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

through out the world

13

4 Different caves are studied for microbial diversity in different

countries

37

5 List of minerals obtained from smast-5 floor and smast-7 wall of

Kashmir smast samples

72

6 Concentration of elements in sample collected from cave floor and

outside cave soil (control)

72

7 Viable Cell Count of bacterial consortium in different media

compositions incubated at 37°C.

109

8 Mn(II) oxidizing Bacteria Isolated from Kashmir cave soil and

speleothem.

186

List of Figure

S.No. Title

Page

No.

1 Different types of caves and its formation including dissolution and

weathering.

4

2 Solutional cave formation 5

3 Limestone cave formation 5

4 Distribution of major groups of microbial communities in cave

environments by 16S rRNA gene sequencing

12

5 The process of calcium carbonate precipitation by bacteria 30

6 Bacteria serving as nucleation site for CaCO3 precipitation in the

sand particle

31

7 Distribution of major groups of microbial communities in cave

environments by 16S rRNA gene sequencing

36

8 Kashmir cave (smast), Nanseer Buner, Khyber Pakhtunkhwa,

Pakistan. White arrows show location of the cave, black arrow

shows entrance to the cave.

65

105

9 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)

71

10 Quantitative analysis of minerals, A. Wall soil sample smast-7, B.

Floor soil sample smast-5

73

11 Infrared spectra of Smast-7 wall 73

12 TGA (Thermogravimetric Analysis) plots of Kashmir smast

(sample-5 Floor and sample-7 wall)

74

13 FE-SEM micrograph & EDS spectra of (a) smast-7 wall and (b)

smast-5 floor

75

14 Nutrient agar plate showing the zones of inhibition against the

clinical isolates

76

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

antimicrobial activity of B. licheniformis KC2-MRL against M.

luteus, S. aureus, Klebsiella and E. coli

79

17 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.

80

18 Comparison of FTIR spectra of control (Bacitracin) and the

antibacterial compound produced by B. licheniformis KC2-MRL

81

19 Fourier-transform infrared spectra from control and different media

after incubation at 37°C for one month

112

20 Scanning electron microscopy of low-density polyethylene samples

under specified treatment after incubation with bacterial consortia at

37°C for one month.

114

21 Limestone cave (Kashmir Smast Pirsai Mardan, Pakistan) 126

22 Evolutionary relationships of taxa Lysinibacillus sphaericus KC5-

MRL.

130

23 Effect of size of inoculum on the growth of Lysinibacillus

sphaericus KC5-MRL and production of lipase.

131

24 Lipolytic, amylolytic and proteolytic activity of Lysinibacillus

sphaericus KC5-MRL at 30°C

132

25 Stability of crude extracts of lipase, amylase and protease of

Lysinibacillus sphaericus KC5-MRL at different pH

134

26 Stability of crude extracts of lipase, amylase and protease of

Lysinibacillus sphaericus KC5-MRL at different temperature

136

27 Effects of detergents on the lipolytic, amylolytic and proteolytic

activity of isolate Lysinibacillus sphaericus KC5-MRL

137

28 Effects of metal ions on lipase, amylase and protease activity of

isolate Lysinibacillus sphaericus KC5-MRL

138

29 Effects of organic solvents on lipase, amylase and protease activity

of Lysinibacillus sphaericus KC5-MRL

139

30 Effect of inhibitors on lipase, amylase and protease activity of

Lysinibacillus sphaericus KC5-MRL

139

31 Molecular Phylogenetic analysis by Maximum Likelihood method 154

32 Calcium precipitates induced by bacteria in crystals form A)

Paracoccus Limosus, B) Brevundimonas naejangsanensis.

155

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

incubation.

156

36 Electron microscopy at different wavelength (A) at 500 μm, (B) at

200 μm, (C) at 100 μm, (D) 50 μm, (E) 1 micro meter.

157

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

Mn600, 30oC and No Mn600, 25oC), 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)

190

49 Growth curves of Bacillus Safensis C6 at 30oC and 25oC (No

Mn600, 30oC and No Mn600, 25oC), 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).

191

50 Growth curves of Bacillus pumilus C7 at 30oC and 25oC (No

Mn600, 30oC and No Mn600, 25oC), in the presence of MnCl2

(Mn600, 30oC), after adding 1 and M of Super Oxides Dismutase

192

(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,

30oC and No Mn600, 25oC), 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).

193

52 Growth curves of Bacillus acidiceler C11 at 30oC and 25oC (No

Mn600, 30oC and No Mn600, 25oC), 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).

194

53 Variation in Mn(II) oxidation at different pH and Ca+2 ion

concentration.

195

54 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 are from 4 h to 36 h.

196

197

198

199

55 Effect of metals on Mn oxide production by cavernicoles after 24 h

of incubation.

200

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

of 25oC 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.

FeS2 + 3.5O2 + H2O Fe2+ + 2SO42- + 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).

4Fe2+

+ O2 + 4H+ → 4Fe3+

+ 2H2O

FeS2 + 14Fe

3+ + 8H

2O → 15Fe

2+ + 2SO

4 2- + 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 +2H

2S → CH

2O+2S+H

2O

2CO2 + H

2S + 2H

2O → 2(CH

2O) + H

2SO

4

Oxidation of Sulfur and Related Compounds

2S+3O2+2H-OH → 2H

2SO

4

(Thiobacillus thiooxidans)

12FeSO4+3O

2+6H-OH → 4Fe

2(SO

4)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

Cave Name Length

(km) Location Discovery Reference

Mammoth cave 651.8 Kentucky, USA 1791

Gulden, 2016

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

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,

Cytophagal/Flexibacter/Bacteroides, Acidobacterium,

Actinobacteria and green-sulfur bacteria

Schabereiter-Gurtner et al. 2002

Cuezva et al, 2009

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

Park New Mexico, USA

α, β and γ-Proteobacteria, Enterobacteriaceae,

Xanthomonas, Bacillus/Coliform group and

Lactobacillaceae

Northup et al. 2003

Llonin La Garma Asturias, Northern Spain Proteobacteria, Actinobacteria, Gram-positive bacteria Schabereiter-Gurtner et al. 2004

Kartchner Caverns Arizona, USA Bacillus, Brevibacillus, Rhizobium, Sphingomonas,

Staphylococcus, Pseudomonas, and other uncultured

β-Proteobacteria

Ikner et al. 2006

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

interact with them and release of Ca2+, Na+, Mg2+, K+ and SO42- 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;

CaCO3 (solid) + H2O + CO2 (gas) <--------->Ca2+ (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)

CaCO3 (s) + H2CO3(aq) ↔ Ca2++ 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).

CaCO3 (s) + H+ ↔ Ca2+ + HCO3------(4)

CaCO3 (s) + H2O ↔ Ca2+ + HCO3- + OH------(5)

CaCO3 (s) + CO2 (aq) + H2O ↔ Ca2+ + 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

H2CO3 2H+ + 2CO3-2

NH3 + H2O NH+4 + OH-

Ca+2 + CO3-2 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+

Cl− + HCO3− + NH3 → NH4Cl + CO32−

Cell−Ca2+ + CO32−→ 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.

2Mn+2+O2(g)+2H2O → 2MnO2(s)+4H+

4Fe+2+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

novel microorganisms

Holmes et al., 2001

Altamira Cave Spain Proteobacteria, Plantomycetales,

Cytophagal/Flexibacter/Bacteroides,

Acidobacterium, Actinobacteria and

green-sulfur bacteria

Schabereiter-Gurtner

et al., 2002 Cuezva et

al., 2009

Reed Flute Guilin Guangxi

China

Knoellia sinensis and K.

subterranea (Actinobacteria)

Groth et al., 2002

Lower Kane

Cave

Wyoming, USA ε-Proteobacteria Engle et al., 2002

Lechuguilla

Spider

Carlsbad Caverns

National Park New

Mexico, USA

α, β and γ-Proteobacteria,

Enterobacteriaceae, Xanthomonas,

Bacillus/Coliform group and

Lactobacillaceae

Northup et al., 2003

Llonin La

Garma

Asturias, Northern

Spain

Proteobacteria, Actinobacteria,

Gram-positive bacteria

Schabereiter-Gurtner

et al., 2004

Kartchner

Caverns

Arizona, USA Bacillus, Brevibacillus, Rhizobium,

Sphingomonas, Staphylococcus,

Pseudomonas and other uncultured

β-Proteobacteria

Ikner et al., 2006

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 4 th 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)

Soil samples Metals (mg/kg)

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-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)

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)

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)

Cluster I

Fig. 3.1.8. Phylogenetic tree of all four species with related sequences in NCBI

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)

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)

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)

Cluster II

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).

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

0

5

10

15

20

25

30

35

40

45

KC1 KC2 KC3 KC4

VA30

NA30

CTX30

Amp25

Amc30

IPM10

Met5

CN10

LEV(5)

Zon

e o

f In

hib

itio

n (

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

around 1020 cm-1 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

mineral salt medium (g L−1): [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 (days)

MSM + PE + bacterial consortium

MSM + PE + calcium carbonate + bacterial consortium

MSM + Glucose + PE + bacterial consortium

MSM + glucose + calcium carbonate + PE + bacterial consortium

MSM + glucose + calcium carbonate + yeast extract + PE + bacterial consortium

MSM + calcium carbonate + yeast extract+ PE + bacterial 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).

Fig.

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).

Lysinibacillus sphaericus(JN377786)

Lysinibacillus sphaericus(JN377784)

Lysinibacillus sphaericus(JN377785)

Lysinibacillus fusiformis(FJ418643)

Lysinibacillus sphaericus(GU204967)

Lysinibacillus sp(HM222673)

Lysinibacillus sp(AM910304)

Lysinibacillus sp(AB689752)

Lysinibacillus sp(JN695724)

Lysinibacillus sp(JX566617)

Lysinibacillus sphaericus-(EU880531)

Lysinibacillus sphaericus(CP000817)

Lysinibacillus sphaericus(FJ544252)

Lysinibacillus sphaericus KC5-MRL (KF010827)

31

85

43

64

20

12

8

11

14

2

2

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

were observed at temperature and pH of 25oC 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

react with the CO32− 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

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

A B

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).

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

A B

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.

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.

A B

C D

E

Fourier transform infrared spectroscopy (FTIR) analysis

Calcium carbonate purchased from sigma was used as a reference with bacterial precipitated

calcium carbonate sample. The control exhibited characteristic peaks of CaCO3 at 871cm-1, 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).

20 30 40 50 60 70 80

0

100

200

300

400

500

600

700

800

In

ten

sity (

arb

.un

its)

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.

A

C

v

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

saturation index to [Ca2+]/[CO32−] 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

of aerobic mineralization of organic compounds, CO2, H2O, NO3¯, SO42¯, PO4

3¯, whereas

anaerobic mineralization results in the production of CH4, CO2, NH3, H2S, PO42¯ (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).

Mn (II) + O−2 + 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

in Bacillus sp., whereas K+, Na+, Sr2 and NO3- ions had no effect. At 10 mM Ca2+ 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

(–COOH) and finally complete mineralization to produce CO2, H2O, NO3-, SO4

2- 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).

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 1 st + 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).

Bacillus pumilus (KR780404)

Bacillus pumilus (KR528376)

Bacillus pumilus (KR780437)

Bacillus pumilus (CP000813)

Bacillus pumilus (KT273321)

Bacillus pumilus (KT624615)

Bacillus pumilus (KT624616)

Bacillus safensis (KP235236)

Bacillus pumilus (KP235237)

Bacillus pumilus (KP235238)

Bacillus safensis (KT719214)

C3

C6

Bacillus pumilus (LN890660)

C7

Bacillus acidiceler (KU254664)

Bacillus acidiceler (LN890177)

C11

C8

Bacillus cereus (KT720292)

Bacillus cereus (KT720291)

Bacillus cereus (KT719760)

100

100

98

100

0.005

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

No Mn 600, 25oC), in the presence of MnCl2 (Mn600, 30oC), 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

No Mn600, 25oC), in the presence of MnCl2 (Mn600, 30oC), 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

No Mn600, 25oC), in the presence of MnCl2 (Mn600, 30oC), 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

No Mn600, 25oC), in the presence of MnCl2 (Mn600, 30oC), 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

and No Mn600, 25oC), in the presence of MnCl2 (Mn600, 30oC), 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).

Fig. 4.11. Variation in Mn(II) oxidation at different pH and Ca+2 ion concentration.

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

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

Mn

ox

ida

tio

n r

ate

(µ

M/h

)

Ca+2 conc (µM) in pH

Mn Oxidation at different pH and Ca+2 concentration

B. pumilus C3

B. Safensis C6

B. pumilus C7

B. cereusC8

B. acidiceler C11

(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).

Fig. 4.13. Effect of metals on Mn oxide production by cavernicoles after 24 h of incubation.

0

0.2

0.4

0.6

0.8

1

1.2

Mn Mn+Ca Mn+Zn Mn+Cu

O.D

(620)n

m

Mn+Metals

Effect of metals on Mn oxidation

B. pumilus C3

B. safensis C6

B. pumilus C7

B. cereus C8

B. acidicelar C11

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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