyzed through agarose gel (1.5 g in TBE), and stained with ethidium bromide.
Sequencing of 16S rRNA was performed for identification of bacterial isolates. Amplification of full length gene was carried out using 27F’ (5’-AGAGTTTGATCCTGGCTCAG-3’) and 1494R’ (5’-CTACGGCTACCTTGTTACGA-3’) bacterial primers (Zeng et al., 2008). The 20 mL PCR reaction mixture consisted of 1 mL DNA sample, 2 mL PCR buffer, 2 mL deoxynucleotide triphosphate (dNTP) mix, both forward and reverse primers 2 mL each, 0.5 mL Ex-Taq DNA polymerase (Takara Shuzo, Otsu, Japan) and 10.5 mL distilled water. Initially, the reaction mixture was incubated at 96°C for 4 min. Then 35 amplification cycles were performed at 94°C for 45sec, 55°C for 60sec, and 72°C for 60sec. The reaction was incubated further for 7 min at 72°C. A positive control of Escherichia coli genomic DNA and a negative control were included in the PCR. Purification of PCR products were carried out using Montage PCR Clean up kit (Millipore) in order to get rid of distinct PCR primers and dNTPs from PCR products. Sequencing of the purified PCR product was performed by using 2 primers, 518F’ (5’- CCAGCAGCCGCGGTAATACG-3’) and 800R’ (5’-TACCAGGGTATCTAATCC-3’). Sequencing was done through Big Dye terminator cycle sequencing kit v.3.1 (Applied Bio- Systems, USA) and sequencing products were resolved on an Applied Bio-Systems model 3100 automated DNA sequencing system (Applied Bio-Systems, USA) by Macrogen, Inc., Seoul, Korea.
Chimeras of the obtained sequences were examined via Check-Chimera program of Ribosomal Database Project (RDP) (http://rdp.cme.msu.edu/seqmatch/seqmatch_intro.jsp) and also compared with 16S rRNA gene sequences present in public database GenBank (NCBI) by using BLAST search program (http://www.ncbi.nlm.nih.gov/BLAST/). The 16S rRNA gene sequences of various bacteria that are closely related to the studied sequence, as shown by BLAST search, were obtained from GenBank database and were aligned with the new sequences using BioEdit 6.0. The phylogenetic tree was constructed by the Maximum Likelihood method with robustness
of 1000 bootstrap value in MEGA 6.0 (Tamura et al., 2004). All the sequences obtained were submitted to NCBI GenBank and the accession numbers have been assigned.
CaCO3 bio-mineralization
For bacterial precipitation of CaCO3 crystals B4 medium was used. Composition of B4 medium used was (g/L); yeast extract, 4; dextrose, 10; calcium acetate, 2.5 and agar 15 g, (pH 7), autoclaved and poured into plates followed by standard streaking of isolates on the plates in such a way that one line was drawn in middle of the plate and incubated at 25°C. Results of CaCO3 precipitation were checked after 4 days of incubation at 25oC.
Effect of temperature and pH on growth of bacteria and CaCO3 precipitation
Effect of temperature (15, 25, 35, 40°C) and pH (5, 7, 8, and 9) on bacterial growth and CaCO3 precipitation was determined in order to optimize the growth and precipitation conditions on B4 medium.
X-ray Diffraction (XRD) of the crystals X-ray powder Diffraction (XRD) is a rapid analytical technique used for phase identification and characterization of unknown crystalline materials (e.g. minerals, inorganic compounds) and identification of fine-grained minerals such as clays and mixed layer clays that are difficult to determine optically (http://serc.carleton.edu/research_education/ geochemsheets/ techniques/ XRD.html). Calcium carbonate crystals were properly washed with distilled water and dried in oven. XRD patterns were obtained from the samples using (X’Pert-APD Philips, The Netherlands) with an X-ray generator (3 kW) and anode (LFF Cu). The Cu Kα radiation was administered at a wavelength of 1.54 Å. The X-ray generator tension and current were 40 kV and 30 mA, respectively. The step-scan data were continuously collected over the range of 5 to 80°2θ. The time constant was set at 2s to check the composition of precipitated calcium carbonates. Scanning electron microscopy Scanning electron microscopy (SEM) (Philips XL30CP) was performed to observe the morphology of crystals precipitated by bacterial strains.
Fourier Transform Infrared spectroscopy
Fourier transform infrared spectroscopy (FTIR) was performed to study the secondary structure of
CaCO3 crystal. CaCO3 crystals synthesized by bacteria were picked from the plate were placed in a holder and exposed to infrared beam for analysis through Fourier Transform Infrared Spectrometer (Jasco FT/ IR– 620). The samples were scanned from 4000-400 cm-1 at resolution of 6.0 cm-1.
Results
Metals analysis Mean concentration (mg/Kg) of elements in cave samples were in the following order; calcium (Ca) > Sodium (Na) > Magnesium (Mg) > Potassium (K) > Iron (Fe) > Manganese (Mn) > Zinc (Zn) > Chromium (Cr) > Lead (Pb) > Nickel (Ni) > copper (Cu) (Table 1). Mean concentration of these elements in cave samples were higher as compared to simple garden soil as a reference.
Isolation of Bacteria Number of viable cell count was calculated as 4.6×104 CFU/g. In this study, total 25 bacterial strains were isolated. The CaCO3 mineralization potential of all the isolates was investigated and only 3 isolates named GSN-11, TFSN-14 and, TFSN-15 was able to mineralize CaCO3.
Identification of bacteria
The 16S rRNA gene sequences of CaCO3 precipitating bacteria have been submitted to NCBI GenBank. The isolates GSN-11, TFSN-14 and TFSN-15 were identified as Bacillus toyonensis GSN-11, Paracoccus carotinifaciens TFSN-14, and Brevundimonas naejangsanensis TFSN-15 (Fig. 3.4.1).
Figure.3.4.1. Phylogenetic analysis by Maximum Likelihood method The evolutionary history was inferred by using the Maximum Likelihood method based on the Tamura-Nei model [1]. The tree with the highest log likelihood (-1525.1520) is shown. The percentage of trees in which the associated taxa clustered together is shown next to the branches. Initial tree(s) for the heuristic search were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of pairwise distances estimated using the Maximum Composite Likelihood (MCL) approach, and then selecting the topology with superior log likelihood value. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 22 nucleotide sequences. Codon positions included were 1st+2nd+3rd+Noncoding. All positions containing gaps and missing data were eliminated. There were a total of 751 positions in the final dataset. Evolutionary analyses were conducted in MEGA6.
Calcium carbonate precipitation Clear zone around streak was observed after 10 days of incubation (Fig. 3.4.2.A). These plates were further incubated and after 20 days increased precipitation was observed on streak surface (Fig.3.4.2. B).
A B
Fig. 3.4.2. Calcium precipitates induced by bacteria in crystals form A) Paracoccus Limosus, B) Brevundimonas naejangsanensis.
Parameters optimization for Calcium carbonate precipitation
Temperature effect Temperature was optimized for bacterial precipitation of Calcium carbonate in B4 medium. The maximum precipitation was observed at 25°C and beyond this, no precipitation was observed (Fig. 3.4.3).
Fig. 3.4.3. Compound microscopy of precipitates produced at 25°C. pH effect
Different pH values were checked for calcium carbonate precipitation in B4 medium. It was revealed that maximum crystal formation was observed at pH 5 (Fig. 3.4.4).
Fig. 3.4.4. Compound microscopy of precipitates produced at pH 5.
Incubation time effect Different incubation time was used for calcium carbonate precipitation. At optimum temperature and pH in B4 medium, optimum incubation time for maximum precipitation was observed 20 days (Figure. 3.4.5).
Fig. 3.4.5. Compound microscopy of precipitates produced after 20 days of incubation.
Scanning electron microscopy Scanning electron microscopy was performed for calcium carbonate crystals precipitated by bacterial isolates on B4 medium. Clear precipitations of the calcium carbonates were observed in scanning electron microscopy at different magnification (Fig. 3.4.6). The clear crystals formation
revealed that these cave bacteria are capable to precipitate the calcium carbonates that are present in abandoned amount in the natural environment of cave.
A B
C D
E
Fig. 3.4.6. Electron microscopy at different wavelength (A) at 500 µm, (B) at 200 µm, (C) at 100 µm, (D) 50 µm, (E) 1 micro meter.
Fourier transform infrared spectroscopy (FTIR) analysis Calcium carbonate purchased from sigma was used as a reference with bacterial precipitated
-1 - calcium carbonate sample. The control exhibited characteristic peaks of CaCO3 at 871cm , 712cm 1, 1794cm-1, 1405cm-1, all the samples analyzed, showed peaks in the same regions which indicated calcium carbonate presence in the samples precipitated by cave bacteria (Figure. 3.4.7).
Fig. 3.4.7. FTIR analysis of CaCO3 with control.
X-ray diffraction (XRD) analysis XRD was performed for the quantitative determination of calcium carbonate polymorphs. Fig 3.4.8 shows the XRD patterns of three polymorphic calcium carbonate crystals as calcite, aragonite and vaterite. All synthesized calcium carbonate samples were crushed into fine powder form for characterization (Fig. 3.4.8).
800
700 C
600 A
500
400
300
Intensity (arb.units) Intensity 200 v
100
0
20 30 40 50 60 70 80 2 (degrees)
key: C= calcite, A= aragonite and V= vaterite. Fig. 3.4.8. XRD analysis of the polymorphs of Calcium carbonate.
Characterization of CaCO3 mineralizing bacteria
All the CaCO3 mineralizing bacteria were subjected into cultural and biochemical characterization. Among four, 2 isolates, GSN-11 and TFSN-14 were gram negative, while TFSN-15 and GSN-22 were gram positive. Conventional biochemical tests were performed for all four isolates and reported in Table.2. All the isolates were optimally grow at 25°C and fall in mesophilic range, while at lower temperature the growth was negligible. The pH ranges of all four isolates were fall in acidic range and showed optimum growth at pH 5-8 11 and 9 while no growth was observed beyond this pH. Optimum incubation time for calcium carbonate preceptation was 20 days for all isolates.
The isolates named GSN-11, TFSN-14, TFSN-15 and GSN-22 were studied for phylogenetic analysis through 16S rRNA sequencing. Sequences obtained from these isolates exhibited variable similarities with the reference sequences from NCBI GenBank. Isolate TFSN-14, has 99% identity with Paracoccus Limosus, isolate TFSN-15 has 99% identity with Brevundimonas diminuta and isolate GSN-11 has 99% identity with Bacillus toyonensis.
Nucleotide sequence accession numbers The nucleotide sequence described in this study can be found from NCBI nucleotide sequence database under accession numbers
Discussion Low nutrient availability, low temperature pH and high humidity can be consider as paradigm of extreme environment in caves for all form of life. Microbes inhabiting caves play important role in cave formation via different process like bio-mineralization, speleogenesis and other geochemical process. The current investigation was aimed to isolate and characterised the possible role of cave microbes in bio-mineralization of calcium carbonate from Kashmir cave Buner Pakistan. A total of 3 strains were isolated in this study and characterized molecularly (16S rRNA). All the isolates clustered in to alpha Proteobacteria and Firmicutes, further differentiated in to Bacillus sp. GSN 11, Parcoccus sp. TFSN14 and Brevundimonas sp. TFSN15. All these isolates were good candidate for calcium carbonate precipitation. Generally, the precipitation processes are not related to a particular bacterial group, and have been reported from numerous ecosystem. Previous investigation shows that halophilic bacteria Exiguobacterium mexicanum was isolated from sea water and tested for biomineralization potential under different salt stress conditions (Bansal et al., 2016). Bacillus strains were most common calcifying isolates collected from Stiffes Cave (Ercole et al., 2001). Gamma- Proteobacteria (45%) group is the most dominant group of bacteria investigated in caves followed by Bacteroidetes, Firmicutes and Actinobacteria (Porter, 1971). Constitutive production of carbonic anhydrase, urease and secretion of EPS are common features of many species of soil bacteria (Moya et al., 2008; Burbank et al., 2011; Erecole et al., 2012).
The implementation of microbially produced calcium carbonate (CaCO3) in different industrial and environmental applications has become an alternative for conventional approaches to induce
CaCO3 precipitation. However, there are many factors affecting the bio-mineralization of CaCO3, which may restrict its application (Seifan et al., 2016). The effect of pH, temperature and incubation time period was investigated on growth and calcium carbonate precipitation by all the three isolates. B4 was the most common medium used in general organo- mineralization studies and has been used to characterize mineral precipitation potential. More than 50% of environmental bacteria are being tested to date are able to precipitate CaCO3 on B4, these include members of the Bacillus, Arthrobacter, Kingella and Xanthomonas (Sprocati et al., 2008). In the present study, the metabolic activity of bacterial strains and temperature are key factors in carbonate deposition. The optimum temperature of 25°C have a positive effect on bacterial precipitation of calcite, increasing the ability of the strains to form crystals. Laiz et al. (2003) reported bacteria from cave soil samples were able to grow comparatively well in broad range of temperature (13-45°C). Based on the results, pH dropped from 7 to 5 the enhanced precipitation of calcium carbonate was observed which leads to enrichment of CO2, and NH4, consequently, acidifying the surrounding
(Gat et al., 2014). pH variation during the precipitation of CaCO3 which is due to factors including
(i) NH3(g) dissolution, (ii) CO2(g) dissolution, and (iii) acid generation during the bacterial production of CaCO3 (Jiménez-López et al., 2001; Lopez et al., 2003). The process of precipitation is a complex mechanism. This mechanism is a function of the cell concentration, ionic strength and the pH of the medium. The media for the growth of the microorganisms are supplemented with a calcium source such as calcium chloride which is precipitated as calcium carbonate. The maximum precipitation was achieved by all the three isolates after incubation of 21-25 days. Precipitation of calcium carbonate was observed on the basis of zone and crystals formation. After proper incubation period the bacteria will attract the calcium toward its self-producing clear zones. Later on the calcium is precipitated by mean of different exopolymeric substances (EPS) at the top of media. Good results were obtained from Bacillus sp. GSN11. The crystal formation by calcification was analyzed for further studies.
The carbonate crystals formed by the Bacillus sp. GSN11 were characterized by FTIR, XRD and SEM. The aim was to investigate the crystals formation for calcium carbonate precipitation. SEM analysis revealed the crystal size varied from 50 to 500 µm and crystalline structure that were seen
embedded. The SEM analysis revealed distinct calcite embedded in bacterial cells. The association of calcite crystals with bacterial cells indicates that bacterial cells served as nucleation sites during the bio-mineralization process (Chahal et al., 2011; Achal et al., 2009). X-ray diffraction results showed that calcite is the major phase formed followed by aragonite and vaterite. XRD analysis showed that carbonate formation form which the microbes were isolated are formed from pure calcite (Ercole et al., 2001). Several studies have been conducted to understand the enigma behind the formation of a variety of carbonate polymorphs, which are found to be dependent on different factors such as growth media, substrate type, pH temperature, bacterial species, organic matter and
2+ 2− saturation index to [Ca ]/[CO3 ] ratio (Rodriguez-Navarro et al., 2012). Some authors also reported that, in case of saline environments, calcite formation is inhibited and aragonite formation increases with increasing concentration of Mg (Cailleau et al., 1977; Kitano et al., 1979; Sayoko and Kitano, 1985). FTIR spectra of the sample were prepared from plate to find the nature of the precipitated calcium carbonate crystals. The spectrum obtained from FTIR (400-400 cm-1) showed substantial similarity to that of standard calcium carbonate crystals (control). The presence of peaks at 1402.57, 871 and 712 cm-1 confirm the presence of calcium carbonate crystals formation. Three major peaks observed due to vibration of carbon oxygen double bond in the carbonate ion (Vahabi et al., 2015).
Conclusions In this study we investigated the bio-mineralization of calcium carbonate by cave microbes. The current study proved the feasibility of cave bacterial isolates in bio- mineralization and different geochemical processes in cave formation and bio- cementation. The outcome of the current study supports the potential of this technology for application in several fields such as remediation of concrete structures and stabilization of beach sands.
Assessment of Kashmir Cave Bacterial Isolates for the Oxidation of Manganese (Mn) and optimization of environmental parameters
Abstract
Mn (II) oxides are present abundantly in every environment, and very active in biogeochemical cycle of nutrients, carbon, contaminants, and other elements. It is believing that bacteria play a key role in Mn oxides precipitation in environment. Manganese oxidizing bacteria (MOB) are reported mostly from marine or other aquatic environment, and few from terrestrial. The current knowledge is on the precipitation of Mn oxides by five Kashmir cave bacterial isolates Bacillus pumilus C3, B. safensis C6, B. pumilus C7, B.cereus C8 and B. acidiceler C11. These Mn(II) oxidizing bacterial strains were isolated and purified on carbon rich K-medium. The Mn(II) oxidation by these isolated bacterial strains was enzymatically controlled reaction. The activity of Mn(II) oxidation was optimum at pH 5-7 and a temperature of 25-30oC and was lost at high temperature. Calcium ion (Ca+2) concentration affected the Mn(II) oxidation dramatically, while the Zn and Cu ions had no such high effect on the growth and Mn(II) oxidation. This demonstrates that cave bacteria are involved in the production of biogenic manganese oxides in cave environment.
Key words: Mn Oxidation, Mn (II) oxides, , Kashmir cave, biomineralization
Introduction
Caves are colonized by variety of microorganisms such as bacteria, fungi, algae and protozoans. Bacteria could be present in cave water, surface and subsurface soil. They may attach to walls or reside in guano. Hoeg, (1946), found microbes which were attached to the walls of Norwegian caves. An earlier review on cave microorganisms was provided by Caumartin (1963). Most of the microbes identified till today are either opportunistic or grow in suitable conditions (Dickson and Kirk, 1976; James, 1994; Jones and Motyka, 1987). Many of the microbes inside the cave are transient microbes i.e. they are brought inside the cave through water, sediment, air and animals. Besides, they may form a parasitic relationship with troglobiotic animals and epibionts (Golemansky and Bonnet, 1994). Cave explorers or humans going inside the cave may act as a means of pathogen transfer from cave to outside environment (Li et al., 2010). Researchers and tourists who deal with bats and cave guano are vulnerable to cave pathogens (Juardo et al., 2010).
Microorganisms are geologically substantial for a number of processes, such as mineral decomposition, mineral formation, biogeochemical cycles and sedimentation. They decompose or form minerals by the help of their enzymes or metabolic products. Their anabolic or catabolic products, along with their respiratory mode (aerobic/anaerobic), either constitute or decay the surroundings. Their mode of nutrition defines and distributes microbes into either chemolithotrophs, the bacteria which utilizes inorganic products such as H2, Fe (II), Mn (II) and
H2S, or photolithotrophs, the sulfur deposits by assimilation of CO2 and utilization of H2S. In case 2 3 of aerobic mineralization of organic compounds, CO2, H2O, NO3¯, SO4 ¯, PO4 ¯, whereas 2 anaerobic mineralization results in the production of CH4, CO2, NH3, H2S, PO4 ¯ (Ehrlich, 2002). Up till now, many studies have shown the connection of microorganisms with speleothems such as carbonate, silicates, sulfur molecules, oxides of iron and manganese and compounds containing potassium nitrate (Northup et al., 1997). Microorganisms carry out the process of precipitation either in a passive manner through sites of nucleation (Went, 1969), or actively by yielding enzymes that intern precipitate the minerals (Danielli and Edington, 1983). Metabolic by-products resulted by acidification reaction of microbes can dissolute the cave topography. Microbial interaction with cave topology gives the idea of general processes of precipitation and dissolution that results in the genesis of different types of speleothems (Northup et al., 2000). Among these processes, Mn oxidation occurs that play an essential role for the survival of microorganisms.
Manganese in Caves
The term ‘Mn oxides’ is collectively used for the oxides, hydroxides and oxyhydroxides, which are highly reactive phases of mineral which play an important role in geochemical cycles of elements. The Earth’s crust is composed of about 0.1% manganese (Nealson, 1983). In Earth’s crust manganese secured fifth position in transition metals (Tebo et al., 2004). Manganese is present in 7 different oxidation states extending from 0 to +7 while naturally it is present in +II, +III and +IV states (Tebo et al. 1997, 2004). Mn have higher redox potential than iron due to which the reduction of Mn is easier than Fe, and tough to oxidize than Fe (Kirchner and Grabowski, 1972).
Oxyhydroxides are found in abundant after sulfate and carbonate minerals in caves. Many reports are available which showed the presence of iron and manganese in abundant form in caves (Hill and Fort, 1997; White et al., 2009; Gazquez et al., 2011) to irregular surfaces on the walls usually on top of visibly altered carbonates (Northup et al., 2003; Spilde et al., 2005; Gazquez et al., 2012a). With carbonate and silicate speleothems, sulfur compounds, oxides of iron and manganese and saltpeter, the microbial associations have been reported (Northup et al. 1997). Manganese compounds are present in caves as in clastic deposit form layer on wall or speleothems (Gascoin 1982; Hill 1982) or as crust (Jones 1992: Moore 1981). In cave, manganese is present in the form of birnessite very common (Hill and Forti, 1997), and some low quantity of crystals oxides and hydroxides like pyrolusite, chalcophanite, cryptomelane, hausmannite, romanechite, rancieite, todorokite and rhodochrosite are also reported (Onac et al.1997a; Onac et al. 1997b). From karst solutional cavity, the manganese is also isolated (Jones, 1992). Peck (1986) and Northup et al. (2003) provided evidences that manganese oxides reported from caves are almost biogenic in nature. Mn (IV) oxide are present in aquatic as well as in terrestrial environment (Post, 1999).
Manganese oxidation by cave microorganisms
The cavernicoles play important role in mineral precipitation either actively by producing enzymes or other metabolites which change the microenvironment (Danielli and Edington 1983), or passively by acting as nucleation site (Went, 1969). Cave microbes also play an important role in cave dissolution by producing the acid as a byproduct (Ehrlich, 1996).
Manganese is a vital trace element that is required by all living organisms. It acts as a cofactor in a variety of enzymatic processes that are necessary for metabolic activities and antioxidant, as well as superoxide dismutase and photosystem II (Tebo et al., 2004). Generally, Mn is found in three oxidative states. In surface environment Mn is found as soluble reduced form Mn (II), and also in oxidized form as Mn (IV). In between these two species of manganese, an intermediate oxidizes phase on Mn also occurs as Mn (III) that can form complex bonds with organic compounds (Madison et al., 2013). Mn cycle occurs between these three states and it has a direct role in environmental health as well as on humans. The oxidized species of manganese i.e. Mn (III/IV) has a major impact on the fate of several nutrients along with pollutants (Spiro et al., 2010; Geszvain et al., 2012). Mn (III) also oxidizes carbon compounds and metals (Trouwborst et al., 2006; Madison et al., 2013). Even though Mn oxidation can occurs through abiotic reactions but the efficiency and production of Mn oxides is more in biological processes because microorganisms have the capability to oxidize Mn (II) at quicker kinetic rate than abiotic processes (Nealson et al., 1988; Tebo, 1991). Those bacterial species that are adequate to oxidize Mn (II) (Tebo et al., 2004; Tebo et al., 2005) through several pathways using enzymes, such as multicopper oxidases (Corstjens et al., 1997; Dick et al., 2008; Butterfield et al., 2013; Geszvain et al., 2013; Su et al., 2013), heme-peroxidase (Anderson et al., 2009), and two-component regulative protein (Geszvain and Tebo, 2010). Although biogenic Mn oxidation is known, the physiological purpose of bacterial manganese oxidation is not yet acknowledged (Learman et al., 2014). At neutral pH, abiotic Mn oxidation occurs and results in the production of phyllomanganate forms of bernessite mineral group that has hexagonal or triclinic structure having poor crystalline Mn oxide to crystalline bernessite.
Microorganisms of caves are capable of producing extracellular polymeric substances (EPSs) and some other metabolites with acidic functional groups, which stress the pH lowering due to metal oxidation. Due to deprotonation of organic functional group, the cell walls of bacteria have negative charges at low pH, which may act as nucleating sites for the cation like iron and manganese (Fein, 2009).
Under optimum pH conditions, manganese is thermodynamically oxidized through reactive oxygen species (ROS) super oxide (O−2). In the previous studies, scavenging of superoxide through Mn oxides has also been reported (Barnese et al., 2008) because Mn (II) is very significant
antioxidant in natural processes. Its nano molar levels have been observed in sea water that could scavenge superoxide which leads to a fast Mn cycle (Hansard et al., 2011). Conversely, another study showed that superoxide, in the presence of organic carbon can induce Mn (II) oxidation at faster rate (Nico et al., 2002). During this reaction Mn (II) is oxidized and superoxide is reduced to hydrogen peroxide (H2O2).
−2 + Mn (II) + O + 2H → Mn (III) + H2O2
Enzymatically superoxide production and its effect on manganese oxidation has been observed in marine bacteria as well, including Roseobacter sp. AzwK-3b, Roseobacter clade (Learman et al., 2011). This manganese oxidation through biological superoxide leads to the formation of manganese intermediate Mn (III). In fungi, ascomycetes have been studied to oxidize manganese Mn (II) with the aid of extracellular superoxide (Learman, 2013; Hansel et al., 2012; Tang et al., 2013).
In the ecosystem, biogenic manganese oxidation has also been affected by a variety of ions. However, there is brief knowledge about the role of these ions on kinetics of manganese oxidation by bacteria. Previous studies have shown the effect of Ca2+ concentration on manganese oxidation + + 2 - 2+ in Bacillus sp., whereas K , Na , Sr and NO3 ions had no effect. At 10 mM Ca concentration, Mn (II) oxidation was increased up to 4-5 times. Studies have suggested that calcium ions have a direct role on Mn oxidizing enzymes by possibly increasing the bridge amongst polypeptide components (Toyoda et al., 2013).
As Mn oxidants are extremely reactive species and can carry out oxidation-reduction reactions at broad range of pH. These redox reactions have an essential role in bioavailability and mobility of toxic heavy metals including Cu, Ni, Co, Pb, Ba, Zn, Ag, Hg and Tl (Sherman and Peacock, 2010; Peacock and Moon, 2012). Up till now, little studies have been done on Mn oxidation at low pH however it is necessary to evaluate biogenic efficiency of Mn precipitation at low pH because most of the mining processes are characterized by acidic pH. Some Streptomyces sp. have been reported to oxidize Mn at pH ranging from 4.5-5, whereas Chlorococcum humicolum algae and Cephalosporium sp. of fungi is capable of oxidizing Mn (II) at pH 4.5. In another study, Mn oxides could immobilize heavy metals in biogeochemical barrier layers formed during Mn oxidation at low pH (4.7 to 5.1). The bacterial samples were isolated from a former mining area of uranium. Six bacterial species were found to produce hexagonal bernessite Mn oxide in its layers, which could immobilize Ba, Cu, Ni, Zn, Co, Cd and Ce. These bacteria were Bacillus safensis, Bacillus altitudinis, Bravibacilllus reuszeri (Gram positive spore formers), Arthobacter and Frondihabitans (Gram positive Actinobacteria) and Sphingomonas (Proteobacteria).
Fig.4.1. Mn cycle of oxidation states found in nature
The biogenic manganese oxides are highly influenced by factors which determine its oxidation state. The Mn(II) oxidation is naturally thermodynamically favorable reaction but a very slow process (Diem and Stumm, 1984; Nealson et al., 1988). The abiotic oxidation of Mn(II) is a very slow process and catalyzed by environmental conditions like high pH and high oxygen pressure, or the oxidation of Mn(II) is also catalyzed by adsorption of Mn(II) ions on mineral surfaces such as iron oxides and silicates (Morgan and Stumm, 1964; Sung and Morgan, 1981; Hem and Lind, 1983; Murray et al., 1985; Davies and Morgan, 1989). In the presence of reducing agents and in anaerobic condition, low pH, or in Mn(II) complexing agents, Mn(IV) is reduced to Mn(III)(II).
Fig. 4.2. Four possible
mechanisms of Mn+2 oxidation by bacteria.
Enzymatic Mechanism of Bacterial Mn(II) Oxidation
From the advancement in identification of enzymes involved in Mn+2 oxidation produced by bacteria and the mechanism by which the oxidation occurs, many questions remain debatable. The involvement of multicopper oxidase enzymes in the oxidation of Mn+2 must be differentiate from actual direct catalysis of Mn+2 oxidation by multicopper oxidase enzymes (MCO). While in the study of Leptothrix discophora has a potential link made between Mn oxidase and gene responsible for the coding of multicopper oxidase enzymes (MCO) (Corstjens et al., 1997). Bacterial Mn oxidase enzyme has not been purified in quantity for detailed biochemical investigation, and no MCO encoding gene is successfully expressed to produce functional enzymes in foreign host. Thus the direct involvement of MCO in Mn+2 oxidation is a hypothesis. In spite of it seems debatable that MCO in bacteria could directly involve in Mn+2 oxidation because (a) genetic and biochemical studies showed the involvement of MCO in Mn+2 oxidation in many unrelated bacteria, (b) some MCO isolated from eukaryotic are known to oxidize Mn(II) directly (Hofer and Schlosser, 1999; Schlosser and Hofer, 2002), and (c) the Fe(II) oxidizing MCO occur in both eukaryotes (Solomon et al., 1996) and bacteria (Kim et al., 2001; Huston et al., 2002).
Fig.4.3. Enzymatic pathway of Mn(II) oxidation
Importance of Mn oxide
Manganese oxides are important because Mn (III/IV) act as cofactor and are responsible for a number of redox reactions. They are also utilized as trace nutrients. In aquatic system, photosynthetic organisms require Mn oxides in the photosystem II and water molecule is split to obtain energy and thus Mn stores oxygen in surrounding environment. Even though manganese was considered biochemically important, its cycle is not completely reported therefore it lead to the further studies of the ionic species of manganese. In a recent study, Mn speciation was identified by filtration method, in which a sample containing manganese was passed through 0.2 µm filter. Dissolved portion was rich in Mn (II) whereas the unfiltered fraction contained soluble form of Mn (III/IV) i.e. oxidation speciation of manganese. Previously Mn (III) was considered thermodynamically unstable and its efficiency was neglected. Nevertheless, in a study conducted by Luther (2010), Mn (III) was found significant for such redox reactions in which one electron is transferred from donor to acceptor molecule. During the process of oxidation, when the transfer of an electron occurs between Mn (II) and Mn (IV), Mn (III) acts as an intermediate ionic state that overcomes the ionic orbital distance. Mn (III) state occurs for a short period of time but when provided with a ligand, Mn (III) state is stable because in complexes Mn (III) ions have higher electrostatic charge and shorter radius than Mn (II) or Mn (IV). Mn (III) also has higher bonding affinity with siderophore than Fe (III) and thus competes with Fe (III) and decreases its uptake in microorganisms and plants (Oldham et al., 2015).
In addition to heavy metal immobilization, Mn oxides are also being used for pollution degradation and dye removal in waste water treatment plants. Besides, Mn oxides combined with gold nanoparticles are used to remove volatile organic compounds (VOC) from atmosphere. In case of VOC removal, Mn oxides efficiently remove hexane, toluene, nitrogen oxides and sulfur oxides, are major compounds present in polluted air (Sinha et al., 2007). Organic dye present in water have led to water contamination and is a huge source of environmental pollution. Due to the physiochemical properties of manganese oxides (Mn III/IV), several studies have been carried out for efficient dye removal through absorption and catalytic breakdown processes. Studies have reported the degradation of RhB dye through redox reaction using Mn (III/IV) oxides having different crystalline structures. Even at low pH (2-6), alpha, beta and gamma structures of manganese oxides can decolorize RhB by cleaving ethyl groups, degradation of carboxylic group - 2- (–COOH) and finally complete mineralization to produce CO2, H2O, NO3 , SO4 and NH4+ (Cui
et al., 2015). In a recent study, methylene blue and methyl orange dye degradation has been achieved using MnO2 coated diatomite. Fabricated Mn oxides have the potency to treat waste water, and remediation of ecosystem (Trung et al., 2016). In another study MnO2 based micromotor were used to have double effect, i.e. dye degradation and bubble separation through absorption. This dual technique resulted in decolorization of dye in waste water more than 90%. This revealing the efficiency of Mn oxide micromotors, for the treatment of waste and portable water reservoirs for removal of dyes (Wani et al., 2015).
Material and Methods
The study is about manganese oxidation by bacterial strains isolated from soil and speleothem collected from Kashmir (smast) cave (Fig.4.4). The cave is located in the Babozai mountain located between District Mardan and Buner having coordinates (34o25’42.12”N 72o13’10.82”E). The length of Kashmir cave is 188 meters, and width is ~ 29 meters. The height of entrance zone is about 30 meters and higher in twilight and dark zone. Kashmir cave is a series of limestone caves. Like other caves, it has low temperature i.e. 10oC and pH of 5-6. The surface was wet due to drifting of water from the cave ceiling. Speleothems, secondary metabolites were formed from entrance to dark zone.
Fig.4.4. Kashmir Smast (Cave) entrance zone
Sampling
In August 2015, soil and speleothem samples were collected from Kashmir cave. The soil and speleothems samples were collected from the dark zone of cave. Soil samples were collected from the surface layer (at depth of 0 cm to 50 cm). While the speleothem samples were collected from
the ceiling of cave. All the soil and speleothem samples were stored in sterile zipper bags, placed in an ice bath, transported to laboratory and kept at 4oC until used for experiments.
Fig.4.5. Speleothems isolated from Kashmir smast (Cave)
Isolation of Mn(II)-oxidizing bacteria The glassware used in whole research was acid washed for 24 hours in 2 M HCl and rinsed with milli-Q water. For isolation of manganese oxidizing bacterial strains from the cave soil samples, 1 g soil sample was serially diluted in normal saline and then was spread on K medium plates, aseptically. Then plates were incubated aerobically at 35°C. Brown coloured colonies were isolated and purified on fresh K medium plates. For the confirmation of the ability of manganese oxidation by these bacterial strains, drops of 0.04% (w/v) leucoberbelin blue solution in 45mM acetic acid was added on the colonies. Formation of blue color due to the reaction of LBB with Mn(III/IV) oxides, confirm the ability of these microbes to oxidize manganese.
Similarly, the bacterial strains were isolated from speleothem by sprinkling the speleothem powder on nutrient agar plate and purified the cultured bacterial strains.
Screening of bacteria for the Mn oxidation
All the isolated bacterial strains were grown on defined medium i.e. K agar medium plates, containing FeSO4.7H2O (0.001 g/l), MnCl2 (0.2 g/l), peptone (2.0 g/l), yeast extract (0.5 g/l), and
10 mM HEPES buffer (N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid; pH 7.5). For the Mn oxidizers, Lecoberbelin Blue (LBB) was used as indicator. LBB was prepared by suspending
0.04% LBB in 45 mM acetic acid. K medium was prepared by adding 25 µL of 100 µM MnCl2 and 5 ml HEPES buffer in autoclaved 500 ml of deionized water having 0.5 g of peptone, 0.3 g of yeast and 2% of agar. The plates were incubated at 15oC for 48 hrs. After incubation, few drops of LBB were poured on bacterial colonies for the confirmation of Mn oxidizers.
Screening of Mn(II) oxidzation bacterial strains
For the production of biogenic manganese oxide, 2 ml cultures at log phase were inoculated in a volume of 75 ml of organic rich K broth medium in a 300 ml flask, and incubated in a shaker incubator of 150 rpm and at 25oC for 60 hours. Control was also incubated without inoculating bacterial strains. About 0.3 ml of sample was taken after every 4 hours for the detection of manganese oxidation during the growth process. Lecoberbelin blue (LBB) solution of 0.9 ml was added to 0.3 ml of growing cells at a ratio of 3:1 and incubated at temperature of 25oC for 15 minutes in dark condition. After incubation, the sample was centrifuged for 5 minutes at 10,000 rpm. The supernatants were taken in a cuvette and measured its absorbance by using UV-Vis at 620 nm. For the establishment of calibration curve, potassium permanganate was used. While in a control, K broth medium and LBB was used. Growth curve was checked by growing the culture in K broth medium and measuring the growth at 600 nm.
Molecular identification of Mn oxidizers
DNA extraction
DNA extraction was performed by centrifuging 1 ml of broth culture at 10,000 rpm for 5 minutes, cells were pelleted out. The pelleted cells were rinsed twice in 400 µL TE buffer after removing the supernatant. After rinsing, the cells were centrifuged at 10,000 rpm for 5 minutes, the pellets were resuspended in 200 µL TE buffer.
Then 100 µL Tris-saturated phenol (pH 8.0) was added to these tubes, followed by a vortex-mixing step of 60 sec, to lyse the cells. To remove the aqueous phase from organic phase, the samples were centrifuged at 13,000 rpm at 4°C for 5 minutes. Then 1.5 ml tube was taken and 160 μl of upper aqueous phase was poured to it. About 40 µL of TE buffer was added to make 200 µL and
mixed with 100 µL of chloroform: isoamyl alcohol (24:1) and centrifuged for 5 min at 13,000 rpm at 4°C. Chloroform: isoamyl alcohol (24:1) extraction was used for the purification of lysate, when there was no longer a white interface, and the same method was repeated twice or thrice. Purified DNA was present in the aqueous phase and was stored at -20°C for further use.
Preparation of the Agarose Gel
About 1.5 g of agarose in 1x TBE, was kept in microwave oven for making the gel. Then 2-5 μl ethidium bromide was added to this agarose solution and mixed by shaking. It was then transferred to gel tray and wells were made in the gel for loading samples by inserting a comb in the liquid gel. It was allowed to solidify for about 15 minutes. About 1 μl of loading dye and 5 μl of supernatant were mixed and loaded on agarose gel (1.5%). The gel was run for a period of 45 min at 110 V and 400 mA and visualized under UV.
Phylogenetic analysis
Phylogenetic analysis was carried out to study sequences using ClustalW program implemented in MEGA4.0 (Thompson et al., 1994). The similar sequences were downloaded from NCBI. All sequences were aligned and the phylogenetic tree was constructed using Neighbor Joining method in MEGA 4.0 bootstrap analysis (1000 replicate) was used for the significance of the generated tree.
Examination of soluble extracellular oxidation
For the examination of extracellular Mn(II) oxidation, all the Mn(II) oxidizing strains were grown up to mid log phase and then filtered in sterile condition. After filtration, 100µM of Mn(II) was added and incubated at 30oC and centrifuged at 150 rpm for 48 hrs. Sample was taken after every 4 hr to check for Mn(II) oxidation by adding 0.9 ml of LBB to 0.3 ml, and measured its absorbance by using UV-Vis at 620 nm.
Examination of superoxide oxidation (SOD)
For the confirmation of oxidation by superoxide dismutase (SOD), two sets of K broth medium one containing 100 µM of MnCl2 and 1 µM SOD and the second contained 100 µM MnCl2 and 5 µM SOD was inoculated with mid log phase culture and incubated at 25oC and 150 rpm for 60 hr. The SOD was syringe filtered into the medium. The control was K broth without SOD. The Mn(II) oxidation was examined by measuring the absorbance after every 4 hr of incubation at 620 nm on UV-Vis spectrophotometer. The experiment was run in duplicate.
Effect of temperature on Mn(II) oxidation
To check the optimum temperature for Mn(II) oxidation by isolated strains, all five isolates were grown at 25oC and 30oC in duplicate in liquid K medium in a shaker incubator at 150 rpm. Mn(II) oxidation was checked by measuring the formation of manganese oxides in a quantitative LBB colorimetric assay after every 4 hr of growth by adding LBB at a ratio of 3:1 to the centrifuged samples. The absorbance of the Mn(II) oxide produced was measured at 620 nm on a UV-Vis spectrophotometer.
Effect of pH on Mn(II) oxidation
Effect of pH on the ability of bacterial strains to oxidize Mn was determined by growing them on modified K agar medium (adjusted to pH 4, 5, 6, 7, and 8) plates. The pH of the medium was adjusted by adding NaOH or HCl. Plates were incubated at 25oC and the growth was checked visually, while the oxidation of Mn(II) was confirmed by adding LBB spot test.
Effect of metal ions on Mn (II) oxidation by bacteria
Tolerance of the isolated strains to copper, zinc and calcium concentration was evaluated in K broth medium. All the five strains were cultured in duplicate. Zinc, copper, and calcium were added in the growth medium at final concentration of 100 µM and inoculated by the selected isolates and incubated for 36 hrs at 25oC. Growth was measured by determining optical density at
600 nm (OD600). Mn(II) oxidation by the isolates was evaluated by measuring production of Mn oxides through quantitative LBB colorimetric assay after 48 h of growth. About 0.1 ml of centrifuged sample was added to 0.3 ml of LBB and the mixture was incubated in dark for 15 min
at room temperature. After incubation, absorbance of the reaction was measured at 620 nm on a
UV-Vis spectrophotometer. Standard curve was plotted with KMnO4.
Results
Manganese (Mn) is an essential metal which is not easily oxidized like iron. Many bacterial strains have the ability to oxidize Mn(II) to Mn(III/IV) five times faster than chemical oxidation. Mn oxides are strongest oxidants next to oxygen in aquatic environment. Mn oxidizing bacteria produce enzymes which not only help to scavange Mn but also other associated elements, thus playing an important role in biogeochemical cycles. In this study we present the isolation of bacterial strains from Kashmir cave having the ability of Mn(II) oxidation to Mn(III/IV).
Isolation of Mn(II) oxidizing bacteria For the isolation of Mn(II) oxidizing bacteria from Kashmir cave soil and speleothem samples, K agar medium was used. About 30 different bacteria were isolated from soil and 4 from the pinkish layer of speleothem. Among 30 soil bacterial isolates, 5 were positive for oxidization of Mn(II) in initial screening, while 3 of 4 from the speleothem were also Mn(II) oxidizers (Table 4.1).
Table. 4.1. Mn(II) oxidizing Bacteria Isolated from Kashmir cave soil and speleothem. Bacterial isolates from Kashmir cave soil Bacterial isolates from Kashmir cave speleothem S.No. Isolate Strain Isolate Strain
1 C 3 Bacillus pumilus S 1 Bacillus cereus
2 C 6 Bacillus safensis S 2 Bacillus cereus
3 C 7 Bacillus pumilus S 4 Bacillus cereus
4 C 8 Bacillus cereus
5 C 11 Bacillus acidiceler
On basis of ICP-MS analysis of the soil, the Kashmir cave soil contains 0.246% wt of manganese. The pH of the soil was 5-6. All the isolates showed the oxidation of Mn(II) by a positive LBB spot test. Initial confirmation was performed by plate assay shown in (Fig 4.6). The color of the isolates on K agar medium plate was brown, and after adding 2-3 drops of LBB on the colonies the color was changed to bluish.
Fig. 4.6. Initial screening of Mn(II) oxidizing bacterial strains from cave soil.
Stereoscopic confirmation The ability of the Mn(II) oxidation was also confirmed by stereoscopy. The color of the colonies was changed to brown by adding LBB shown in (Fig 4.7).
Fig. 4.7. Stereoscopy of the isolates.
Molecular identification of isolated Mn(II) oxidizing bacteria Bands of extracted DNA from Mn(II) oxidizers isolates from Kashmir cave soil and speleothem were clearly observed on gel when subjected to UV illuminator, shown in (Fig 4.8)
Fig 4.8. DNA bands of Mn(II) oxidizing isolates.
Phylogenetic Analysis of Mn oxide producers
The sequences from 16S rRNA gene obtained from Macrogen Incorporation were used for BLAST in NCBI gene bank database, which revealed Mn(II) oxidizers belong to genus Bacillus. The phylogenetic tree was constructed through MWGA5 software (Fig 4.9).
Bacillus pumilus (KR780404) Bacillus pumilus (KR528376) Bacillus pumilus (KR780437) Bacillus pumilus (CP000813) Bacillus pumilus (KT273321) Bacillus pumilus (KT624615) 100 Bacillus pumilus (KT624616) Bacillus safensis (KP235236) Bacillus pumilus (KP235237) Bacillus pumilus (KP235238) 98 Bacillus safensis (KT719214) C3 C6 Bacillus pumilus (LN890660) C7 Bacillus acidiceler (KU254664) 100 Bacillus acidiceler (LN890177) C11 C8 Bacillus cereus (KT720292) 100 Bacillus cereus (KT720291) Bacillus cereus (KT719760)
0.005 Fig. 4.9. Phylogenetic analysis by Maximum Likelihood method
Evolutionary history was inferred by using the Maximum Likelihood method based on Tamura- Nei model (Tamura and Nei, 1993). Tree with the highest log likelihood (-1391.0539) is shown in Fig. (4.9). The percentage of trees in which the associated taxa clustered together is shown next to the branches. Initial tree for the heuristic search was obtained automatically as follows. When the number of common sites was < 100 or less than one fourth of the total number of sites, the maximum parsimony method was used; otherwise BIONJ method with MCL distance matrix was used. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 11 nucleotide sequences. Codon positions included were 1st + 2nd + 3rd + Noncoding. All positions containing gaps and missing data were eliminated. There were a total of 740 positions in the final data set. Evolutionary analyses were conducted in MEGA5 (Tamura et al., 2011).
Growth curves of Mn(II) oxidizing cave isolates Due to the presence of Mn in cave soil, the cavernicoles (cave microbes) have intrinsic capability to oxidize Mn(II) to Mn(III/IV). The optimum conditions were checked for the production of Mn oxides by these isolates. The isolates from speleothem showed low ability of Mn(II) oxidation therefore were not processed further. Soil isolates showed more oxidation of Mn(II) as compared to bacteria from speleothem. Soil isolates were grown in K broth medium and their growth was checked at O.D600 after every 4 hours of incubation. Growth was checked in the presence and absence of MnCl2, after adding 1 and 5 µM of Super Oxide Dismutase (SOD) and 100 µM of calcium acetate in K broth medium. The growth of all isolates is shown in (Fig. 4.10).
Fig. 4.10a. Growth curves of Bacillus pumilus C3 at 30oC and 25oC (No Mn 600, 30oC and
o o No Mn 600, 25 C), in the presence of MnCl2 (Mn600, 30 C), after adding 1 and 5 M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600).
Fig. 4.10b. Growth curves of Bacillus Safensis C6 at 30oC and 25 oC (No Mn600, 30oC and
o o No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30 C), after adding 1 and 5 M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600).
Fig. 4.10c. Growth curves of Bacillus pumilus C7 at 30oC and 25 oC (No Mn600, 30oC and
o o No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30 C), after adding 1 and 5 M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600).
Fig. 4.10d. Growth curves of Bacillus cereus C8 at 30oC and 25 oC (No Mn600, 30 oC and
o o No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30 C), after adding 1 and 5 M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600).
Fig. 4.10e. Growth curves of Bacillus acidiceler C11 at 30oC and 25 oC (No Mn600, 30 oC
o o and No Mn600, 25 C), in the presence of MnCl2 (Mn600, 30 C), after adding 1 and 5 M of Super Oxides Dismutase (SOD) (Mn+SOD1,600. Mn+SOD5,600), and in the presence of 100M Calcium acetate (Mn+Ca, 600).
Examination of soluble extracellular oxidation After 48 hrs of reaction, no visible biogenic Mn oxide production was observed. Addition of colorimetric dye, leucoberbalin blue (LBB), which oxidizes and changes to blue color in the presence of Mn(III) and Mn(IV), showed no reaction, confirming the lack of Mn oxidation.
Examination of Superoxide oxidation (SOD) Following 36 hrs of experiment, no visible changes in the precipitation of Mn oxides were observed in the presence and absence of SOD, or change in color of LBB after adding it to the reaction solution (Fig. 4.12). The results confirm that the production of Mn oxide is enzymatic not chemical.
Optimization of Mn oxide production
Ability of the cave isolates to precipitate Mn oxide was dependent on pH of the medium and growth phases. Mn oxidation was checked at different pH like 5, 6 and 7 and they all showed growth at range of 5 – 7 pH, similarly Mn oxidizers were checked at different temperature and optimally grew at temperature 25 and 30oC. B. cereus C8 and B. acidiceler C11 showed maximum oxidation under pH and in the presence of various Ca+2 concentrations. Whereas, less ability of Mn(II) oxidation was observed in case of B. pumilus C3, B. safensis C6 and B. pumilus C7 (Fig. 4.11). The optimum Mn(II) metabolism was observed at 25oC (Fig. 4.12).
Mn Oxidation at different pH and Ca+2 concentration
1.6 1.4 1.2 1 B. pumilus C3 0.8 B. Safensis C6 0.6 0.4 B. pumilus C7 0.2 B. cereusC8
Mn oxidation rate (µM/h) rate oxidation Mn 0 B. acidiceler C11 0/5.0 1/5.0 5/5.0 0/6.0 1/6.0 5/6.0 0/7.0 1/7.0 5/7.0 Ca+2 conc (µM) in pH
Fig. 4.11. Variation in Mn(II) oxidation at different pH and Ca+2 ion concentration.
(a) B. pumilus C3
(b) B. safensis C6
(c) B. pumilus C7
(d) B. cereus C8
(e) B. acidicelar C11 Fig. 4.12. Mn(II) oxidation capacity and Mn(III,IV) oxide concentration as a function of reaction time in C rich media K-medium. The ages of the Mn oxide were from 4 h to 36 h.
Effect of metals (Zn, Cu and Ca) on Mn oxide precipitation by Cave bacteria Growth of the isolates was affected by the tested metals, with direct effect on Mn oxide precipitation. Growth and Mn(II) oxidation by Kashmir cave isolates was very low in early stage of incubation in the presence of zinc and copper ions but after Growth and Mn oxidation by bacteria increased after 18 h of incubation. Ca+2 dramatically encourage the growth and Mn oxide formation (Fig. 4.13).
Effect of metals on Mn oxidation 1.2
1
0.8 B. pumilus C3 0.6 B. safensis C6
B. pumilus C7 O.D (620)nm O.D 0.4 B. cereus C8 B. acidicelar C11 0.2
0 Mn Mn+Ca Mn+Zn Mn+Cu Mn+Metals
Fig. 4.13. Effect of metals on Mn oxide production by cavernicoles after 24 h of incubation.
Discussion
The presence of Mn oxides and oxyhydroxides which are insoluble, are a good sign for the microbial activities (Mann et al., 1990). The reported biogenic Mn oxides produced by bacteria in laboratory are dominantly Mn(IV) as opposed to lower oxidation states (Tebo et al., 1997). The biogenic Mn oxidation is a significant process for the production of biosignature, because at neutral pH the abiotic Mn(II) oxidation is kinetically very slow mechanism. Microbes play a key role in the oxidation of Mn(II) in terrestrial environments because they speed up the rate of Mn(II) oxidation (Nealson et al., 1988).
In most terrestrial environments, manganese accompanies iron in mineral suites, because together they play an important role in lives of many microorganisms (Ghiorse and Ehrlich, 1992). From cave environments, manganese minerals are also reported occur with iron minerals. At pH>6, the abiotic oxidation of soluble Fe(II) occurs very rapidly because it is highly pH dependent reaction. On the other hand, the reduction of Fe(III) and Mn(IV) is not favored thermodynamically under oxic conditions but may readily occur under low pH (Nealson et al., 1983).
Mostly Mn(II) oxidizing bacteria have been reported from marine environments, only a few number of bacterial strains have been isolated from terrestrial environments which have the ability of Mn(II) oxidation. The gene responsible for the Mn(II) oxidation has not been identified from soil bacteria (Waasbergen, et al., 1996). Thus the soil-borne bacteria specially from cave environment needs to be investigated for the Mn(II) oxidation. For this investigation various culturable cave bacterial strains should be isolated from cave soil for the mechanistic studies of biogenic manganese oxidation in cave environment. In present study, soil samples were collected aseptically from Kashmir Cave in which Mn was found 0.24 wt%. The Kashmir cave soil sample (nutrients limited and pH 5-6) was first time studied for the diversity of culturable bacterial strains having the capability of Mn(II) oxidation. The culturable bacterial strains on K-medium showed the Mn(II) oxidizing activities. Most reports on the production of biogenic Mn oxides production, the pH has been maintained at neutral or alkaline. We isolated manganese oxidizing bacteria (MOB) from the nutrient limited and acidic soil of pH 5.5. Five bacterial strains were isolated, and characterized by 16S rRNA. The isolates were assigned into a single cluster or phyla firmicutes Bacillus pumilus C3, Bacillus safensis C6, Bacillus pumilus C7, Bacillus cereus C8 and Bacillus
acidiceler C11. Bacillus are spore forming bacteria and able to oxidise Mn after initiating spore forming (Mayanna et al., 2015). Bacillus genera belongs to -proteobacteria which grow at low nutrients availability and play a key role in biogeochemical cycles for their energy. Several other MOB have also been reported from other bacterial phyla, like actinobacteria, and proteobacteria (Santelli et al., 2014; Akob et al., 2014), and some studies on biogenic manganese oxides production in laboratory by using different bacterial strains including L. discophora SS-1 (Nelson et al., 1999), P. putida GB-1 (Tebo et al., 2005; Zhu et al., 2010), Bacillus SG-1 (Webb et al., 2005a), P. putida MnB1 (Villalobos et al., 2003) and Acremonium sp. KR21-2 (Tanaka et al., 2010).
The isolated MOB strains from Kashmir cave soil (pH 5) showed growth and Mn(II) oxidation at low pH 5. These strains also showed the ability of Mn(II) oxidation till neutral pH 7. One of the possible reason for these microbes to carried out Mn(II) oxidation on neutral and acidic pH may be due to the adaptation of these microbes to the extreme environment with in caves, with retain in their capability to grow at neutral pH as well. At low pH bacterial Mn(II) oxidation is a unique character of bacteria. In acidic condition, the Mn(II) oxidation is predicted to be thermodynamically unfavorable (Tebo, et al., 2007; Nealson, 2006). Mn oxides precipitation by bacteria at neutral or slightly alkaline pH (from 7 to 9) is thermodynamically favorable but this process is very slow in the absence of bacterial colonies. At low pH a large activation energy is required, which may not be suitable for the bacterial strains, especially Mn oxidation by microbes is not thought to provide energy for cell (Tebo et al., 2004).
Mn(II) oxidation is enzymatically controlled reaction (Ehrlich, 1968). From the present study it was revealed that all the Kashmir cave isolates oxidized Mn(II) enzymatically not chemically. By using the molecular biological techniques in the field of Mn(II) oxidation, the report was led to identification of involvement of many genes in Mn(II) oxidation. From this reports it was the first report of using same tools by different bacteria in Mn(II) oxidizing process (Brouwers et al., 2000). Reports from the three Mn(II) oxidizing bacteria suggest that multicopper oxidases (MCO) play a key role in the Mn(II) oxidizing systems in bacterial strains. The sequence of the Mn(II) oxidizing protein have similarity to MCO enzyme were involved in Mn(II) oxidation (Larsen et al., 1999). Geszvain et al., (2013) reported a cluster of gene from P. putida GB-1 which regulates the oxidation of Mn(II). The reported clusters were composed of PputGB1_2447 and PputGB1_2665
which encode two MCO enzymes, each one can independently oxidize Mn(II) and Mn(III). In different Alphaproteobacteria specie other pathways of Mn(II) oxidation are observed other than MCO enzyme like A. manganoxydans SI85-9A1 is catalyzed by mopA, heme peroxidase like enzyme (Anderson et al., 2009), while superoxide pathway is using by Roseobacter sp. AzwK-3b (Learman et al., 2011).
Metals ions affect the growth and Mn (II) oxidation capability of microorganisms (Miyata et al., 2007). We examined the metals tolerance of the Bacillus pumilus C3, Bacillus safensis C6, Bacillus pumilus C7, Bacillus cereus C8 and Bacillus acidiceler C11. All the five strains grew and oxidized Mn(II) in the presence of Ca+2 while the growth and oxidation ability was very low in the presence of zinc (Zn) and copper (Cu) ions in initial incubation time but after incubation of 16hr these strains start growth and Mn(II) oxidation. In Kashmir cave soil calcium is most abundant element (21.11% wt), while Zn and Cu are in less quantity (0.047 and 0.032 wt%). However, the isolated strains tolerated higher concentration of metals than what typically inhibits Mn(II) oxidation by model MOB strain Leptothrix discophora SS-1 [Zn(II) at 10 µM and Cu(II) at 100 µM] (Miyata et al., 2007). From this report we suggest that Kashmir cave Mn oxidizing bacteria are well adapted to the metals contaminated environment.
Calcium ion have a dramatic effect on the growth and Mn(II) oxidation by Kashmir cave isolates. The Ca+2 ions bind to the MnxG enzyme on the spore coat of Bacillus sp.SG-1 at low pH which affects the conformation and activity of enzymes which conclude the role of Ca+2 in MCO. The Mn(II) oxidation rate in the presence of Ca+2 is two and a half time higher in magnitude than that of Ca+2 free condition
Conclusions
• The Kashmir cave was first time explored microbiologically. • Total 34 bacterial strains were isolated from different samples on the basis of colony morphology. Of these, 4 showed antimicrobial activity against Gram positive, Gram Negative and clinical and non-clinical isolates. • The FTIR results demonstrated that antimicrobial metabolites produced by the selected bacterial strain resembled Bacitracin. • Optimum conditions recorded for antibiotic production were 35oC, pH 5, incubation time of 48 hrs and Nutrient broth as growth and production medium. • The study isolates (Serratia sp. KC1-MRL, Bacillus licheniformis KC2-MRL, Bacillus sp. KC3- MRL and Stenotrophomonas sp. KC4-MRL) showed ability to degrade polyethylene plastic. • The isolates were also able to produce valuable commercial enzymes including; Proteases, Lipases, Amylases • The geochemical analysis revealed that the samples were rich in different elements and compounds with the most abundant element detected was Ca followed by Fe. • The microbes were able to mineralize different polymeric forms of Ca, these forms are calcite, vetarite and argonite. • Reports for mineralization of vetarite are very limited; our isolates are very efficient in vetarite mineralization. • Study isolates were also potent Mn oxidizers. Enhance oxidation of Mn was recorded in the presence of Ca.
Future Prospects
1. Application of cavernicolic compounds in different industrial fields 2. There are more than 50 caves in Pakistan, they need to be explored microbiologically, geochemically and geomicrobiologically. 3. Need deep metagenomic studies for the detection of more potent cavervicoles. 4. Search for novel isolates which have potential role in biogeochemical processes in biomineralization. 5. Simulation studies for extraterrestrial life signatures
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