universita' degli studi di verona

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I shows the EDAX spectra for area (figure A1-3, B1-3) and II shows the EDAX ..... properties similar to those of sulfur and tellurium (its periodic table adjacent.

UNIVERSITA’ DEGLI STUDI DI VERONA DEPARTMENT OF BIOTECHNOLOGY DOCTORATE SCHOOL OF LIFE AND HEALTH SCIENCES

PhD in: MOLECULAR, INDUSTRIAL AND ENVIRONMENTAL BIOTECHNOLOGIES CYCLE XXVIII

Study of the biogenic potential of nanoparticle formation from selenite and tellurite by two environmental strains of Burkholderia fungorum and assessment of their resistance as planktonic cells or biofilms to polyaromatic hydrocarbons S.S.D. BIO/19 Coordinator: Prof. Roberto Bassi Tutor: Prof. Giovanni Vallini Co Tutor: Dr. Silvia Lampis PhD Candidate: Nazanin Seyed Khoei

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ACKNOWLEDGEMENT I thank the collaboration of Prof. Vallini’s Group in Verona (E. Zonaro, M. Andreolli and A. Bulgarini). Part of this work was carried out while I was at the University of Calgary. I would like to thank Prof. Turner and his Group in Calgary (Dr. Lemire, Dr. Demeter and Dr. Bay) for many useful discussions. I also thank Mr. Bernardi and Ms. Benati for technical support for electron microscopy analyses.

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TABLE OF CONTENTS ACKNOWLEDGEMENT ......................................................................................2 TABLE OF CONTENTS .......................................................................................3 TABLE OF FIGURES ............................................................................................ 6 ABBREVIATION ...................................................................................................9 LIST OF PUBLICATIONS AND CONFERENCE PRESENTATIONS ........10 ABSTRACT ...........................................................................................................12 INTRODUCTION.................................................................................................14 1

Pollution Endangering Environment (Organic and non-Organic) ....................... 14

1.1

Organic Pollutants .................................................................................................... 14

1.1.1

Polycyclic Aromatic Hydrocarbons (PAH), Heterocyclic Compounds and their

toxicity14 1.1.2

Bacterial Transformation of DBT, Tolerance and Bioremediation ....................... 16

1.1.3

Biofilm ................................................................................................................... 20

1.2

Non-Organic Pollutants ............................................................................................ 22

1.2.1

Selenium ................................................................................................................ 24

1.2.1.1

Selenium Sources................................................................................................ 24

1.2.1.2

Selenium Deficiency ........................................................................................... 25

1.2.1.3

Selenium Toxicity ............................................................................................... 26

1.2.1.4

Cycling of Selenium ........................................................................................... 26

1.2.1.5

Microbial Transformation in Selenium Cycling ................................................. 28

1.2.1.5.1

Bacterial Reduction of Selenium Oxyanions ................................................... 29

1.2.1.5.2

Bacterial Oxidation of Selenium...................................................................... 31

1.2.1.5.3

Bacterial Methylation of Selenium Compounds .............................................. 32

1.2.1.5.4

Bacterial Demethylation of Methylated Selenium Compounds ...................... 33

1.2.2

Tellurium ............................................................................................................... 34

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1.2.2.1

Tellurium Sources and Use ................................................................................. 34

1.2.2.2

Tellurium Use in Medicine ................................................................................. 35

1.2.2.3

Tellurium Toxicity .............................................................................................. 35

1.2.2.4

Cycle of Tellurium .............................................................................................. 35

1.2.2.5

Microbial Reduction in Tellurium Cycling......................................................... 36

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Possible Exploitation of Microbial Processes for the Treatment of Se and Te

Polluted Matrices ............................................................................................................. 38 3

Nanoparticles.............................................................................................................. 40

4

Burkholderia Genus ................................................................................................... 43

OBJECTIVES ....................................................................................................... 45 MATERIALS AND METHODS ......................................................................... 47 1

Strains ......................................................................................................................... 47

2

Chemicals.................................................................................................................... 47

3

Study of Transformation of Organic Hydrocarbons and Tolerance toward them 48

3.1

Transformation of DBT ............................................................................................ 48

3.2

Biofilm Formation .................................................................................................... 48

3.3

Counting CFUs of Bioflm for Making the Growth Curve ........................................ 48

3.4

Morphologic Analysis of Biofilm ............................................................................. 49

3.5

Evaluating Tolerance ................................................................................................ 49

3.5.1

Tolerance of Biofilm in the Presence of DBT ....................................................... 49

3.5.2

Tolerance of Biofilm in the Presence of PAHs ...................................................... 49

3.5.3

Tolerance of Planktonic cells Grew in the Presence of DBT and PAHs ............... 50

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Study of Burkholderia Strains in the Presence of Metalloids ................................. 50

4.1

Minimum Inhibitory Concentration .......................................................................... 50

4.2

Microbial Growth ..................................................................................................... 50

4.3

Capability of Strains to Reduce Selenite and Tellurite ............................................. 51

4.3.1

Determination of SeO3 2- Amount .......................................................................... 51

4.3.2

Determination of TeO3 2- Amount .......................................................................... 51

4.4 4.4.1

Exploration of the Mechanism .................................................................................. 52 Finding the Bacterial Compartment in which Reduction Occurs .......................... 52

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4.4.1.1

Subcellular Fractions .......................................................................................... 52

4.4.1.2

Extracellular Fractions ........................................................................................ 52

4.4.2

Selenite and Tellurite Reduction Activity Test ...................................................... 52

4.4.3

Reduction in the Presence of BSO ......................................................................... 53

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Assessment of Elemental Selenium and Nanoparticles (Se, Te) ............................ 53

5.1

Measurement of Produced Elemental Selenium Amount ......................................... 53

5.2

Separation of Biogenic Se and Te Nanoparticles (SeNPs and TeNPs)..................... 54

5.3

Morphology, Localization and Characterization of Nanoparticles ........................... 54

5.3.1

Scanning Electron Microscopy (SEM) .................................................................. 54

5.3.2

Transmission Electron Microscopy (TEM) ........................................................... 54

5.3.3

UV–visible Spectral Analysis of Nanoparticles .................................................... 55

5.3.4

Dynamic Light Scattering (DLS) and Zeta Potential Analysis of Nanoparticles .. 55

RESULTS AND DISCUSSION ...........................................................................56 1

Transformation of DBT ............................................................................................ 56

2

Biofilm Formation ..................................................................................................... 61

3

Evaluating Tolerance................................................................................................. 64

3.1

Tolerance of Biofilm and Planktonic cells in the Presence of DBT ......................... 65

3.2

Tolerance of Biofilm and Planktonic cells in the Presence of PAHs........................ 67

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Study of Burkholderia Strains in the Presence of Metalloids ................................. 69

4.1

Minimum Inhibitory Concentration .......................................................................... 69

4.2

Microbial Growth ..................................................................................................... 71

4.3

Capability of Strains to Reduce Selenite and Tellurite ............................................. 73

4.4

Exploration of the Mechanism.................................................................................. 78

4.4.1

Selenite and Tellurite Reduction Activity Test ...................................................... 78

4.4.2

Reduction in the Presence of BSO ......................................................................... 80

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Assessment of Elemental Selenium and Nanoparticles (Se, Te) ............................ 85

5.1

Measurement of Produced Elemental Selenium Amount ......................................... 85

5.2

Morphology, Localization and characterization of Nanoparticles ............................ 88

CONCLUSION .....................................................................................................98 REFERENCES ....................................................................................................101

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TABLE OF FIGURES Figure 1: Structure of 16 PAHs on the EPA priority pollutant list ................................................ 16 Figure 2: Kodama pathway showing the degradation of DBT to HFBT. (I) dibenzothiophene; (II) cis-1,2-dihydroxy-1,2-dihydrodibenzothiophene; (III) 1,2-dihydroxy-dibenzothiophene; (IV) cis-4[2-(3-hydroxy)-thionaphthenyl]-2-oxo-3-butenoate;

(V)

trans-4[2-(3-hydroxy)-

thionaphthenyl]-2-oxo-3-butenoate; (VI) 3- hydroxy-2-formyl benzothiophene (HFBT). ... 18 Figure 3: Developmental of bacterial biofilm cycle ....................................................................... 22 Figure 4: Selenium cycle................................................................................................................ 28 Figure 5: Mechanisms involved in selenium cycle. ....................................................................... 31 Figure 6: Tellurite reduction mechanisms. ..................................................................................... 38 Figure 7: Burkholderia, a rod-shaped bacterium. ........................................................................... 44 Figure 8: Degradation of 500 mg l-1 of DBT and growth of bacteria. A is B. fungorum DBT1, B is B. fungorum 95. Three factors were measured together. Degradation of DBT (triangle), Growth using DBT as carbon source in DM medium (black circle), DBT concentration in control DM medium without bacteria (dotted line with star), and growth in control DM medium without DBT (dashed line). Results are expressed as the mean with standard deviation. Different letters in degradation curves are statistically different (P < 0.05, Tukey’s test). .............................. 58 Figure 9: Change of color in bottles containing DM medium with 500 mg l-1 of DBT for 95 (B), DBT1 (C) after 72 hours. A is control with bacteria but no DBT (both strains showed same results for control). ................................................................................................................ 61 Figure 10: Panel A: Growth curves of biofilm formation. B. fungorum DBT1 is circle and B. fungorum 95 is square. Panel B: CLSM analysis for biofilm of strains after 72 hours. A is 95 and B is DBT1. ...................................................................................................................... 64 Figure 11: Growth of planktonic cells (grew with DBT) and exposure of biofilm cells to DBT (after 72 h). Planktonic cells of strain 95 (black circle), planktonic cells of strain DBT1 (white square), biofilm of strain 95 (white circle), biofilm of DBT1 (black square). Results are expressed as the mean of 3 measurements and standard deviation. Different letters and symbols are statistically different (P < 0.05, Tukey’s test). ................................................................. 66 Figure 12: CLSM analysis for biofilm of strains after 72 h of exposure to DBT. A is strain 95 and B is strain DBT1. A1 and B1 are exposed to 8 mg l-1 of DBT, A2 and B2 are exposed to 128 mg l-1 of DBT and A3 and B3 are exposed to 2048 mg l-1 of DBT. .......................................... 66 Figure 13: Growth of planktonic cells (grew with PAH) and exposure of biofilm cells to PAH (after 72 h). Planktonic cells of strain 95 (black circle), planktonic cells of strain DBT1 (white square), biofilm of strain 95 (white circle), biofilm of strain DBT1 (black square). Results are

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expressed as the mean of 3 measurements and standard deviation. Different letters and symbols are statistically different (P < 0.05, Tukey’s test). ................................................................ 68 Figure 14: CLSM analysis for biofilm of strains after 72 h of exposure to PAH. A is strain 95 and B is strain DBT1. The structure didn’t change with increasing concentration of PAH. ........... 69 Figure 15: Growth of B. fungorum 95 (A) and B. fungorum DBT1 (B) on Nutrient Agar plates without salts (A, B) or with 2.0 mM sodium selenite (A1, B1) and with 0.2 mM potassium tellurite (A2, B2) ................................................................................................................... 70 Figure 16: Minimum inhibitory concentration for selenite (A) and tellurite (B). circle for B. fungorum 95 and squares for B. fungorum DBT1 ................................................................ 71 Figure 17: Growth of B. fungorum 95 (A) and B. fungorum DBT1 (B) as control (dot line), in the presence selenite (A1 and B1) and tellurite (A2 and B2). Growth in the presence of 0.5 mM selenite (dash line), in the presence of 2 mM selenite (gray line). Growth in the presence of 0.1 mM tellurite (black line), in the presence of 0.2 mM tellurite (square). ............................... 73 Figure 18: A and B: Selenite reduction in the presence of 0.5 mM selenite (A) and 2 mM selenite (B): Selenite reduction of B.95 (star) and B.DBT1 (white circle) (n=3). .............................. 77 Figure 19: Panel I: Cytoplasmic fraction for both strains fractioned in stationary phase (B. fungorum 95 (A) and B. fungorum DBT1 (B)) react with selenite (right) and tellurite (left) and electron donor (NADH (A1 and B1) and NADPH ((A2 and B2)). ..................................................... 79 Figure 20: Change of color in presence of selenite and BSO, this Picture is taken after 72 hours. Written concentrations on pictures belong to BSO. A shows strain 95 and B shows strain DBT1..................................................................................................................................... 85 Figure 21: Selenite reduction and selenium production for strain 95 (A and B) and strain DBT1 (A and C), in the presence of 0.5 mM selenite (A) and 2 mM selenite (B and C). Selenite reduction of B.95 (star) and B.DBT1 (white circle) selenium formation of B.95 (white triangle) and B.DBT1 (black circle) (n=3). ................................................................................................ 87 Figure 22: Spectrum of Se nanoparticles produced by B. fungorum 95 (left) and B. fungorum DBT1 (right) after 24 (dot line), 48 (line) and 72 (dash line) hours. ................................................ 88 Figure 23: SEM analysis for B. fungorum DBT1 (A-A3) and B. fungorum 95 (B-B3) showing spheric elemental SeNPs (shown by circles) and rod-shaped bacterial cells. A and B are controls without selenite. A1 and B1 are strains grown with 2 mM selenite after 24 hours, A2 and B2 are after 48 hours and A3 and B3 are after 72 hours. Graphs are EDAX spectra for both strains with selenium (I with Se peak shown by an arrow) and control (II without Se peak).92 Figure 24: TEM analysis for both strains with 2 mM selenite after 24 hours. B. fungorum DBT1 is A and C. fungorum 95 is B and D. Circle shows nanoparticles outside of bacterial cells and dead cell. Arrows show nanoparticles produced inside of cells. ........................................... 93

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Figure 25: DLS analysis and Zeta potential: A is the size distribution of SeNPs from B. fungorum 95 and B is the size distribution of SeNPs from B. fungorum DBT1 cultures. C and D are Zeta potential of SeNPs generated by B. fungorum 95 (C) and B. fungorum DBT1 (D) cultures. 94 Figure 26: SEM analysis for B. fungorum DBT1 (A-A3) and B. fungorum 95 (B-B3). A and B are controls without tellurite. A1 and B1 are strains grown with 0.2 mM tellurite after 24 hours, A2 and B2 are after 48 hours and A3 and B3 are after 72 hours. Graphs are EDAX spectra for both strains with tellurium (I and II are with Te peak shown by arrows) and control (III without Te peak). I shows the EDAX spectra for area (figure A1-3, B1-3) and II shows the EDAX spectra for white spots shown with circles in figure A3 and B3. ........................................... 95 Figure 27: TEM analysis for both strains with 0.2 mM tellurite after 24 hours. B. fungorum DBT1 is A and B. fungorum 95 is B. Arrows show needle like nanoparticles localized inside of cells and square shows dead cells along with nanoparticles released outside of bacterial cells by cell lysis........................................................................................................................................ 96 Figure 28: DLS analysis and Zeta potential: A is the size distribution of TeNPs from B. fungorum DBT1 and B is the size distribution of TeNPs from B. fungorum 95 cultures. C and D are Zeta potential of TeNPs generated by B. fungorum DBT1 (C) and B. fungorum 95 (D) cultures.97

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ABBREVIATIONS AO, Acridine Orange; AOBS, Acoustic Optical Beam Splitter; ARR, Arsenate Respiratory Reductase; BSA, Bovine Serum Albumin; BSO, S-n-Butyl homocysteine Sulfoximine; CBD, Calgary Biofilm Device; CFUs, Colony Forming Units; CLSM, Confocal Laser Scanning Microscopy; CNR, National Research Council; DBT, Dibenzothiophene; DDTC, Diethyldithiocarbamate; DLS, Dynamic Light Scattering; DM, Defined Mineral Medium; DMDSe, Dimethyl Diselenide; DMDTe, Dimethyl Ditelluride; DMS, Dimethylsulfide; DMSe, Dimethyl selenide; DMTe, Dimethyl Telluride;

EPA, environment protection agency; EPS,

Extracellular Polymeric Substance; GSH, Glutathione; GSSG, Diglutathione; HFBT, 3-Hydroxy-2-Formylbenzothiophene; LCS, Leica Confocal Software; LD50, Lethal Dose; MIC, Minimum Inhibitory Concentration; NA, Nutrient Agar; NADH, Nicotinamide Adenine Dinucleotide; NADPH, Nicotinamide Adenine Dinucleotide Phosphate; NB, Nutrient Broth; PAHs, Polycyclic Aromatic Hydrocarbons; PBS, Phosphate-Buffered Saline; PCBs, Polychlorinated Biphenyls; QDs, Quantum Dots; ROS, Reactive Oxygen Species; SeCys, Selenocysteine; SefA,

Se

factor

A;

SEM,

Scanning

Electron

Microscopy;

SeMet,

Selenomethionine; SeNPs, Selenium Nanoparticles; SeOB, Selenium Oxidizing Bacteria; TeNPs, Tellurium Nanoparticles TEM, Transmission Electron Microscopy; TrxR1, Thioredoxin Reductase; TxSS, Type 1–6 Secretion Systems.

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LIST OF PUBLICATIONS AND CONFERENCE PRESENTATIONS PUBLISHED PAPER Bioremediation of diesel contamination at an underground storage tank site: a spatial analysis of the microbial community. M Andreolli, N Albertarelli, S Lampis, P Brignoli, Seyed Khoei N, G Vallini. World Journal of Microbiology and Biotechnology, 2016, 32 (1), 1-12 SUBMITTED PAPERS 

Insights into selenite reduction and biogenesis of elemental selenium nanoparticles by two environmental isolates of Burkholderia fungorum. Seyed Khoei N., Lampis S., Zonaro E., Yrjälä K., Bernardi P., Vallini G. (Submitted in New Biotechnology Journal) (2016).



Comparative response of two Burkholderia fungorum strains grown as planktonic cells versus biofilm to dibenzothiophene and selected polycyclic aromatic hydrocarbons. Seyed Khoei N., Andreolli M., Lampis S., Vallini G., Turner RJ. (Submitted in Canadian Journal of Microbiology) (2016).

CONFERENCE PROCEEDING BOOK 6th European Bioremediation Conference, Chania, Crete, Greece 2015 (summer): 

How the presence of organic pollutants and metalloids can influence dibenzothiophene -degrading bacterial strains. Seyed Khoei N., Lampis S., Turner RJ., Vallini G.

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CONFERENCE PRESENTATIONS 6th International Conference on Environmental, Industrial and Applied Microbiology, Barcelona, Spain 2015 (autumn):

▪ Insights into selenite reduction and biogenesis of elemental selenium nanoparticles by two environmental isolates of Burkholderia fungorum. Seyed Khoei N., Lampis S., Zonaro E., Vallini G. ▪

Does dibenzothiophene (DBT) and poly aromatic hydrocarbons effect biofilm versus planktonic growth of DBT degrading strains? Seyed Khoei N., Lampis S., Vallini G., Turner RJ.

6th Congress of European Microbiologists, Maastricht, Netherlands 2015 (spring): ▪ How DBT degrading strains behave in presence of organic and metalloid

pollutants. Seyed Khoei N., Lampis S., Turner RJ., Vallini G.

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ABSTRACT Here we report on two strains of Burkholderia fungorum (DBT1 and 95) and their ability in both planktonic and biofilm modes to resist high concentrations of hydrocarbons in order to be exploited in bioremediation protocols. In nature, bacteria often attach to surfaces by establishing biofilms. B. fungorum DBT1 was isolated from an oil refinery drainage, while B. fungorum 95 was isolated from the inner tissues of a hybrid poplar plant cultivated in a soil contaminated by polycyclic aromatic hydrocarbons (PAHs). The hydrocarbons tested were dibenzothiophene (DBT) as a sample of thiophenes and a mixture of PAHs, namely: naphthalene, phenanthrene and pyrene. Moreover, their ability to transform toxic metalloid oxyanions (namely selenite and tellurite) to non-toxic elemental form was evaluated. This transformation not only eradicate the toxic metalloid compounds in contaminated area, but also can be utilized in order to obtain elemental form of metalloid in the form of nanoparticles with applications in technology and medicine. Our results showed that both strains degraded high concentration of dibenzothiophene and both forms of biofilm and planktonic of bacteria resisted up to 2000 mg l-1 of this compound. Moreover, B. fungorum DBT1 showed reduction in tolerance to PAHs mixture (naphthalene 2000 mg l-1, phenanthrene 800 mg l-1 and pyrene 400 mg l-1) as biofilm and planktonic forms. In contrary, formation of biofilm helped B. fungorum 95 to resist PAHs in these concentrations while planktonic form could not resist. Confocal laser scanning microscopy pictures showed that by exposing biofilm to DBT and PAHs, the structure changes. In fact, high concentration of DBT caused the formation of aggregation in biofilm. On the other hand, the result of both strains behavior in the presence of metalloids showed that strain DBT1 was able to reduce 0.5 mM selenite and 0.1 mM tellurite, while strain 95 reduced more than 1 mM selenite and 0.05 mM tellurite to elemental forms within 96 hours of aerobic incubation. B. fungorum 95 produced 1 mM selenium in the presence of 2 mM selenite. Produced selenium nanoparticles were spherical and

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zero charged with average hydrodynamic diameter of 170 nm (for strain 95) and 200 nm (for strain DBT1). However, produced tellurium nanoparticles were needle like and positive charged with average hydrodynamic diameter of 120 nm and 170 nm for strains 95 and DBT1 respectively. Scanning and transmission electron microscopy analyses showed both extracellular and intracellular selenium nanoparticles. Selenite reduction activity test evidenced cytoplasmic enzymatic activation by accepting electron from electron donors. Since nanoparticles occurred extracellularly but they are produced intracellularly according to selenite reduction activity test, either they exit by secretion or after cell lysis. However, tellurium nanoparticles are produced and occurred intracellularly by cytoplasmic activity. In conclusion, the findings for the resistance against hydrocarbons provide new perspectives on the efficiency of using DBT-degrading bacterial strains in bioremediation of contaminated sites containing high concentration of poly aromatic hydrocarbons and thiophenes and low concentration of metalloid oxyanions of selenium and tellurium. Production of selenium and tellurium nanoparticles under aerobic conditions by strains DBT1 and 95 could be due to intracellular reduction mechanisms. These biogenic nanoparticles of both kinds present size compatible with medical and technological applications which are currently under study.

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INTRODUCTION 1

Pollution Endangering Environment (Organic and non-Organic)

Throughout different categories of pollutants that can endanger the environment, hydrocarbons (organic) and metals (inorganic) have the most importance (Förstner et al. 1981). 1.1

Organic Pollutants

Hydrocarbons in different forms are organic pollutants of most concern (Chikere et al. 2011). Soil contamination by hydrocarbons (oil hydrocarbons as the major part) is getting more dominant across the world. The reason is reliance on petroleum as a main source of energy around the globe, population surge, industrialization and ignorance for environmental issues. Intentionally and unintentionally release of these carcinogenic and neurotoxic hydrocarbons in the environment causes interruption in equilibrium between living beings and their surrounded habitat, which leads to polluted terrestrial and aquatic ambiance (Holliger et al. 1997; Das and Chandran, 2010). 1.1.1

Polycyclic Aromatic Hydrocarbons (PAH), Heterocyclic Compounds and their toxicity

Polycyclic aromatic hydrocarbons (PAH) is a group of hydrocarbons comprising two or more fused benzene rings. Therefore, they have poor water solubility, high hydrophobicity and lack of degradation that result in accumulation of these compounds in environment. Their properties cause major concern for environment which makes 16 of them as priority pollutants by environment protection agency (EPA), including naphthalene, phenanthrene and pyrene (Figure 1) (Seo et al. 2009). The persistency in environment of these compounds increases with the number of aromatic rings (Conte et al. 2001; Hawrot-Paw, 2012). Naphthalene is a common pollutant of water. It can bind to molecules in liver, kidney and lung tissues and cause toxicity in these organs. Moreover, acute poisoning by naphthalene causes

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haemolytic anaemia and nephrotoxicity and ophthalmological changes in human. Naphthalene is an inhibitor of mitochondrial respiration. Phenanthrene is a photosensitizer of human skin, an allergen. Phenanthrene is also a mutagen to bacterial systems and is classified as carcinogen. Since PAHs are found as a mixture in the environment, their carcinogenic effect increases (Andreolli, 2010). Heterocyclic compounds (mostly sulfur substituted) are components of crude oil which often co-exist with PAH in the environment (Seo et al. 2009, Andreolli et al. 2011). They are highly toxic and persistent, comprising of mono-, di-, tri- and tetraalkyl-substituted thiophene, benzothiophene and dibenzothiophene (Samokhvalov, 2011). Dibenzothiophene (DBT) is the most abundant organo-sulfur compound in oil. Thus, since 1999, its use has been introduced as a reference compound in research in the field of condensed thiophene bioremediation (Mohebali & Ball, 2008).

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Figure 1: Structure of 16 PAHs on the EPA priority pollutant list

1.1.2

Bacterial Transformation of DBT, Tolerance and Bioremediation

The efficient, cost effective and environmentally friendly remediation method for hydrocarbon and metal polluted soils is bioremediation, which exploits indigenous microorganisms that can use contaminant hydrocarbons and metals as source of energy and convert them into safe products (Yousefi Kebria et al. 2009; Chorom et al. 2011; Ramos et al. 2011; Abioye, 2011). There are two known types of pathway used by bacteria in metabolism of DBT:

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1- Ring destructive pathway or Kodama pathway (Kodama et al. 1970, 1973), in which bacteria use carbon in the ring as a source of energy by hydroxylation of DBT carbon skeleton and subsequently cleavage of aromatic ring without removing the sulfur (Figure 2). This process leads to the accumulation of 3-hydroxy-2-formyl benzothiophene (HFBT) as a water-soluble end product, with a lower carbon content than DBT. The important fact about Kodama pathway is that the colorful intermediate and last product act as Kodama pathway indicators. The intermediate (trans-4[2-(3-hydroxy)-thionaphthenyl]-2-oxo-3-butenoate) is red and the last product (HFBT) is yellow, so together they produce orange color. So if we see this color in reaction medium, it means Kodama pathway is running.

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Figure 2: Kodama pathway showing the degradation of DBT to HFBT. (I) dibenzothiophene; (II) cis-1,2dihydroxy-1,2-dihydrodibenzothiophene; (III) 1,2-dihydroxy-dibenzothiophene; (IV) cis-4[2-(3-hydroxy)thionaphthenyl]-2-oxo-3-butenoate; (V) trans-4[2-(3-hydroxy)-thionaphthenyl]-2-oxo-3-butenoate; (VI) 3hydroxy-2-formyl benzothiophene (HFBT).

2- Hydrocarbon backbone conserving pathway (4S pathway or biodesulfurization) through which bacteria use sulfur as a source of energy by oxidation of sulfur atom. In fact, sulfur removing takes part without affecting the carbon skeleton (Mohebali and Ball, 2008).

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The degradation of DBT through Kodama pathway was already identified in Beijerinckia sp. B8/36 (Laborde and Gibson 1977) and Pseudomonas species (Monticello et al. 1985). Waldau and colleagues (2009) studied isolates of biphenyl utilizing bacteria capable of transforming DBT through Kodama pathway. Moreover, DBT oxidation by four strains of Pseudomonas through Kodama pathway were investigated previously (Kropp et al. 1997). Alternatively, the degradation of DBT in Sphingomonas subarctica T7b, the isolate of soil, is desulfurization (Gunam et al. 2006). Shewanella putrefaciens strain NCIMB 8768 is another example of a DBT desulfurizing microorganism (Ansari et al. 2007). DBT metabolic pathway in Mycobacterium strain G3 is proved to be 4s pathway (Okada et al. 2002). Another example of DBT desulfurizing bacteria is Paenibacillus validus strain PD2, isolated from oil contaminated soils, contains dszC gene which confirms desulfurization of DBT through 4S pathway (Derikvand et al. 2015). The key to success for bioremediation is dependent on not only the availability of target compound degrading microorganisms, but also high tolerance of bacteria to toxic compounds (Seo et al. 2009; Mandal et al. 2012; Zingaro et al. 2013). Toxic elements accumulate in the cell membrane and cause leakage of vital cellular inner components such as RNA, proteins, phospholipids (Lăzăroaie, 2010). As a result of overall cellular malfunction especially with regards to membrane proteins, bacteria die. However, toxin resistant bacteria (mostly Gram negative) have an impermeable membrane that enable them to survive in presence of toxic components (Lăzăroaie, 2010). Tolerance to toxic compounds has the restriction impact on industrial bioprocessing and bioremediation application of bacteria and only high resistant bacteria are target for these applications in practice (Zingaro et al. 2013). By conventional methods PAHs compound can be removed from environment, but such methods are expensive and spread the pollutant. Therefore, alternative environmental friendly methods are required. Bioremediation is an alternative safe

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approach which converts pollutants to less hazardous forms in a less expensive and less energy consuming way. Bioremediation is a natural process which uses indigenous microorganisms by adding them to contaminated environments in order to accelerate transformation of harmful substances to nontoxic end products (Mohan et al. 2006). There are numerous examples of application of bioremediation in management of environmental pollution. For example, Liu and colleagues (2009) used field-scale bioremediation at Shengli oilfield in northern China to treat the oily sludge. They concluded that the method was successful to decrease the toxicity of oily sludge especially by using Photobacterium phosphoreum T3. In other study, Festuca arundinace decreased toxicity of sludge by using two bioremediation technologies. Besides, THC was decreased by 5–7% (Ouyang et al. 2005). Mishra and colleagues used a bioremediation plot for one year using Acinetobacter baumannii strains and they found out that both physical and chemical properties of the soil was improved significantly (2001). Other work presented the successful bioremediation of oil by the naturally adapted Pseudomonas putida which is a promising adjunct in oil spillage operations (Raghavan et al. 1999). 1.1.3

Biofilm

When clusters of microbial cells attach to a surface, biofilms form (Figure 3). Biofilms occur in moisty environment, where there are surface and sufficient nutrient flow. They consist of a single bacterial species or multi species (bacteria, algae, fungi and protozoa) (Singh et al. 2006). It is suggested that the dominant state of microorganisms in nature is as biofilm, which provides high tolerance to various stresses (Singh et al. 2006; Gorbushina et al. 2009). Although biofilms are natural state of bacterial strains in environment, there is not enough information about formation and survival of biofilms exposed to different contaminants. There are some advantage for using biofilm instead of planktonic cells n bioremediation process which have been realized recently, including natural immobilization. Microbial cells in biofilm state are trapped in self-produced extracellular polysaccharide (EPS) of biofilm. This has an economic advantage for using

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microbial strain in bioprocess. Another advantage is robustness of microbial cells in biofilm state which makes bioremediation through biofilm a good alternative to planktonic cell due to better toleration of biofilm to toxic materials. In fact, biofilm provides a physical barrier for cells to avoid penetration of toxic compounds (Halan et al. 2012). Moreover, this structure can cope with the deficiency of culture-based techniques. Golby et al. (2011) used the Calgary biofilm device (CBD) for direct culturing of community biofilms from oil sands tailings. They demonstrated that this method enable cultivation of complex bacterial community indigenous to oil sands without problems of traditional agar plating. Most Burkholderia strains are capable of establishing biofilm in order to be resistant against chemical stress (Pibalpakdee et al. 2012; Castorena et al. 2008). There are very few studies available about tolerance of biofilm versus planktonic forms against organic pollutants. One available example is a study of tolerance of Pseudomonas pseudoalcaligenes KF707 as planktonic and biofilm in the presence of polychlorobiphenyls as an organic pollutant and some metals (Tremaroli et al. 2010). Another example is the study of tolerance and degradation capability of single and mixed microbial community in both states of planktonic and biofilm, against a mixture of naphthenic acids (Demeter et al. 2015). These authors concluded that exploiting biofilm of mixed species (which is more tolerant than planktonic state) offers a good strategy in remediation of oil sand area. Other examples of bacterial strains that produces more tolerant biofilm to some organic pollutants than planktonic cells are Zymomonas mobilis (Li et al. 2006) and Pseudomonas sp. strain VLB120ΔC-BT-gfp1 (Halan et al. 2011).

21

Figure 3: Developmental of bacterial biofilm cycle

1.2

Non-Organic Pollutants

Metals are the main non-organic pollutants of environment. They are categorized in 4 groups: 1) Essential for metabolism (Na, K, Mg, Ca, V, Mn, Fe, Co, Ni, Cu, Zn, Mo and W); 2) Toxic heavy metals (Hg, Cr, Pb, Cd, As, Sr, Ag, Si, Al, Tl); 3) Radionuclides (U, Rn, Th, Ra, Am, Tc); 4) Semi-metals or metalloids (B, Si, Ge, As, Sb, Te, Po, At, Se) and they exist in environment in cationic and anionic forms (Perpetuo et al. 2011). Generally, they co-exist with organic pollutants in environment. Investigations showed existence of metals like lead, cadmium and selenium in polluted sites (Biernacka and Małuszyński, 2006). Agricultural drainage water, sewage sludge, fly ash from coal-fired power plants, oil refineries, and mining sites are also contaminated by selenium (Hamilton, 2004). Moreover, the combustion of coal at electric generators in water reservoirs mobilize selenium and contaminate the water and soil (Lemly, 1996). Tellurium is a similar element to selenium which can also be found in polluted sites (Yurkov et al. 1996). Microorganisms use biosorption as the main mechanism for heavy metal detoxification and redox reaction to immobilize metal ions, metalloid and organometals. Prokaryotes can obtain energy by oxidizing metals like “Mn2+, Fe2+, Co2+,

22

Cu, AsO2-, Seo or SeO3 2-” and reducing “Mn4+, Fe3+, Co3+, AsO4 2-, SeO3 2- or SeO4 ” (Perpetuo et al. 2011).

2-

Two main detoxification processes for metalloids are defined: I.

Reduction or immobilization of toxic metalloid oxyanions to non-toxic elemental state: for example transformation of selenate (SeO4 2- ) and selenite (SeO3 2- ) to elemental selenium (Seº) with lower bioactivity due to being in lesser oxidation state (White et al. 1995).

II.

Methylation of metalloids, metalloid oxyanions or organo-metalloids to methyl derivatives: for example conversion of arsenate, AsO2- and methylarsonic acid to trimethylarsine.

Maiers and colleagues (1988) explained that the conversion of selenate to elemental selenium by bacteria isolated from selenium-rich site of California results in red precipitate (Seº). They described that from one week incubation of bacteria, 100 mg l-

1

of selenate was reduced (75 mg l-

1

reduced completely to red elemental

selenium). Wolinella succinogenes on the other hand was treated with high concentrations of selenate (10 mM) and selenite (1 mM). It could reduce them to elemental selenium during plateau phase of growth (Tomei et al. 1992). Likewise, immobilization of toxic Tellurite (TeO3 2-) to elemental tellurium (Teº) is a way of detoxification by bacteria because tellurite is toxic to most microorganisms at the concentration of 4µM (White et al. 1995). Reduction of tellurite to elemental tellurium followed by its deposition inside cells and near the cytoplasmic membrane was explained previously (Taylor et al. 1988; Lloyd-Jones et al. 1994). Blake and colleagues (1993) evaluated tellurite reduction by Pseudomonas maltophila strain O-2. They explained that production of black or gray color shows the presence of insoluble elemental tellurium (Teº). Rhodobacter sphaeroides, a photosynthetic bacterium, was able to grew in the presence of 0.4 mM (100 mg l‾¹) of tellurite and transform it to elemental tellurium (Moore and Kaplan, 1992, 1994).

23

1.2.1

Selenium

Selenium (Se) is a chemical element with atomic number 34. It is a metalloid with properties similar to those of sulfur and tellurium (its periodic table adjacent elements). In 1818, for the first time, it was discovered as deposits in sulfuric acid by the Swedish chemist Jӧns Jacob Berzelius who initially confused it with tellurium due to sulfur-like properties. He named it after the Greek goddess of the moon (Selene) (Weeks, 1932). Initially, it was considered as only harmful and toxic element; until, after one hundred and forty years, Schwarz and Foltz identified it as essential trace element for animal health for protection against liver necrosis (Schwartz and Foltz, 1957). Moreover, in 1979 they used it in the prevention and treatment of Keshan disease. This abnormality is a lethal cardiomyopathy which is caused by selenium deficiency (Keshan disease research group). Therefore, it is important to consider the fact that although this naturally occurring metalloid is essential to living organisms in trace quantities, it can be toxic at elevated concentrations (Mathew et al., 2014). Elemental selenium is polyatomic and has several allotropic states and can provide either crystalline or amorphous structure. Colloidal elemental selenium or so-called amorphous selenium, produce a redorange precipitate (Nuttall, 2006). Although elemental selenium can exist in different allotropic forms (crystalline, metallic, and amorphous), it has commonly been considered an unavailable form in natural environments due to its low solubility (Minaev et al. 2005). 1.2.1.1

Selenium Sources

The sources of selenium are both natural and anthropogenic. Selenium is found naturally in large extent in ores, rocks, sediments, fossil fuels, few carbon layers of the soil and in the soils of volcanic areas (Ohlendorf, 1989; Fordyce, 2013). Moreover, anthropogenic source of selenium are due to transportation of drainage water, petroleum extraction, mining exploitation and a number of raw material processes. Glass, rubber, pigment, ceramic, plastic and electronic industries are other examples. Not to mention using selenium in supplementary food for animals

24

and even medications for human. Therefore, selenium concentrations in soils range from 0.01 mg kg−1 to 1200 mg kg−1 as result of natural and accidental release (Fordyce, 2013; Winkel et al. 2015). 1.2.1.2

Selenium Deficiency

Selenium has a narrow range between the level of deficiency and toxicity. They have been estimated that intake of selenium of 30 μg×day is not enough for human health, while exceeding 900 μg×day can cause damage (Winkel et al. 2015). The reason that our body needs selenium is the presence of this element in the amino acids which contain selenium such as selenocysteine (SeCys) and selenomethionine (SeMet). SeCys is named the 21st amino acid due to the fact that it is required in a small set of selenoproteins that have redox functions (Cone et al. 1976). The function of selenium is related to selenoproteins which were first discovered by Turner and colleagues (Turner and Stadtman, 1973). Selenoproteins (namely, glutathione peroxidases and thioredoxin reductases), not only protect cells from oxidative damage, but also regulate intracellular redox state. They have other functions as well such as involvement in thyroid hormone metabolism (Fairweather et al. 2010). Nowadays, selenium is known to be an essential and critical trace element for the health and survival of mammals and humans. It is involved in varied functions such as immune function, successful reproduction, optimal brain function, protection against cardiovascular disease, aging, myopathy, osteoarthropathy, normal function of the thyroid gland, anti-viral activity, ant-oxidant and anti-carcinogenic effects. (Rayman, 2000; Hatfield et al. 2006; Papp et al. 2007). At moderate to high doses, redox active selenium compounds are able to regulate cellular redox homeostasis by generating reactive oxygen species (ROS). Consequently, increased ROS levels cause cytotoxic effects on cancer cells which are more sensitive to ROS than normal cells (Brodin et al. 2015).

25

1.2.1.3

Selenium Toxicity

Apart from the total selenium content of food sources, the chemical form of selenium is important because it affects the bioavailability of this metalloid. Different chemical forms of selenium show different properties regarding to sorption, bioavailability, mobility and toxicity (Wallschläger and Feldmann, 2010; Winkel et al. 2015). Toxicity of selenium is either acute or chronic. Different chemical forms of selenium have vastly different toxic potentials. For example, the average oral Lethal Dose (LD50) for rats is 7 mg Se/kg body weight for sodium selenite, while selenium sulfide has LD50 of 138 mg Se /kg. Elemental selenium is the safest form of selenium in biosphere with LD50 of 6700 mg Se/kg (Cummins and Kimura, 1971). Some of selenium compounds toxic effects are tachycardia, hypotension, nausea, vomiting, diarrhea, and abdominal pain. If there is also pulmonary edema, it can be a serious complication. Neurologic symptoms may also exist such as: tremor, muscle spasms, restlessness, confusion, delirium, and coma (Nuttall, 2006). Selenium intake levels from food depends on the concentration plus bioavailability of selenium forms in food sources. The latter factor itself depends on many other factors such as selenium content of soils, irrigation water and animal feed. Moreover, selenium content varies considerably across food sources and geographical origins (Rayman et al. 2008; Steinnes, 2009). Prevention of Selenium toxicity, at the same time maintenance of adequate levels of this element in dietary food will be provided by having the knowledge and control of distribution processes of selenium in agricultural soils and plants as selenium cycles across the soil-plant-atmosphere interfaces (Winkel et al. 2015). 1.2.1.4

Cycling of Selenium

This metalloid in the environment is in different chemical forms, for example, oxyanions [selenite (Se (IV) or SeO3 2-) and selenate (Se (VI) or SeO4 2-)]. Selenate

26

and selenite are highly soluble, therefore bioavailable which makes them toxic to human, plants and animals. Fortunately, the oxyanion forms of selenium can be reduced and transformed chemically or biologically to elemental form of selenium (Seº). This form has poor water solubility and stability, therefore cannot be easily taken up by living beings. Thus elemental selenium is not toxic in low dose (Nancharaiah and Lens, 2015a). However, elemental selenium in nature is occurring rarely. Elemental selenium can be reduced to selenide (Se2-) only under highly reducing conditions. Selenide is usually found bonded to metals (in rocks) and organic compounds. Additionally, selenide can also form H2Se (volatile and highly toxic) which is a structural analogue of H2S. Other forms of selenium are organoselenium compounds which contain selenocysteine (mostly) and selenomethionine in proteins, which are formed by incorporation of selenide into aminoacids. These forms can be produced by assimilatory reduction of selenite or selenate. In nature, the transformation (oxidation and reduction) of selenium is occurring both chemically and biologically. Microorganisms play a prominent role in selenium cycle in nature by carrying out both oxidation and reduction reactions (Figure 4). The evidence of selenium metabolism is found in all domains of life, including Bacteria, Archaea, Eukarya, and viruses (Nancharaiah and Lens, 2015b).

27

Figure 4: Selenium cycle.

1.2.1.5

Microbial Transformation in Selenium Cycling

Generally, there are four biological transformations of Se: reduction (assimilatory, dissimilatory (anaerobic), detoxification (aerobic)), oxidation, methylation, and demethylation. We are going to discuss each of the biological transformations of Se in detail to provide information on the important role of each mechanism in selenium cycle in the environment. It is important to know that among these mechanisms, aerobic and anaerobic reduction and methylation are significant with respect to bioremediation schemes.

28

1.2.1.5.1

Bacterial Reduction of Selenium Oxyanions

Dissimilatory reduction of selenite and selenate in the environment involves conservation of metabolic energy for microorganisms (Nancharaiah and Lens, 2015a). Some aquatic and soil bacterial strains can tolerate selenium oxyanions and reduce them to less bioavailable forms: Seº or methylated form of selenium. These strains can be utilized for bioremediation in order to decontaminate polluted ambient (Sarret et al. 2005). Selenium oxyanions are reduced by bacteria either anaerobically or aerobically by involvement of enzymes or chemically. Finally, elemental form of selenium accumulates inside or outside of bacterial cells as selenium nanoparticles (SeNPs) (Zheng et al. 2014). The mechanisms for reduction of selenium oxyanions are not completely understood. Archaea and Bacteria domains members can use selenite and selenate as terminal electron acceptors and reduce them to insoluble elemental selenium through dissimilatory reduction under anaerobic conditions (Nancharaiah and Lens, 2015a). Selenate accepts electron, catalyzed by selenate reductase, under anaerobic condition. The example is Thauera selenatis (DeMollDecker and Macy, 1993). Also nitrite reductase in bacteria can catalyze selenite reduction. The example is Rhizobium sullae (Basaglia et al. 2007). Under aerobic conditions, selenium oxyanions can also be reduced to elemental form by several bacterial strains, through either detoxification or redox homeostasis in phototrophic bacteria (Nancharaiah and Lens, 2015a). The reduction of selenite to elemental selenium was generally known to be mediated by thiols such as glutathione in cytoplasm as part of a microbial detoxification strategy (Turner et al. 1998; Kessi and Hanselmann, 2004) (Figure 5). The involved enzyme is NADPH-glutathione reductase which receive electron mainly from NADPH. Different mechanisms have been proposed for reduction of selenite (Kessi and Hanselmann, 2004). One of the reduction mechanisms is Painter-type reactions. Other mechanisms are thioredoxin reductase system, sulfide or siderophore mediated reduction and dissimilatory reduction. Although selenate reduction is environmental significant process, only a few strains can perform selenate reduction. Reduction of selenite produces 529.5

29

kJ/mol of acetate and 164 kJ/mol lactate. It leads to acetate (first equation) or pyruvate (second equation) with oxidation of lactate as electron donor.

C2H4OHCOO‾ + SeO3 2- + H+ → CH3COO‾ + Seº + HCO3‾ + H2O C2H4OHCOO‾ + SeO3 2- + H2 + 2H+ → CH3COCOO‾ + Seº + 3H2O

In this way colorless selenite is reduced to red insoluble elemental selenium (Nancharaiah and Lens, 2015b). Pearce et al. (2009) compared reduction profiles of Geobacter sulfurreducens, Shewanella oneidensis, and Veillonella atypica. All of them could transform selenite to elemental selenium with the presence of electron donor. Among them, V. atypica was the most efficient strain. G. sulfurreducens and S. oneidensis formed a crystalline intermediate. The selenium oxyanion reduction rate and formed selenium are influenced by the reduction mechanism which microorganisms use. Li proposed the reduction in S. oneidensis MR-1 is through respiratory electron transport pathway (Li et al. 2014). However, elemental selenium can be reduced microbiologically even further to soluble selenide. This form is in combination with metal ions, forming insoluble metal selenides. Moreover selenide can be emitted as H2Se, which is volatile and highly reactive. H2Se in the presence of oxygen can be oxidized immediately to elemental selenium. Organoselenium and methylated selenium also contain selenide (Nancharaiah and Lens, 2015a).

30

Figure 5: Mechanisms involved in selenium cycle.

1.2.1.5.2

Bacterial Oxidation of Selenium

Selenium-oxidizing bacteria (SeOB) can oxide selenide and elemental selenium to selenite or selenate and complete the other half of selenium cycle (Figure 4). In

31

1923, oxidation of elemental selenium was reported by autotrophic soil bacteria. Lipman and Waksman added elemental selenium to a fresh soil. They observed that elemental selenium was oxidized and caused an increase in soil acidity. This rodshaped bacterium even could use elemental selenium as a source of energy (Lipman and Waksman, 1923). Sarathchandra and Watkinson (1981) reported bacterial oxidation of elemental selenium to selenite by Bacillus megaterium. In this transformation, selenite was the dominant product, while trace amounts of selenate was also produced. Dowdle and Oremland (1998) performed a study on oxidation of elemental selenium in soil slurries to selenite or selenate. The transformation was inhibited by killing the bacteria by using autoclave or antibiotics. On the other hand, adding acetate, glucose, or sulfide boosted the oxidation of selenium by bacteria which shows the involvement of chemoheterotrophs and chemoautotrophic thiobacilli. Cultures of Thiobacillus ASN-1 and Leptothrix MnB1 and a heterotrophic soil enrichment, oxidized selenium with selenate as the major end product. Same year, Losi and Frankenberger demonstrated microbial oxidation and solubilization of precipitated elemental selenium in soil. They found out that the oxidation of selenium was correlated to pre-exposition of soil to Se. They also showed that oxidation of elemental Se in soils is largely mediated by biotic mechanisms and adding an inorganic carbon source (NaHCO3) resulted in a more efficient selenium oxidation than adding glucose. This fact implies that oxidation is chemoautotrophic. Generally, oxidation rates depend on the dissolved oxygen concentrations; consequently, the process can be different based on the environmental conditions, for example, rivers, lakes or ponds. Generally, oxidation rates are 3 to 4 times lower than those of reduction in selenium cycle. This can lead to larger amounts of elemental selenium or selenides in sediments (Nancharaiah and Lens, 2015b). 1.2.1.5.3

Bacterial Methylation of Selenium Compounds

Methylated Selenium compounds are producing from oxyanions and organo-Se compounds. This transformation is occurring in soil, sediment and water (Doran,

32

1982). Volatile dimethyl selenide (DMSe) and dimethyl diselenide (DMDSe) are the most common forms of methylated selenium with selenium in its fully reduced state (selenide) (Winkel et al. 2015). Other volatile methylation compounds are methaneselenone, and methaneselenol. Although the importance of Se methylation is not clearly understood, volatile Se compounds are diluted in atmosphere, hence are less hazardous. Mc Carty and colleagues (1993) performed study on the species of phototrophic non-sulfur bacteria and found that they contribute in selenium and sulfur cycling by producing volatile methylated compounds. Selenate was transformed to dimethyl selenide and dimethyl diselenide by Rhodocyclus tenuis and Rhodospirillum rubrum. Also selenite was transformed by R. tenuis to dimethyl selenide. The mechanism of reducing selenocysteine to hydrogen selenide is based on the reduction of glutathione via selenocysteine-glutathione selenenyl sulfide (Sayato et al. 1997). Ranjard and colleagues (2003, 2004) found two different methyltransferases in species of Pseudomonas. One is bacterial thiopurine methyltransferase. This enzyme transforms selenite and selenocysteine to dimethyl selenide and dimethyl diselenide (Ranjard and Cournoyer, 2003). The other is a homolog of calichaemicin methyltransferase which produces same products. The enzyme, MmtA, is a new group of methyltransferases which has homologs in other species of bacteria (Ranjard et al. 2004). 1.2.1.5.4

Bacterial Demethylation of Methylated Selenium Compounds

DMSe and its analogue DMS (dimethylsulfide) undergo biological demethylation reactions in the environment. Demethylation is removal of a methyl group form a methylated compound. Several soil microorganisms are capable of demethylation of volatile selenium methylated compounds. An example is demethylation of DMS by methylotrophic methanogens (Stolz et al. 2006; Vriens et al. 2014).

33

Oremland and co-workers (1989) hypothesized that methylotrophic bacteria perform demethylation, while certain hydrogen-oxidizing methanogens may be involved in reductive methylation. The importance of demethylation reactions in the environments has yet to be determined and can be implemented as a remediation technique. 1.2.2

Tellurium

Franz Joseph Mūller, who was born in 1740 in Vienna, discovered tellurium. In 1782 he extracted tellurium from a bluish-white ore of gold (called aurum problematicurn, aurum paradmum, or aurum album) in Transylvania where his father served as the treasurer. On January 25, 1798, Klaproth read an article on the gold ores of Transylvania. He reminded his hearers of the forgotten element, and suggested the name tellurium for it (Weeks, 1932). Tellurium comes from the Latin word tellus which means earth. 1.2.2.1

Tellurium Sources and Use

Tellurium is found in native form and telluride form in combination with gold or other metals. It is produced commercially from muds of electrolytic refining of blister copper in especially U.S., Canada, Peru and Japan. Tellurium is 75th most abundant elements of earth with the concentration in crust of 0.002 ppm (Bagnall, 1966; Cooper, 1971; Haynes, 2011). The appearance of crystalline tellurium is silvery-white liquid with metallic luster. Amorphous tellurium is a result of precipitating from a solution of tellurous acid. Tellurium is a semiconductor and have some applications in this matter such as blasting caps, in ceramics and photovoltaic cells (Haynes, 2011) and is popular as a coloring and propertymodifying agent in different types of glasses. It is also exploited as a reagent on silverware to produce black shade as tellurium chloride and tellurium dioxide. Elemental tellurium and tellurium diethyldithiocarbamate are vulcanizing agents which allow rubber to tolerate change in temperature and increase its half-life. Tellurium is also a catalyst in a variety of reactions. Te is also used to improve the

34

properties of cast iron, lead, and copper alloys (Zannoni et al. 2008). Other applications of tellurium are in electronics and rechargeable batteries. Also there are some biological use for example as quantum dots (QDs) for probe in biological detection (Turner et al. 2012). The use of tellurium in these industries cause more release of it in the environment and cause toxicity in living beings (Kim et al. 2013). 1.2.2.2

Tellurium Use in Medicine

Previously tellurium was used for cure of microbial infections before discovery of antibiotics. In 1926 they used it in the treatment of syphilis. Moreover, tellurite with antibiotic properties has been used in microbiology since the 1930s (Fleming, 1932; Fleming and Young, 1940). In 1984, a potential anti-sickling effects of tellurite on blood cells in the treatment of sickle cell anemia was suggested (Asakura et al. 1984). Other applications of tellurium compounds in medicine are treatment of AIDs, protection of bone marrow stem cell during chemotherapy, defense against oxidative and nitrosative stress and other applications (Zannoni et al. 2008). 1.2.2.3

Tellurium Toxicity

Despite many chemical similarities between selenium and tellurium, a nutritional role has not been identified for tellurium. Moreover, tellurium even at low concentrations induces both acute and chronic toxicity in a variety of organisms (Goulle´ et al. 2005). Generally, it is considered that the toxicity of tellurite is a consequence of its strong oxidizing properties. Also there are evidences linking tellurite toxicity to the production of ROS (Tremaroli et al. 2007). 1.2.2.4

Cycle of Tellurium

Tellurium in the environment has a same cycle as selenium in 4 inorganic states: elemental form (Teº), inorganic telluride (Te 2- ), tellurite (Te (IV) or TeO3 2- ) and tellurate (Te (VI) or TeO4 2-). Moreover, there are organic forms (methylated form) of tellurium as well, such as dimethyl telluride (DMTe) or (CH3TeCH3) and dimethyl ditelluride (DMDTe) with garlic smell. Similar to selenium, oxyanions of

35

tellurium are toxic and more common than nontoxic elemental form. The tellurium oxyanions, tellurite and tellurate, are toxic because they are strong oxidizers. Telluride is chemically reactive and tends to combine with organic compounds forming the final product of organic tellurides (Zannoni et al. 2008). 1.2.2.5

Microbial Reduction in Tellurium Cycling

The redox reaction plays a crucial role in controlling tellurium toxicity due to insolubility of Teº and therefore low bioavailability and toxicity of it. Thus, reduction of oxyanions of tellurium (Te(IV) and Te(VI)) to insoluble and less toxic Teº is an effective solution for environmental detoxifying of tellurate and tellurite. Microbes are contributing naturally to this transformation which can be exploited for bioremediation. For this, microorganisms employ direct enzymatic redox reaction or they use indirect redox active organic molecules such as quinines. However, the exact mechanism for tellurite reduction is not fully understood (Kim et al. 2013). Klett (1900) was the scientist who noticed the biochemical transformation of sodium tellurite into a black, insoluble precipitate. He presumed that to be metallic tellurium. A variety of bacterial aerobic and anaerobic phototrophs, hydrothermal heterotrophs, eukaryotes (fungi and plants) and mitochondria in animal tissues can perform this reduction leading to black tellurium precipitates (Zannoni et al. 2008). The product of the reduction of tellurium oxyanions are elemental tellurium nanoparticles (TeNPs) which are usually deposed intracellularly rather than extracellularly oppose of SeNPs (Baesman et al. 2007). Sepahei and his colleagues (2009) isolated some Bacillus species form copper mine with high-level tolerance to tellurite (1500 to 2000 mg l-1). Produced tellurium nanocrystals occurred intracellularly with different structure including nanorodes. These bacterial strains can be used to produce tellurium nanoparticles as an alternative for bench-scale synthesis. Deinococcus radiodurans reduced 70% of 40 μM potassium tellurite to elemental tellurium within 5 hours (Anaganti et al. 2014). E. coli can reduce tellurite in its cytoplasm (Pérez et al. 2007). Transmission electron microscopy (TEM) shows that TeNPs usually precipitate close to the cell wall and/or

36

lipid membranes (Zannoni et al. 2008). There are 4 types of tellurium oxyanions reduction: I. Painter-type reaction with glutathione similar to selenium counterpart (Turner et al. 2001) (Figure 6) II. Catalytic reduction by oxidoreductases (Avazeri et al. 1997) III. Redox reaction involving the siderophore pyridine-2,6- bis(thiocarboxylic acid) (Zawadzka et al. 2006) IV. Reduction (direct or indirect) by aid of terminal oxidase from the membranebound respiratory chain (Trutko et al. 2000) Due to the similarity between the chemistry of tellurium and selenium, the first three mechanisms of reduction are similar in both. However, tellurium oxyanions may act differently in site-specific interaction(s) with components of the bacterial respiratory chain (the fourth mechanism).

37

Figure 6: Tellurite reduction mechanisms.

2

Possible Exploitation of Microbial Processes for the Treatment of Se and Te Polluted Matrices

In recent years, economic boom and industrialization in developing countries has caused massive progress in modern agriculture. Such development has many environmental side effects such as pollution. One example is pouring industrial discharge and drainage waste water and material into agricultural lands. Toxic elements and metals can remain in soil and water for a long time presenting numerous health hazards to all living forms. They also decrease fertility of soil and

38

biodiversity. Typical inorganic and organic contaminants in urban areas are metals and petroleum derived products. The presence of both types on the same site demands more technical and economic force in order to decontaminate them. No doubt that biological approaches offer efficient, cheaper and safer solution to this problem. Bioremediation is the technology of using living organisms to remediate polluted matrices. They can transform or eliminate polluted substances (Pal et al. 2010). Since 90s scientists are trying to find more appropriate way for remediation of toxic Se oxyanions released to water and soil. One of the environmental friendly and low cost way is using biological systems (microbes and plants) to remove this metalloid oxyanions through bioremediation. There are several bioremedial approaches including utilizing bacteria in bioreactors for reducing toxic forms of selenium to non-toxic insoluble elemental form. In this way not only the toxic form will be removed form environment, but also the end product in form of biogenic nanoparticles have promising applications in medical and technological fields. These systems are designed to remove selenite and selenate; tellurite and tellurate from contaminated water (with origin of industrial or agricultural) before or after release into the environment. Apart from reduction, methylation of metalloid oxyanions to volatile methylated forms can be exploited as well by using methylating microorganisms. The example for successful in situ use for treatment of selenium contamination is treatment of seleniferous soils in San Joaquin Valley, California. One of the major concerns in this area is high level of selenium in ponds containing agricultural drainage water. They tried to use a bioremediation program in order to volatilize selenium to detoxify a saline seleniferous sediment of the contaminated pond. Finally, 32% of the Se was removed successfully over 22 months (Frankenberger and Karlson, 1995). There are other bioremediation studies in this matter. The first commercial use of a bioremediation was in 1972 to clean Sun Oil pipeline spill in Ambler, Pennsylvania. Since then, bioremediation became a well-developed way of cleaning up several contaminants. Ability of rhizobacteria

39

(Stenotrophomonas sp.) capable of reducing the selenium oxyanions in contaminated soils was discussed before (Pal et al. 2010). Exploitation of Thauera selenatis, a selenate-respiring bacterium, in a biological reactor for remediation purposes has been described before for decontaminating water (Macy et al. 1993). Also plants can effectively remediate Se contaminated soils and waters (phytoremediation process). For example, Terry and colleagues (1992) showed selenium volatilization in different plants in soil. There are some examples of tellurite converting bacteria such as Basnayake and colleagues (2001) work that used Pseudomonas fluorescens K27 to reduce tellurite and tellurate to dimethyl telluride and elemental tellurium in bioreactor with harvest yield of 34% for Teº in 92 hours which shows the promising application of this strain in bioremediation. Gharieb and colleagues (1999) also used bioreactor experiment with fungi (Fusarium and Penicillium citrinum) capable of tellurite biotransformation.

3

Nanoparticles

As it has been mentioned before in this thesis, the result of selenium and tellurium oxyanion reduction is elemental selenium (Seº) and tellurium (Teº) nanoparticles. SeNPs has important semiconducting and photoelectric properties. Moreover, they have some application in environmental biotechnology such as mercury capturing or in medicine (Jain et al. 2014). SeNPs have excellent biological properties of selenium ions with much lower adverse reaction due to low toxicity which makes them good candidate for pharmaceutical dosage forms instead of other forms of selenium in its cycle (Zhang et al. 2011). The selenium nanoparticles also have biologic activity due to their optimum adsorptive ability resulting from interaction between nanoparticles and functional groups of proteins (namely: NH, C=O, COO‾, and C–N). Studies on biological toxicity of selenium showed that SeNPs are

40

efficient with regard to increasing the activities of glutathione peroxidase and thioredoxin reductase enzymes (Torres at al. 2012). SeNPs are more investigated thanTeNPs; however, there are some examples of applications of TeNPs using their semiconductor and thermoelectrical properties, or antibacterial and antifungal potential (Kim et al. 2011; Zare et al. 2013; Zonaro et al. 2015). There are several studies revealing the promising application of nanoparticles in medical field: The use of SeNPs as supplements with high biological activity and low toxicity explained by Yazdi and colleagues (2012). He also mentioned that selenium nanoparticles have effects on different disorders such as malignant diseases in human. Moreover, this author showed that SeNPs produced by Lactobacillus plantarum stimulates immune response of mice with 4T1 breast cancer tumor. The mechanism is through the elevation of IFN-γ and IL-12. Same author after one year used SeNPs through oral administration to treat X-ray irradiated mice. They conclude that nanoparticles are effective for recovery of BM suppression by increasing lymphocytes and neutrophils in patients (Yazdi et al. 2013). The antioxidant effects of selenium nanoparticles (Huang et al. 2003; Torres at al. 2012; Forootanfar et al. 2014) and anticancer properties have been mentioned by different authors (Huang et al. 2003; Faghfuri et al. 2015). Not to mention several antimicrobial,

anti-parasite

and

antifungal

effects

reported

previously

(Mahmoudvand et al. 2014; Kheradmand et al. 2014; Srivastava and Mukhopadhyay, 2015a). In recent years there is an increasing interest in synthesis of nanoparticles due to their promising applications. Chemical methods of synthesizing SeNPs need high temperatures and pressures, therefore are expensive and hazardous to environment. Nowadays biological systems, mainly bacteria, are good replacement for chemical methods in order to obtain selenium nanoparticles (Zhang et al. 2011). The biogenesis of selenium using bacteria have been reported before. Oremland et al. (2004) reported the biogenesis of SeNPs under anaerobic conditions by Se-respiring bacteria, such as Sulfurospirillum barnesii, Bacillus selenitireducens, and Selenihalanaerobacter shriftii, able to convert selenite and

41

selenate to SeNPs. Also, Klonowska et al. (2005) explained selenite and tellurite reduction by Shewanella oneidensis and production of their elemental forms. However, anaerobic conditions have some limitations making biosynthesis processes challenging, such as culture conditions and isolate characteristics. Selenium-tolerant aerobic microorganisms provide the opportunity to overcome mentioned limitations (Torres at al. 2012). There are few studies dealing with biologically aerobic formation of SeNPs (Hapuarachchi et al. 2004). Pseudomonas aeruginosa and Bacillus sp. are other examples (Yadav et al. 2008; Prakash et al. 2009). The size of nanoparticles is important to have both industrial and medical applications. It has been mentioned that the size of 5–200 nm is optimum for having anti-oxidant effects for example (Torres at al. 2012). Bajaj and Winter (2014) in their study dealing with metalloid oxyanion reduction by Duganella violacienigra, concluded that their bacterial cultures could be exploited commercially to bioremediate both selenite and tellurite contaminated matrices. Moreover their strains provide the possibility of green synthesis of extracellular selenium and tellurium nanospheres. Zonaro and colleagues (2015) obtained

selenium

and

tellurium

nanoparticles

from

two

strains

of

Stenotrophomonas maltophilia SeITE02 and Ochrobactrum sp. MPV1 which possess antimicrobial and anti-biofilm properties against Escherichia coli JM109, Pseudomonas aeruginosa PAO1 and Staphylococcus aureus ATCC25923. Tanaka and colleagues (2010) used Magnetospirillum magneticum AMB-1 which is capable of crystalizing tellurium inside the cells and forming tellurium nanocristals. They mentioned that their magnetotactic strain can crystalize both magnetite and tellurium which would have a promising application in bioremediation and magnetic recovery of tellurite. There are other reports for tellurium nanoparticle biogenesis (Narayanan and Sakthivel, 2010). Srivastava and colleagues (2015b) used Halococcus salifodinae BK3 tolerant to 5.5 mM tellurite and obtained TeNPs with antibiotic effects effective towards Gram positive and negative bacteria. Zare and colleagues (2013) evaluated the antifungal activity of biogenic Te NPs against Candida

42

albicans ATCC14053. They showed that TeNPs (0.2 mg ml-1) inhibit the squalene monooxygenase enzyme and subsequently increase the expression of the ERG1 gene. Kim et al. (2014) synthesized bismuth tellurium selenide nanomaterials for thermoelectric applications. Lin et al. (2012) also prepared Te nanomaterials (NMs) with different shapes and evaluated their antibacterial activity against Escherichia coli.

4

Burkholderia Genus

Burkholderia (Figure 7) is a genus of Proteobacteria. The genus was named after Walter H. Burkholder, plant pathologist at Cornell University. There are about 82

species

in

this

genus.

The

pathogenic

members

of

this

genus

include Burkholderia mallei, responsible for glanders disease; Burkholderia pseudomallei, causative agent of melioidosis; and Burkholderia cepacia, pathogen of pulmonary infections in patients with cystic fibrosis. The Burkholderia genus was previously known as a part of Pseudomonas genus. The name of this genus refers to a group of virtually ubiquitous Gram-negative, motile, obligatory aerobic rod-shaped bacteria. The species in this genus have a wide range including both animal and plant pathogens, and some environmentally important species. In particular, B. xenovorans (which used to called B. fungorum) showed the ability to degrade chloro organic pesticides and polychlorinated biphenyls (PCBs). The use of Burkholderia species for agricultural purposes (such as biodegradation, biocontrol, and plant growth-promoting rhizobacteria) is possible after confirmation that the species do not belong to pathogenic species of Burkholderia (Burkholderia mallei, pseudomallei, and cepacia). The specie Burkholderia fungorum was proposed by Coenye and colleagues (2001) for isolates recovered from the environment, and animal and human clinical samples.

43

Figure 7: Burkholderia, a rod-shaped bacterium.

A few single strain with DBT degrading properties have been isolated; Burkholderia fungorum DBT1, isolated from an oil refinery sewage drainage showed promising biodegradation efficacy, converting DBT to soluble end product in three days (through ring destructive Kodama pathway) (Di Gregorio et al. 2004). The novel hybrid poplar endophytic strain, Burkholderia fungorum 95, is another effective strain. However, their tolerance and growth pattern in the presence of different concentrations of this organic hydrocarbon and other PAH are unknown. Additionally, their behavior in the presence of metalloids needs to be evaluated.

44

OBJECTIVES In contaminated sites DBT is found with other PAH pollutants and metals (Seo et al. 2009). To evaluate if DBT degrading strains can be used in industrial bioremediation, the growth and tolerance of these strains with DBT and PAH as biofilm should be investigated and compared to planktonic form. It is also important that these strains be able to resist and transform inorganic pollutants as well. Here two strains of Burkholderia fungorum (DBT1 and 95) were studied. DBT1 was previously isolated from oil refinery drainage in Pisa at the formerly Soil Microbiology Centre of the Italian National Research Council (CNR) (Andreolli et al. 2011); while Burkholderia fungorum 95 was obtained in axenic culture at the Department of Biosciences, University of Helsinki (Yrjälä et al. 2010). They had been preliminarily assessed for their promising capability to

degrade

dibenzothiophene (DBT) and polycyclic aromatic hydrocarbons (PAHs) (Andreolli, 2010). But this is the first time that these Burkholderia strains are exposed to high concentrations of mixture of PAHs and DBT to compare their tolerance as planktonic and biofilm forms in order to take a step ahead for using them in bioremediation protocols. The studied strains were isolated from oil refinery sewage drainage and plant grown on PAH polluted soil. Therefore, there is high probability for existence of selenium in those polluted sites as we mentioned before. Increasing amount of selenium and tellurium compounds especially toxic oxyanions in the environment cause problem for all living type of life. Thus, it’s necessary to set up effective systems for the remediation of polluted effluents such as microbial transformation by bacteria extracted from the same contaminated matrices which seems cost effective and environmental friendly choice (Fujita et al. 2002). Moreover, the product of this transformation (metalloid nanoparticles) has application in biotechnological fields. So, metalloid oxyanions of selenium and tellurium were chosen as sample inorganic pollutants to be tested with the strains.

45

Therefore, the present work is divided in two parts. The first part is focusing on evaluating the resistance of strains exposed to PAHs. The objectives of this part are to: I. Understand if these strains can build biofilm structure II. Analyze the DBT degradation profiles of strains compared to each other III. Evaluate the tolerance of strains against PAHs mix and DBT both as planktonic and biofilm IV. Visualize biofilm through CLSM to observe the effects of PAHs and DBT on the structure of biofilm. Therefore, these findings shed light on the possibility of utilizing biofilm of these bacterial strains in bioremediation of contaminated sites. The second part focuses on exposure of strains to metalloids. The reduction of selenite and tellurite by selected two strains of Burkholderia fungorum was investigated. The objectives of this work are to: I. Determine the efficiency of aerobic selenite and tellurite reduction within the strains of Burkholderia fungorum considered and to compare two strains to know which one is more effective. II. Clarify the possible mechanism(s) of reduction. Understanding selenium and tellurium metabolism pathways in Burkholderia fungorum strains improves our knowledge about the aerobic reduction mechanisms in the biogeochemical cycle of selenium and tellurium. III. Characterize the elemental selenium and tellurium nanoparticles synthesized by the bacterial strains and evaluate if they have appropriate potential properties to have application in medicine or technology which needs to be investigated in future. IV. Visualize strains with produced nanoparticles to get more information about localization, shape and size of nanoparticles.

46

MATERIALS AND METHODS 1

Strains

The two bacterial strains used in this study were Burkholderia fungorum DBT1 and Burkholderia fungorum 95 which are environmental isolates. Burkholderia fungorum DBT1 has been isolated from discharge of an oil refinery near Leghorn (Tuscany, Italy) (Andreolli et al. 2011). Burkholderia fungorum 95 isolated from surface-disinfected leaves, stems and roots of hybrid Populus tremula grown in PAH-polluted soil (Yrjälä et al. 2010).

2

Chemicals

Chemicals purchased from Sigma-Aldrich (Milan, Italy) were all analytical grade. Nutrient broth, and technological agar were provided through Oxoid Italia Spa (Garbagnate Milanese, Italy). Sodium selenite and potassium tellurite were prepared as a 100 mM and 10 mM stock solutions in deionized water respectively. After preparation they were sterilized by filtration with 0.2 µm Millipore filter. The growth media that was utilized in tolerance against polyaromatic hydrocarbons was yeast mannitol broth (YMB) composed of 0.5 g l‾¹ K2HPO4; 0.1 g l‾¹ MgSO4·7H2O; 0.1 g l‾¹ NaCl; 0.4 g l‾¹ yeast extract; 10 g l‾¹ mannitol. The colony forming units (CFUs) measure was performed on plates of yeast mannitol agar (YMA), prepared by adding 15 g l‾¹ of agar to YMB medium (Andreolli et al. 2011). Defined mineral medium (DM) without carbon source was used for DBT degradation test with the composition of 2.2 g Na2HPO4, 0.8 g KH2PO4, 3.0 g NH4NO3, 0.05 g yeast extract, and deionized H2O to one liter (Frassinetti et al. 1998).

47

3

Study of Transformation of Organic Hydrocarbons and Tolerance toward them

3.1

Transformation of DBT

DM medium supplied with 500 mg l-1 DBT was used to perform this test. During the incubation time of 72 hours on shaker at 27ºC, a spectrophotometric method used to measure remaining concentration of DBT. Two control negatives (without bacterial strain and without DBT) were set as well. For measuring the concentration of DBT, every 24 hours three samples from each flasks were taken. Then, DBT was extracted with 2 ml of ethyl acetate followed by shaking vigorously for 10 seconds and vortexing for 30 seconds. Afterwards a few drops of saturated NaCl solution was added to separate the emulsion into two phases. Finally, the absorbance of top layer was measured at 324 nm. The test was performed for triplicate samples. 3.2

Biofilm Formation

Biofilms were prepared using Calgary biofilm device (CBD) (Ceri et al. 1999). The colonies from both strains in YMA plates were suspended in to 5 ml of 0.9% NaCl matching the opacity of 1.0 McFarland standard (correlate to 3.0 × 10⁸ CFUs ml ‾¹). Then 150 µl of 30 fold of diluted saline-strain mixture in YMB medium was put to each well of a 96-well microtiter plate. Then, the CBD was put on top of the 96-well microtiter plate. The assemble device then was placed on a gyrorotary shaker (shaking with a speed of 150 rpm) for 96 hours at 25ºC (Harrison et al. 2006). 3.3

Counting CFUs of Bioflm for Making the Growth Curve

After each 24 hours of incubation, the pegs of CBD were rinsed twice in 0.9% NaCl. The following step was putting the pegs into a new 96-well microtiter plate filled in each well with 200 µl of a solution composed of 0.9% Saline and 0.1% Tween 20. The assembled device (called recovery plate) then was placed in Aquasonic 250HT ultrasonic device (VWR International, Mississauga, ON, Canada) set at 60 Hz for 30 min. Then the viability of bacterial cells were evaluated by using MBEC™ recovery protocol suggested by Harrison et al. (2010). The recovery plate was

48

diluted and spot plated on YMA petri dishes. CFUs counts were performed on YMA plates incubated for 48 hours at 25ºC. All of the standard deviations were based on triple replicates. 3.4

Morphologic Analysis of Biofilm

Some pegs form CBD were removed for confocal laser scanning microscopy (CLSM) analysis. They fixed in 200 µl of Glutaraldehyde 5% for 30 minutes at 37ºC then stained with acridine orange (AO) (0.1% w/v in phosphate buffered saline (PBS)). After staining, pegs were covered with aluminum foil to be protected from light until visualization time. Then they were put on glass slide with one drop of distilled water. Biofilms around the pegs were observed by Leica DM IRE2 spectral confocal and multiphoton microscope with a Leica TCS SP2 acoustic optical beam splitter (AOBS) (Leica Microsystems, Richmond Hill, ON, Canada). In each study, a 63 × water immersion objective was used. Images were analyzed by Leica confocal software (LCS, Leica Microsystems) (Harrison et al. 2006). 3.5 3.5.1

Evaluating Tolerance Tolerance of Biofilm in the Presence of DBT

72 hours old biofilm was exposed to a new 96-well microtiter plate containing different concentrations of DBT (2 mg l-1, 8 mg l-1, 32 mg l-1, 128 mg l-1, 512 mg l1

, 2048 mg l-1). The assemble device then was placed on gyrorotary shaker (shaking

with speed of 150 rpm) for a 72 hours at 25ºC. After incubation time, sample pegs were cut and assessed through counting the CFUs by the method used in 3.3 section. 3.5.2

Tolerance of Biofilm in the Presence of PAHs

72 hours old biofilm was exposed to a new 96-well microtiter plate containing different concentrations of a 5:2:1 mixture of naphthalene, phenanthrene and pyrene. This mixture has been previously used as a petroleum model (Cui et al. 2008a). 6 different concentrations were used, starting from 1.95 mg l-1 of naphthalene, 0.78 mg l-1 of phenanthrene and 0.39 mg l-1 of pyrene as minimum concentrations and 2000 mg l-1 of naphthalene, 800 mg l-1 of phenanthrene and 400 mg l-1 of pyrene as

49

maximum used concentrations. The assembled device then was placed on gyrorotary shaker (shaking with speed of 150 rpm) for a 72 hours at 25ºC. After incubation time, sample pegs were cut and assessed through counting the CFUs by the method used in 3.3 section. 3.5.3

Tolerance of Planktonic cells Grew in the Presence of DBT and PAHs

The normalized planktonic cells (to 1.0 McFarland standard) were inoculated in the presence of DBT in the 96-well microtiter plate containing YMA. Then, after 72 hours they were spot plated on YMA petri dishes and CFUs counted on YMA plates incubated for 48 hours at 25ºC. All of the standard deviations were based on triple replicates. The experiment was repeated in the presence of a mixture of PAHs.

4 4.1

Study of Burkholderia Strains in the Presence of Metalloids Minimum Inhibitory Concentration

The minimum inhibitory concentration (MIC) for both strains with selenite and tellurite was measured by culturing on nutrient agar plates with increasing concentration of selenite (0 to 20 mM) and tellurite (0 to 2 mM). The stock solutions which were utilized for this experiment were 100 mM of sodium selenite (Na2SeO3) and 10 mM of potassium tellurite (K2TeO3). Based on the MIC result, two concentrations of Na2SeO3 (0.5 and 2.0 mM) and two concentrations of K2TeO3 (0.1 and 0.2 mM) were chosen to add in flasks separately for assessment of capability of strains to reduce chosen metalloid oxyanions. Controls were set (without selenite/tellurite and without bacterial cells) 4.2

Microbial Growth

CFUs in nutrient broth flasks without mentioned metalloid oxyanions and with 2 chosen concentrations of them were measured on nutrient agar plates on triplicate base.

50

4.3

Capability of Strains to Reduce Selenite and Tellurite

The aliquots of strains with final optical density of 0.01 in 100 ml of nutrient broth were added to each flask. Two types of control negative flasks were prepared as well (one without oxyanion and one without bacterial strain). Then flasks were incubated on shaker (250 rpm) at 27°C. 4.3.1

Determination of SeO3 2- Amount

The test was performed by the method described in Lampis et al. (2014). In each of 50 ml glass bottles 10 ml of 0.1 M HCl, 0.5 ml of 0.1 M EDTA, 0.5 ml of 0.1 M NaF, and 0.5 ml of 0.1 M of disodium oxalate were added followed by putting 50 μl and 200 μl of sample taken from flasks containing respectively 2 mM and 0.5 mM of selenite to make the total amount of 100 nmol of selenite in each glass bottle. The next step was adding 2.5 ml of 0.1% 2,3-diaminonaphthalene solution (sensitive to light) dissolved in 0.1 M HCl and incubating bottles at 40°C for 40 min and then cooling to room temperature. Final step was extraction of selenium-2,3diaminonaphthalene complex with 6 ml of cyclohexane by shaking the mixture vigorously for 1 min and checking the absorbance of the organic phase at 377 nm by spectrophotometer (UNICAM, Helios β). Calibration curves were made by using 0, 50, 100, 150 and 200 nmol of selenite in nutrient broth. All standard deviations were based on triplicates. 4.3.2

Determination of TeO3 2- Amount

100 µl of aliquot was mixed with 300 µl of 0.5 M Tris.HCL (pH 7) and 100 µl of Diethyldithiocarbamate reagent (DDTC) followed by reading the absorbance of the yellow mixture at 340 nm wavelength (Turner et al. 1992). All standard deviations were based on triplicates.

51

4.4 4.4.1 4.4.1.1

Exploration of the Mechanism Finding the Bacterial Compartment in which Reduction Occurs Subcellular Fractions

For this 3 different subcellular zones were analyzed: cytoplasm, periplasm, membrane. Moreover, supernatant was evaluated for the presence of extracellular reduction activity as well. 48 hours and 24 hours cultures of both strains in nutrient broth for stationary phase and exponential phase, respectively, were centrifuged separately at 10000×g for 10 min at 4°C to provide pellet containing bacterial cells followed by washing twice with 0.9% NaCl solution. The pellet was subjected to form spheroplast in which outer membrane is disrupted by lysozyme and EDTA, periplasmic fraction is solubilized, while the cytoplasmic membrane remains intact according to the method described by Osborn and Munson (1974). Centrifugation at 25000×g for 20 min makes spheroplasts collected as pellet. Then, they were suspended in 10 ml of 50 mM NaCl plus one tablet of complete protease inhibitor. The periplasmic fraction containing protein was collected from supernatant after filtration (0.2 µm Millipore). The suspension of spheroplasts in NaCl was sonicated for 7 cycles (40 seconds sonication/ 40 seconds rest in ice). Then they were centrifuged at 200000×g for 75 min. The membrane fraction was the brownish pellet which was suspended in 10 ml of 50 mM PBS (pH 7.4) containing 0.5% Triton X-100. The cytoplasmic fractions were recovered by filtration (0.2 µm Millipore) from supernatant. 4.4.1.2

Extracellular Fractions

48-hour culture of strains in nutrient broth was centrifuged at 5000 × g for 10 min at 4°C, then supernatant was filtered through 0.2 µm Millipore filter and evaluated. 4.4.2

Selenite and Tellurite Reduction Activity Test

The following combination was added to each well of 96-well microtiter plate: 50 μL of intracellular or extracellular fractions consisting protein (2 mg ml‾¹, measured by Bradford (1976) method), with sodium selenite solution (final concentration 5.0

52

mM) or potassium tellurite solution (final concentration 0.5 mM) and nicotinamide adenine dinucleotide (phosphate) (NADH/NADPH) (final concentration 2.0 mM) buffered with 100 μL of Mcllvaine buffer at different pH values (6.0, 6.2, 6.4, 6.6, 6.8 and 7.0). Finally, water added to final volume of 200 μL. Three negative controls were set up: without fraction, oxyanion and electron donor. The plate was then incubated at room temperature for 24 hours (Lampis et al. 2014). The aim was to see the change of color in medium to red-orange for selenite and gray-black for tellurite, showing the selenium and tellurium particles respectively produced by reduction. 4.4.3

Reduction in the Presence of BSO

25 ml of nutrient broth with final concentration of 0.5 mM of selenite for B. fungorum DBT1 and 1 mM for B. fungorum 95 and 3 different concentrations of Sn-butyl homocysteine sulfoximine (BSO) (0.5, 1.0, and 3.0 mM) were utilized. For tellurite, the initial used concentration was 0.1 mM tellurite for both strains. The remaining oxyanions in the medium after 0, 3, 6, 24, 30, 48, 72 hours of incubation were measured by spectrophotometer at 377 nm. Controls were set (without selenite, without bacterial strains and without BSO).

5 5.1

Assessment of Elemental Selenium and Nanoparticles (Se, Te) Measurement of Produced Elemental Selenium Amount

The method of Biswas and coworkers (2011) was used with slight modifications. The calibration curve was drawn by measuring absorbance (at 490 nm) of the red color resulting from elemental selenium produced by reducing selenite (1 to 10 μmol) by using 25 μmol of HN2 OH·HCl. Samples were withdrawn from flasks after 0, 6, 24, 48 and 72 hours of incubation. Then 10 ml of each sample was centrifuged (5000×g for 10 minutes). The aim of using centrifuge was to separate bacteria and selenite residues from red insoluble selenium collected as a pellet. The following step was washing the pellet twice with 2 ml of 1 M NaCl to remove selenite residue. Then selenium in pellet dissolved in 2

53

ml of 1 M Na2S then being centrifuged to separate cells and finally the absorption of the reddish solution was measured with spectrophotometer at 490 nm. All standard deviations were based on triplicate samples. 5.2

Separation of Biogenic Se and Te Nanoparticles (SeNPs and TeNPs)

Each strain was inoculated in 800 ml of nutrient broth supplied with 2 mM Na2SeO3 for obtaining SeNPs and 0.2 mM K2TeO3 for obtaining TeNPs. Then these flasks incubated aerobically at 27◦C for 24, 48 and 72 hours separately to compare the obtained biogenic nanoparticles during each time period. The next step is recovering cells with produced nanoparticles by centrifuging at 10000×g for 10 min followed by washing with 0.9% NaCl. The pellet were then re-suspended in 50 mM Tris/HCl buffer (pH8.2) and bacterial cells were lysed by ultra-sonication at 100W for 5 min. Then they were centrifuged for 30 minutes at 10000×g. Finally, TeNPs and SeNPs in supernatant were collected as pellet by centrifuging for 30 minutes at 40000×g and re-suspended in deionized water (Zonaro et al. 2015). 5.3 5.3.1

Morphology, Localization and Characterization of Nanoparticles Scanning Electron Microscopy (SEM)

Bacterial samples with nanoparticles of 24, 48 and 72 hours were fixed with 2.5% glutaraldehyde in PBS (pH 7.4), stayed overnight at 4ºC, followed by dehydration with increasing concentrations of ethanol (30%, 50%, 70%, 90% and 100%). Then they were put on metallic specimens stubs and observed with XL30 ESEM (FEIPhilips) equipped with an EDAX micro-analytical system. 5.3.2

Transmission Electron Microscopy (TEM)

Bacterial samples with nanoparticles extracted and then fixed with 2% glutaraldehyde in PBS (pH 7.4) for 2 h. Following step was post-fixation in 1% osmium tetroxide in aqueous solution for 2 h and dehydration in increasing concentrations of acetone. Afterwards, samples were put in a multi-well grid and observed by EM10 electron microscope (Zeiss, Oberkocken, Germany).

54

5.3.3

UV–visible Spectral Analysis of Nanoparticles

Absorbance was measured using Cary60 spectrophotometer at wavelengths between 200 to 800 nm. The NPs dispensed in deionized water were assessed. 5.3.4

Dynamic Light Scattering (DLS) and Zeta Potential Analysis of Nanoparticles

These analyses were performed by using a Zen 3600 Zetasizer Nano ZS from Malvern Instruments (Worcestershire, UK) equipped with a 633 nm Helium–Neon laser light source (4.0 mW), detecting scattering information at a fixed angle of 173°. 300 µL from nanoparticles suspension in water, were transferred to a quartz cuvette (10 mm path length) for measuring Zeta potential and 500 µL from nanoparticle samples were put in disposable cuvette for measuring DLS and data recorded at 25°C with equilibrium time of 30 seconds. All the values were obtained using the Malvern software.

55

RESULTS AND DISCUSSION The studied B. fungorum strains were isolated through enrichment culture method in the presence of DBT as a sole carbon source. It is proved that bacterial strains occurring in contaminated matrices can be involved in biotransformation of both organic and metallic contaminations in such environments (Anderson et al. 1998; Lovley and Anderson, 2000). Moreover, reduction of selenite during bioremediation approach by a Burkholderia strain was previously reported (Lusa et al. 2015). Therefore, here, we evaluated the tolerance of strains DBT1 and 95 of Burkholderia against high concentrations of hydrocarbons and their ability to transform high concentration of DBT and reduce selenite and tellurite in order to use them in bioremediation of contaminated sites by mentioned hydrocarbons and metalloids. Also the possibility of using them as a source of biogenic nanoparticles was assessed. Besides, the evaluation of bacterial mechanism of reduction of metalloid oxyanions by strains with hydrocarbon degradation capacity increases the knowledge in this field. Metalloid reduction profile together with tolerance evaluation of Burkholderia strains are not available in the literature and this study is the first contribution on the topic.

1

Transformation of DBT

In order to evaluate and compare the ability and behavior of two strains to grow and transform DBT (500 mg l-1) as a sole source of carbon and energy, the planktonic cells were grown in DM (minimal medium) in the presence of DBT as the only source of carbon. Growth curves had the highest cell density after 24 hours of incubation by increasing about one logarithmic order of the cell numbers for both strains (Figure 8). However, there was no growth in DM medium without DBT as carbon source. Within 72 hours of incubation, strain DBT1 transformed 89.85 ± 4.3% of the DBT in which 70.1 ± 10.0 was during the exponential phase of growth (Figure 8A). The other strain, however, transformed total amount of 40.25 ± 3.8%

56

of 500 mg l-1 DBT mainly during the stationary growth phase and beginning of death phase (only 5.04 ± 2.4% was in exponential phase). (Figure 8B). This means that transformation of DBT is correlated to the growth kinetics. For strain DBT1 the best performance for DBT transformation was when bacterial cells were in their exponential growth phase, while for the other strain the main transformation happened when bacterial cells were in stationary and death phase. As previously described, DBT is considered essential for formation of amino acids (cysteine, cystine, methionine), some vitamins and other vital compounds for growth of microorganisms (De Araújo et al. 2012). This could be the reason of the maximum transformation of DBT along with growth of strain DBT. In other words, strain DBT1 needs to consume 350 mg l-1 of DBT to achieve maximum growth. The maximum consumption of DBT during exponential phase was observed before (Buzanello et al. 2014). However, strain 95 used only 5% of DBT during exponential phase and 35% in stationary and death phase. Similarly, Rhizobium meliloti Orange 1 transformed the main part of DBT (around 60%) during stationary phase (Frassinetti et al. 1998). It was proved that Rhizobium meliloti Orange 1 has two pathways for metabolism of DBT, the first is by producing HFBT as the end product, and the other leads to DBT-5-oxide production. In latter, bacterial cells do not show increase in their biomass (occurs in stationary phase) (Frassinetti et al. 1998). Probably strain 95 has similar mechanisms.

57

Figure 8: Degradation of 500 mg l-1 of DBT and growth of bacteria. A is B. fungorum DBT1, B is B. fungorum 95. Three factors were measured together. Degradation of DBT (triangle), Growth using DBT as carbon source in DM medium (black circle), DBT concentration in control DM medium without bacteria (dotted line with star), and growth in control DM medium without DBT (dashed line). Results are expressed as the mean with standard deviation. Different letters in degradation curves are statistically different (P < 0.05, Tukey’s test).

Figure 9 shows the appearance of an orange color in inoculated DM medium by 72 hours exposure to 500 mg l-1 DBT due to the production of DBT degradation metabolites. Non-inoculated control resulted in DBT abatement of only 7.86 ± 4.1% by 72 h and no change of color in medium. Therefore, both strains significantly increased (P < 0.01) the removal of DBT in comparison to abiotic culture. Moreover,

58

strain DBT1 notably enhanced (P < 0.01) the abatement of DBT compared to strain 95 (Figure 8). As it was mentioned in introduction, B. fungorum DBT1 transforms DBT through Kodama pathway to HFBT (Di Gregorio et al. 2004). The assumption that strain 95 is capable of transforming DBT was initially based on the color change in DM medium from colorless to orange-red, indicating the biotic release of colored intermediates in the presence of DBT (Figure 9). Moreover, 95 in DM culture supplemented by DBT (as the sole carbon and energy source) showed growth parallel to decrease of DBT concentration levels in the growth medium (Figure 8). Therefore, appearance of the same orange color in the culture of strain 95 in DM medium supplied with DBT and the presence of dbt genes in this strain (Andreolli, 2010) strongly suggest Kodama pathway is the mechanism adapted by B. fungorum 95 for DBT degradation as well. In fact, both Burkholderia fungorum DBT1 and 95 have dbt genes in a similar arrangement, which makes them unique from other known organisms in PAH degradation with nah or phn-like class of genes (Andreolli, 2010). However, strain 95 is less effective compared to strain DBT1 to degrade DBT. There are some other bacterial strains capable of degrading DBT through Kodama pathway. Examples are: Rhizobium meliloti Orange 1 isolated from sediments of an oil drainage able to reduce 30% of initial 100 mg l-1 of DBT by 48 hours (Frassinetti et al. 1998). A natural mixed culture (called ERI-11), isolated from oil contaminated soil, degraded DBT through Kodama pathway as well. One of the bacterial strains in this mixture (called A11) degraded 90 % of 270 µM DBT (Khedkar and Shanker, 2014). Nine isolates from crude oil reservoirs degrade DBT through Kodama pathway as well. Pseudomonas stutzeri A76, Shingomonas sp. A54 and X. polyaromaticivorans 127W were the most efficient ones, able to degrade 50 mg l-1 DBT by 3 days (Kitauchi et al. 2005). Therefore, our results showed that both strains (especially strain DBT1) have much better degradation properties compared to the other strains available in the literature. The reason can be the presence of two set of genes (dbt) encoding for the initial steps of the oxidative Kodama pathway in

59

our strains. The genes involved in DBT transformation through Kodama pathway were cloned from Pseudomonas sp. for the first time (Denome et al. 1993). They were clustered into one operon under the control of a single promoter. They belong to the nah-like class of genes that are responsible for oxidation of naphthalene to catechol. Nah-like class of genes encode enzymes involved in transformation of PAHs with low molecular fraction. Phn-like class are phenanthrene catabolic genes which were also found in Burkholderia sp. strain RP007 (Laurie and Lloyd, 1999). The fact that our studied Burkholderia strains possess dbt instead of nah or phn-like class of genes could make them more efficient than other bacterial strains. The dbt genes are organized in two operons instead of one. So, the potent activation of the two operons by the studied strains may be responsible for the higher efficiency of DBT transformation by them. This was proved for strain DBT1 (Di Gregorio et al. 2004), and hypothesized for strain 95 (Andreolli, 2010). Moreover, since strain 95 is an endophyte strain, it would probably act better in coexistence with a plant and that could be the reason which makes it less effective than strain DBT1, isolated from soil. Both strains can be considered as an effective catalyst of DBT transformation to HFBT as end product. Bressler and Fedorak (2001) previously used mixed cultures in order to mineralize HFBT to CO2. Therefore, Burkholderia fungorum DBT1 with ability to transform DBT rapidly to HFBT molecule, with ability to be mineralized easily, has the potential to be exploited in DBT biodegradation. Besides, Burkholderia fungorum 95 also showed promising efficiency to transform DBT. By using strain 95 synergically with plant (phytoremediation), we may get more potent biodegradation results from it. Because, evidences suggest that organic contaminants are often removed quicker and more efficient from planted soils than from soils without vegetation (Andreolli, 2010). Another suggestion is since both strains transform DBT during different growth phases, it would be promising to

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project a study in which by mixing both strains, we may obtain synergic effects of both strains which last longer (during both exponential and stationary phase).

Figure 9: Change of color in bottles containing DM medium with 500 mg l-1 of DBT for 95 (B), DBT1 (C) after 72 hours. A is control with bacteria but no DBT (both strains showed same results for control).

2

Biofilm Formation

As it was mentioned in previous section, isolation of indigenous bacterial strains able to degrade condensed thiophenes (the organic compounds which tend to bioaccumulate throughout the food chains) and use them as sole source of carbon and energy, may result in applications in bioremediation protocols. Previously, Di Gregorio et al. (2004) showed that strain DBT1 can degrade oxidatively DBT, through the Kodama pathway, within three days of incubation and remove efficiently the initial 100 mg l-1 of DBT from the culture. Moreover, our results showed strains DBT1 and 95 are capable of transforming 450 and 200 mg l-1 DBT

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respectively, which are notable amounts compared to literature. Moreover, Andreolli et al. (2011) showed flexible metabolism of PAHs by strain DBT1. The analyzed PAHs were naphthalene, phenanthrene and fluorene in concentration of 100 mg l-1. Strain DBT1 showed an encouraging traits for the possible application in the clean-up approach of contaminated sites by PAHs. However, these authors exposed only strain DBT1 in planktonic form to individual PAHs. In order to be able to use Burkholderia fungorum strains DBT1 and 95 in DBT bioremediation protocols, apart from high degradation capability, they should have high tolerance not only to high concentrations of DBT but also to a mixture of PAHs, as in most contaminated sites they are found together. In fact, one of the main aspects that should be taken into account to allow bioremediation to succeed is the tolerance by the selected microbes to the mixture of toxic compounds they are expected to degrade at the concentrations present at polluted sites (Seo et al. 2009; Mandal et al. 2012; Zingaro et al. 2013). Recently, use of biofilms of bacteria in biotechnology field have attracted attention. Opposed to the planktonic mode of bacterial state, biofilms show better mechanical stability with better application in bioreactors. One of the mechanisms that makes biofilms more resistant to toxic elements, is the limited penetration of toxicants through biofilm. There are also increasing opportunities of horizontal gene transfer and intercellular social interaction in biofilm that leads to rapid use of xenobiotic substrate (Demeter et al. 2015). Therefore, we tried to evaluate tolerance of our strains in both planktonic and biofilm states against high concentrations of DBT and a mixture of PAHs, to see if they are tolerant and if biofilm state increases the resistance. Our results showed that both strains DBT1 and 95 are able to establish good biofilms. Initial biofilms formation (in 96-well microtiter plate containing YMB) was without presence of any aromatic hydrocarbon source. Growth curve showed that both strains after 24 hours reached stationary phase with similar cell numbers.

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However, after 72 hours strain 95 showed 1 logarithmic point higher cell numbers than B. fungorum DBT1 (Figure 10A). Microscopic observations of microbial biofilms have shown that these sessile bacterial biofilms have a structure with complicated architecture. Some bacterial species such as Pseudomonas aeruginosa develop thick layers of mushroom- like structures divided by water-filled spaces. This structures consist of bacterial cells embedded in EPS matrix or glycocalyx (mushroom-like biofilm). Other forms of biofilms are microcolonies composed of cell groups divided by water channels (structured biofilm) and continuous layer on the surface (uniform biofilm) (Davies et al. 1998). Figure 10B demonstrates that strain Burkholderia DBT1 produces a uniform biofilm, while the Burkholderia 95 has a structured biofilm. It has been proved that three dimensional architecture of biofilm (structured and mushroom-like biofilms) is due to involvement of EPS. Exopolysaccharide has a role in stabilizing the 3D structure of biofilm with mechanism of providing physical barrier and minimizing the intercellular interaction (Watnick and Kolter, 1999). It is also suggested that EPS has a role in cell aggregation (Barrow et al. 1984; Li et al. 2006). Therefore, structured biofilm of Burkholderia 95 can be due to EPS production. Similarly, strain of Zymomonas mobilis is capable of producing biofilm composed of microcolonies embedded in EPS which is divided by open water channels (Li et al. 2006).

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Figure 10: Panel A: Growth curves of biofilm formation. B. fungorum DBT1 is circle and B. fungorum 95 is square. Panel B: CLSM analysis for biofilm of strains after 72 hours. A is 95 and B is DBT1. The strains were grown on the polystyrene pegs in YMA without hydrocarbons.

3

Evaluating Tolerance

The 72-hour biofilm exposed to PAH and DBT to obtain the tolerance of biofilms for both strains, while planktonic cells initially grew with PAH and DBT in YMB in 96-well microtiter plate. The reason that a 72-hours-old biofilm was exposed to

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toxicants was to decrease the cell shock from exposure of DBT and PAHs by exposure of a mature biofilm with better structure and cell numbers (after 72 h). 3.1

Tolerance of Biofilm and Planktonic cells in the Presence of DBT

Tolerance of biofilms to increasing concentrations of DBT were similar for both strains. Both found to be tolerant to 2048 mg l-1, highest used concentration. Both strains in both state of planktonic and biofilm had constant levels of CFUs with increasing concentrations of DBT (Figure 11). Although, statistical analyses showed some difference among the growth pattern, both states of cultures in both strains evidenced steady levels of growth even with higher concentrations of DBT. Figure 12 shows that Burkholderia 95 exposed to low concentration of DBT (8 mg l-1), forms more dense biofilm respect to higher concentration. In the presence of highest used concentration of DBT, the density of biofilm decreases to a microcolony structure. However, this effect for strain DBT1 is less observed and this strain showed the most uniformity at 128 mg l-1 DBT. The overall effect of DBT on the structure of biofilm is formation of microclustere and aggregation. There is no study dealing with evaluating the tolerance of biofilm against condensed thiophene. However, Demeter et al. (2015) showed that oil sand waste water microbial community revealed multispecies biofilm with microcolony structure with naphthenic acids. Bio-webs (also called bridge) previously was described in the presence of PAHs like naphthalene (Rodrigues, 2005). The reason is the cluster arrangement of biofilm decrease the external surface and results in less exposure to external stress (Flemming and Wingender 2010).

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Figure 11: Growth of planktonic cells (grew with DBT) and exposure of biofilm cells to DBT (after 72 h). Planktonic cells of strain 95 (black circle), planktonic cells of strain DBT1 (white square), biofilm of strain 95 (white circle), biofilm of DBT1 (black square). Results are expressed as the mean of 3 measurements and standard deviation. Different letters and symbols are statistically different (P < 0.05, Tukey’s test).

Figure 12: CLSM analysis for biofilm of strains after 72 h of exposure to DBT. A is strain 95 and B is strain DBT1. A1 and B1 are exposed to 8 mg l-1 of DBT, A2 and B2 are exposed to 128 mg l-1 of DBT and A3 and B3 are exposed to 2048 mg l-1 of DBT. * This picture is taken to show only the structural change of biofilm in the presence of DBT. The aim is not to measure the growth and cell density. In order to measure the cell density we need at least 20 different measurement through taking pictures from different zones of biofilm pegs.

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3.2

Tolerance of Biofilm and Planktonic cells in the Presence of PAHs

The only study that evaluated tolerance of bacterial strains (only in planktonic form) to high concentrations of PAH is the study of Zafra and colleagues (2014). They measured tolerance of strains Pseudomonas aeruginosa B7, Klebsiella sp. B10 and Stenotrophomonas maltophilia B14, as planktonic form, to up to 6000 mg l-1 concentrations of PAHs (phenanthrene, pyrene and benzo[a]pyrene). The aim was to find potential strains for PAH biodegradation. However, these authors did not measure tolerance against naphthalene which is another important toxic compound. Six different concentration ratios of PAHs mix were evaluated to sustain growth of the two strains (Figure 13). All mixtures supported growth for both organisms under both planktonic and biofilm incubation conditions until the concentration of 500, 200, 100 mg l-1 of naphthalene, phenanthrene and pyrene, respectively (P < 0.05). However, B. fungorum 95 planktonic cells found the highest concentrations lethal whereas strain DBT1 could tolerate the highest concentrations (naphthalene 2000 mg l-1, phenanthrene 800 mg l-1 and pyrene 400 mg l-1) regardless of biofilm or planktonic growth, but these concentrations caused a reduction of two logarithmic orders for planktonic (P < 0.05) and biofilm (P < 0.01) of strain DBT1. For strain 95 there was no inhibition in biofilm state (P < 0.05) while no growth was detected for planktonic state (P < 0.01) in the presence of highest used concentrations. The reason that biofilm of strain 95 was effective to increase the resistance toward PAHs could be its ability to form structural biofilm which is more effective to reduce the explosion surface with organic pollutions (Watnick and Kolter, 1999). Mangwani et al. (2013) showed that biofilm-mediated bioremediation using P. mendocina NR802 significantly enhanced PAHs degradation by a marine strain. In future, biodegradation capacity of biofilms can be measured and compared with planktonic forms. In contrary of exposure to DBT, with increasing the concentration of PAH, the density of biofilm exposed to PAHs for both strains remained high (Figure 14). The

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effect of exposure to PAH on biofilm of strains is forming a dispersed structure. So, we can say that, the exposure of biofilm to PAHs mixture of naphthalene 2000 mg l-1, phenanthrene 800 mg l-1 and pyrene 400 mg l-1, cause dispersion for B. fungorum 95 and B. fungorum DBT1. Comparisons between control growth and growth in the presence of different concentrations of DBT and PAHs in both biofilm and planktonic modes show that these pollutants are well tolerated for our strains.

Figure 13: Growth of planktonic cells (grew with PAH) and exposure of biofilm cells to PAH (after 72 h). Planktonic cells of strain 95 (black circle), planktonic cells of strain DBT1 (white square), biofilm of strain 95 (white circle), biofilm of strain DBT1 (black square). Results are expressed as the mean of 3 measurements and standard deviation. Different letters and symbols are statistically different (P < 0.05, Tukey’s test).

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Figure 14: CLSM analysis for biofilm of strains after 72 h of exposure to PAH. A is strain 95 and B is strain DBT1. The structure didn’t change with increasing concentration of PAH.

4 4.1

Study of Burkholderia Strains in the Presence of Metalloids Minimum Inhibitory Concentration

The capability of converting sodium selenite to elemental selenium and potassium tellurite to elemental tellurium by considered bacterial strains was initially assessed on the basis of the red and black (respectively) colonies formation on nutrient agar plates containing metalloid oxyanions (Figure 15). Afterwards, minimum inhibitory concentration (MIC) assay showed that both strains are tolerant to selenite up to the dose of 5 mM and to tellurite up to the dose of 0.2 mM, while they do not grow at dose of 7.5 mM SeO32- and 0.5 mM TeO32- (Figure 16). Therefore, both concentrations of 0.5 and 2.0 mM for selenite and 0.1 and 0.2 mM for tellurite were chosen as initial reference amounts of metalloid in the culture flasks throughout the reduction tests. In a bioremediation perspective, the MIC of these B. fungorum strains against selenite and tellurite are relatively low. For example Lysinibacillus sp. strain ZYM-1, which is a potential candidate for heavy metal bioremediation, is

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resistant to 100 mM selenite, 10 mM selenate and 2 mM tellurite and able to make nanoparticles from them (Zhao et al. 2016). Most tellurite-resistant microorganisms can resist tellurite at concentrations between 0.01 to 150 mM. However, Thermoactinomyces sp. QS-2006 is proved to be quite tolerate to tellurite (up to 500 mM) (Amoozegar et al. 2012). Nevertheless, two initial concentrations of each oxyanion were chosen to study the reduction profile and mechanism of transformation in the mentioned strain in order to use them for the biogenic source of nanoparticles.

Figure 15: Growth of B. fungorum 95 (A) and B. fungorum DBT1 (B) on Nutrient Agar plates without salts (A, B) or with 2.0 mM sodium selenite (A1, B1) and with 0.2 mM potassium tellurite (A2, B2)

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Figure 16: Minimum inhibitory concentration for selenite (A) and tellurite (B). circle for B. fungorum 95 and squares for B. fungorum DBT1

4.2

Microbial Growth

The microbial growth without oxyanion salts and in the presence of two concentrations of selenite and tellurite were measured. Without oxyanion salts, both strains increased the number of CFUs by 2 logarithmic points within 24 hours, from 7.26 to 9.55 log10 CFUs ml ‾¹ for strain 95 and 7.27 to 9.43 log10 CFUs ml ‾¹ for strain DBT1. Strain DBT1 stayed in stationary phase 24 hours more than strain 95. Regarding microbial growth in the presence of selenite, adding selenite caused a decrease of 1 logarithmic point in growth after 24 hours for both strains compared to control (8.75 log10 CFUs ml ‾¹ for strain 95 in the presence of 0.5 mM selenite and 8.51 log10 CFUs ml ‾¹ for the other strain under the same conditions) (Figure 17 A1, Figure 17 B1). Increasing the selenite concentration in the culture medium to 2 mM, did not make a difference in the growth of both strains compared to 0.5 mM selenite. Dynamics of cell growth in the presence of selenite was similar to that measured in culture in absence of this oxyanion; however, lower values were reached in stationary phase and this phase lasted less. Therefore, it seems that

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selenite at the present concentrations does not have severe toxic effect on both strains since their growth curves follow the same pattern of the relative controls and selenite does not notably affect cell division process and is not consider as a strong stress factor until 2 mM concentration for both strains. Similarly, the growth of Bacillus sp. strain JS-2 was not affected by increasing concentrations of selenite (Dhanjal et al. 2011). As we see in Figure 17, for B. fungorum 95 in the presence of both used concentrations of tellurite, there was a decreasing pattern in cell growth. This means tellurite is toxic for B. fungorum 95 even at concentration of 0.1 mM. Generally, tellurite is more toxic than selenite even at low concentrations (Amoozegar et al. 2012). However, cell growth of strain DBT1 in the presence of both used concentrations of tellurite followed closely the control growth curve. The effect of tellurite here is similar to selenite in which the presence of metalloid determined the increasing of bacterial growth of one logarithmic point instead of two logarithmic points (in control) within 24 hours (here they increased from 6.5 to 7.5 log10 CFUs ml ‾¹). Therefore, tellurite at used concentrations is not severely toxic for strain

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DBT1.

Figure 17: Growth of B. fungorum 95 (A) and B. fungorum DBT1 (B) as control (dot line), in the presence selenite (A1 and B1) and tellurite (A2 and B2). Growth in the presence of 0.5 mM selenite (dash line), in the presence of 2 mM selenite (gray line). Growth in the presence of 0.1 mM tellurite (black line), in the presence of 0.2 mM tellurite (square).

4.3

Capability of Strains to Reduce Selenite and Tellurite

Selenite and tellurite reduction efficacies of Burkhoderia strains were evaluated in nutrient broth (liquid rich medium) at increasing metalloid concentrations: 0.5 mM and 2.0 mM for selenite; 0.1 mM and 0.2 mM for tellurite. In all cases the abiotic controls were negative and biotic flasks were positive for the reduction (Figure 18). With initial concentration of 0.5 mM selenite in the cultures of both strains, by increasing the cell numbers during the first 24 hours (exponential phase), about three fourths (75%) of selenite depletion occurred, after an initial delay of 6 h. Finally, selenite was reduced completely by 96 hours (by the end of death phase) (Figure 17,

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Figure 18). Nevertheless, with 2 mM selenite, strains showed different responses. B. fungorum 95 could reduce 1.2 mM selenite of which 28% occurring within 24 hours and a further 61% within 48 hours. The remaining 11% of 1.2 mM was transformed within 96 hours. Therefore, the progressive reduction occurred between 24 to 48 hours (end of stationary growth phase and beginning of death phase). On the other hand, B. fungorum DBT1 reduced only 0.5 mM of 2 mM selenite with 79% reduction occurring within 24 hours (exponential growth phase) and remaining amount by the end of 72 hours. The reduction in the presence of 2 mM selenite was slower and less efficient for both strains (Figure 18). Similarly, in the study of Li and colleagues (2014) on Rhodopseudomonas palustris, selenite reduction was correlated to the growth kinetics as well. Also, Kessi et al. (1999) observed SeO3 2bacterial reduction during the transition between the exponential and the stationary growth phases. We can say that reduction in our strains is probably controlled by growth-phase dependent regulatory molecules. Strain 95 in the presence of higher initial concentration of selenite, reduced more amount and major part of reduction took place during stationary phase rather than exponential phase while strain DBT1 always reduced same amount (0.5 mM) in exponential phase. The reason could be in strain DBT1 in the presence of higher concentration of selenite (substrate), a negative feedback control mechanism activates and shuts down the involved enzyme. Therefore, even with more selenite in the culture, the reduction amount is the same. Strain 95 in the presence of 2 mM selenite acted similarly to when it was exposed to 500 mg l-1 DBT, with main part of transformation occurring in stationary phase. Probably strain 95 has two mechanisms (one activating in exponential phase and the other activating in stationary phase) for the transformation of both hydrocarbons and metalloids. As we see in Figure 18, B. fungorum 95 in the presence of 0.1 mM tellurite could gradually transform tellurite to half of the initial amount by the end of 96 hours. However, with initial 0.2 mM tellurite, only less than 15% of initial tellurite was transformed by 24 hours. From 24 hours onward, the concentration of tellurite

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remained same, while the growth was decreasing (Figure 17). After 24 hours the medium color changed to gray-black, revealing the formation of elemental tellurium. The initial lag phase of growth was seen previously in the growth of Pseudomonas pseudoalcaligenes KF707 in the presence of 100µM of tellurite (Tremaroli et al. 2009). Likewise, Slobodkina and colleagues (2007) showed that Bacillus thermoamylovorans SKC1 has a lag phase of 2 days for growth, while 80 % of 0.5 mM initial tellurite was decreased by the end of two days. The mechanism in Bacillus thermoamylovorans SKC1 proved to be detoxification. Therefore, tellurite had a bacteriostatic effect on the growth of Bacillus thermoamylovorans SKC1 and B. fungorum 95, meaning tellurite reduction is not dependent on the bacterial growth and is reduced during the lag phase of growth. In fact, in Bacillus thermoamylovorans SKC1 the growth did not start until tellurite was reduced. Probably tellurite inhibits the synthesis of vital metabolites for growth in strain 95 or induce the production of toxic metabolites that inhibits the growth until the reduction of tellurite completes. It was also observed in E. coli that bacterial growth was inhibited by tellurite, due to activation of oxidative stress (Pérez et al. 2007; Wang et al. 2010). Moreover, superoxide dismutase during the reduction of tellurite produces superoxide radicals that inhibits the growth of bacterial cells (Perez et al. 2007). For B. fungorum DBT1, however, tellurite reduction and microbial growth showed opposite trends, with TeO3 2- decreasing along with cell growth until they reach the stationary phase (by the end of 72 h) (Figure 17, Figure 18). In the presence of 0.1 mM tellurite, half of the amount was reduced gradually by 72 hours (when the cells were in stationary phase) and the other half was reduced promptly between 72 hour and 96 hours (when the cells were in death phase). The transformation of tellurite after 72 hours to 96 hours also was seen in Roseococcus thiosulfatophilus tested by Yurkov and colleagues (1996) while for other 6 studied strains transformation to

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elemental tellurium occurred in 24 hours. However, in the presence of 0.2 mM tellurite only 15% of whole amount decreased by the end of 96 hours in strain DBT1. This means tellurite until the concentration of 0.2 mM is not severely toxic for B. fungorum DBT1 but still B. fungorum DBT1 is not able to transform it completely. On the other hand, 0.1 mM concentration is not only non-toxic but also transformable. Tellurite is more toxic that selenite for our strains, similarly to Bacillus thermoamylovorans SKC1 (Slobodkina et al. 2007). These two strains evidenced very different behaviors compared to each other towards selenite and tellurite, in terms of selenite and tellurite reduction efficiency and growth. In fact, selenite reduction was more efficient in strain 95, while strain DBT1 was more efficient in tellurite reduction. To clarify, in the presence of lower concentration of selenite (0.5 mM) both strain reduced selenite in a similar manner, reducing the whole amount by the end of 96 hours with 75% reduction in exponential growth phase. Their difference was in the presence of higher concentration of selenite (2 mM) in which strain 95 reduced 60 percent (mainly between 24 to 48 hours), while strain DBT1 reduced only 25% after 96 hours of incubation (mainly in exponential phase). The correlation between exponential growth and selenite reduction is consistent with a possible usage of detoxifying mechanisms by these two strains (Lortie et al. 1992). Although the final growth yield decreased by adding selenite to the growth culture (compared to control growth), the increasing of selenite concentrations in the cultures does not affect growth curve. As for tellurium, strain DBT1 reduced 0.1 mM (half amount before 72 h and half after 72 h), while the strain 95 reduced 0.05 mM of tellurite. For both strains selenite reduction was correlated and dependent on growth, starting at mid exponential phase (after 6 hours), while tellurite reduction started from the beginning of incubation period, independent on growth, for both strains: strain 95 did not grow in the presence of tellurite and half of tellurite reduction in strain DBT1 occurred after 72 h when cells were in death phase. Selenite was not severely toxic to neither strains, while, tellurite had bacteriostatic effect on strain 95. Increasing the concentration of both selenite and

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tellurite, had a negative effect on the reduction capability of both strain. By exposing the bacterial cells to increased level of selenite, the reduction capacity of strain 95 decreased 40%, while this drop in capacity was 75% for strain DBT1. However, increased level of tellurite reduced the capacity of strain 95 by 35% while this drop in capacity was 85% for strain DBT1. Likewise, 2mM selenite had an inhibitory effects on Bacillus sp. strain SF1 selenite reduction capacity (Kashiwa et al. 2000). This could be due to saturation of reductases. We suggest to study the synergistic effects of using both strains together in one culture for reduction of selenite and tellurite to benefit from the characteristics of both strains.

Figure 18: A and B: Selenite reduction in the presence of 0.5 mM selenite (A) and 2 mM selenite (B): Selenite reduction of B.95 (star) and B.DBT1 (white circle) (n=3). C: Tellurite reduction in the presence of 0.1 and 0.2 mM tellurite: Tellurite reduction of B.95 in the presence of 0.1 mM tellurite (black square) and 0.2 mM tellurite (white circle). Tellurite reduction of B.DBT1 in the presence

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of 0.1 mM tellurite (white square) and 0.2 mM tellurite (black circle) (n=3). Dash and dot lines are sterile medium plus oxyanions.

4.4 4.4.1

Exploration of the Mechanism Selenite and Tellurite Reduction Activity Test

One of the possible mechanisms of oxyanions reduction in bacteria is through enzymatic reactions. There are different enzymatic systems for the reduction of selenite and tellurite by bacterial cells. The involved enzymes are found in different parts of bacterial cells. For example in E. coli, it has been proved that enzymes involved in selenite reduction are in cytoplasm and periplasm (Debieux et al. 2011).On the other hand, membrane-bound nitrate reductases (NRs) A and Z showed tellurite reduction activity, which is expressed under anaerobic conditions (Avazeri et al. 1997). Moreover, nitrite reductase in Thauera selenatis (DeMollDecker and Macy, 1993) and in Rhodobacter sphaeroides (Sabaty et al. 2001) is able to reduce selenite and tellurite, respectively. Bacillus mycoides SeITE01 has both membrane and extracellular protein involved in reduction of selenite (Lampis et al. 2014). Moreover, Kim and colleagues (2013) showed Shewanella oneidensis MR-1 produces extracellular crystalline tellurium nanorods by tellurite reduction in the presence of iron. In order to localize the cellular compartment where selenite or tellurite reduction takes place, enzymatic in vitro tests were performed. B. fungorum DBT1 and B. fungorum 95 were fractionated in stationary phase to obtain three subcellularcompartments (cytoplasm, periplasm and membrane fractions). Also supernatant was analyzed. These tests showed that for both strains only the cytoplasmic fraction is effective in selenite and tellurite reduction with NADH acting as the most efficient electron donor, as we can see from a more marked change of color with NADH than NADPH, meaning the more reaction in the presence of NADH (Figure 19 Panel I). In case of tellurite, for both strains there was low activity also without electron

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donor. Therefore, selenite and tellurite reduction occurs mainly by cytoplasmic enzymatic reaction in both strains. Since reduction of selenite for both strains was dependent on growth unlike tellurite, selenite reduction activity in subcellular level was performed also on strains fractioned in exponential phase of growth. In these phase, strain DBT1 showed more activity in the presence of NADPH in cytoplasmic and periplasmic fractions, while strain 95 showed the same behavior as stationary phase (Figure 19 Panel II).

Figure 19: Panel I: Cytoplasmic fraction for both strains fractioned in stationary phase (B. fungorum 95 (A) and B. fungorum DBT1 (B)) react with selenite (right) and tellurite (left) and electron donor (NADH (A1 and B1) and NADPH ((A2 and B2)). Panel II) Cytoplasmic fraction (above) and periplasmic fraction (below) for strains DBT1 fractioned in exponential phase reacting with selenite and NADPH. In all pictures, 1, 2 and 3 are three replicates of subcellular fractions reacting with metalloid in the presence of electron donor, while 3 following wells are 3 controls: i-without metalloid, ii-without electron donor and iiiwithout subcellular fraction respectively. Only positive enzymatic activities are shown.

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4.4.2

Reduction in the Presence of BSO

Selenite reduction to elemental selenium in vivo has been demonstrated to be a quite dynamic process, involving many intermediates, such as dimethyl-selenide, dimethyldiselenide, dimethyl selenone and selenodiglutathione (Yu et al. 1997; Kessi and Hanselmann, 2004; Cui et al. 2008b). For instance, in Ralstonia metallidurans CH34 reduction occurs after several hours due to complex assimilatory and detoxification pathways. Finally, Se0 and alkyl selenide are produced in equal proportions (Sarret et al. 2005). One of the first reduction pathways that has been discovered for selenite is chemical reaction between selenite and thiol groups of proteins, so called, Painter-type reaction (Painter, 1941). Glutathione (GSH) has an essential role in neutralizing oxidative stress compounds and is the most abundant thiol in cytoplasm of some prokaryotic cells (Kessi and Hanselmann, 2004). Glutathione or glutathione reductase reduce selenite to selenodiglutathione (GS-Se-SG) and glutathionyl selenol as intermediates and finally to hydrogen selenide (H2Se) and elemental selenium. NADPH is the main electron donor involved in the process (Ganther, 1971; Kessi and Hanselmann, 2004). Since our results evidenced that selenite reduction takes place in the cytoplasm, glutathione may be involved in the reduction process. In order to prove this hypothesis, the microbial cells were cultured with an inhibitor of glutathione synthesis (BSO in 3 different concentrations) and compared with cultures without BSO. This is to evaluate if selenite reduction efficiency is correlated with glutathione synthesis or not. It has been proved that in some microbial isolates the presence of BSO can decrease the amount of selenite reduction and this is correlated directly to the initial concentration of BSO (Antonioli et al. 2007; Kessi, 2006). As we can see in Figure 20 and Table 1, BSO significantly decreased selenite reduction in both strains, especially in B. fungorum DBT1, as it can also be observed

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from less red color in bottles containing BSO with regard to the control bottle. For the strain 95 after 72 hours, 100% of selenite was reduced in control flask, while in the flask containing 3 mM BSO, only 79% was reduced. For the strain DBT1 the reduction in the presence of 3 mM BSO even was lowered to 60%. This correlation between inhibition and initial amount of BSO had already been reported by Antonioli and coworkers (2007). Therefore, glutathione-based selenite reduction is active in these two studied bacterial strains. For B. fungorum DBT1 it is more important because reduction was decreased significantly in the presence of BSO and also selenite reduction activity showed that NADPH is the main electron donor in exponential phase. NADPH is also the main electron donor in glutathionebased mechanism. In previous section we suggested that probably in strain DBT1 there is an enzymatic system which works on the basis of negative feedback control. Likewise, it was suggested that in Redox regulation by glutathione there is negative feedback inhibition involved (Berndt et al. 2014). The reason for better performance of B. fungorum DBT1 in exponential phase is that this strain may express the highest level of reductases and/or proceeds towards the saturation of glutathione synthesis during the exponential phase. Moreover, in the exponential phase this strain showed selenite reduction activity in both periplasmic and cytoplasmic fractions, while only the cytoplasmic fraction could actively reduce SeO3

2-

in the stationary phase.

Probably because of negative feedback control activation in exponential phase, the strain cannot be very active in stationary phase. Since for both strains NADH was the main electron donor in stationary phase, other additional mechanisms could be by NADH related reductases such as oxidoredoxin. Hunter (2014) found a flavin oxidoreductase in Rhizobium selenitireducens that could reduce selenite to elemental selenium using NADH not NADPH. This reductase has more tendency to accept electrons from NADH rather than from NADPH.

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Other involved enzymes in selenite reduction are thioredoxin reductase in Bacillus subtilis (Garbisu et al. 1999) or hydrogenase in Clostridium pasteurianum (Yanke et al. 1995), which can be important also in Burkholderia strains. Similarly, there are some biological molecules that can be involved in selenite reduction such as bacillithiols and cysteine (Lenz et al. 2011; Lampis et al. 2014). Turner and colleagues, in studies dealing with tellurite resistance in Escherichia coli, pointed out that glutathione and other reduced thiols (RSH) in the cytoplasm have role in tellurite reduction and production of tellurium as well (Turner et al. 1995; Turner et al. 1999). Baesman (2007) explained that tellurite reduction can be achieved by enzymes with broader substrate affinities, such as nitrite reductase or dimethyl sulfoxide reductase or by involvement of cytochromes. Moreover, tellurite, which is a strong oxidant, can act as electron acceptor from lactate oxidation. Also, Chasteen and colleagues (2009) provided information about intracellular enzymatic reduction of tellurite by nitrate reductase, dihydrolipoamide dehydrogenase (LpdA) (in E. coli (Castro et al. 2009)), and catalase or by other unspecific reductase enzymes (in E. coli (Pérez et al. 2007)). Tellurite can be an effective substrate for other redox enzymes including catalase, lipoamide dehydrogenase and squalene monooxygenase (Rigobello et al. 2011). Tellurite can also be reduced chemically by reduced thiols. Similarly, other low molecular weight compounds such as cysteine can reduce tellurite. Also tellurite interacts with the thiol-dependent enzymes (such as thioredoxin reductase (TrxR1 and TrxR2)) altering the balance between pyridine nucleotides and thiol redox state (Rigobello et al. 2011). Tellurite reduction was not suppressed by BSO in both DBT1 and 95 strains showing that glutathione is not involved in tellurite reduction (data not shown). Therefore, the possible mechanism can be reduction with reductases dependent on electron donors or thiols other than glutathione. Previously it was reported by Pérez and colleagues (2008) that aldehyde reductase, which is part of a glutathione-

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independent system, can protect E coli from tellurite toxicity. A similar glutathioneindependent system can be in charge in our strains. Strains showed some activity even without the used electron donors in reduction of tellurite which means also abiotic reaction (thiols for example) is involved, or tellurite reductases which also accept electron from electron donors other than what we used here, like lactate. Also quinon redox mediators such as lawsone and anthraquinone can be involved (Ramos-Ruiz et al. 2016). Both strains seem to have different mechanisms for selenite and tellurite. For both strains in the presence of selenite and tellurite, cytoplasmic enzymatic systems are probably the main mechanism involved. For strain DBT1 glutathione is more important than strain 95 for selenite reduction, while for tellurite reduction glutathione is not involved for neither strains. Strain DBT1 has more activity in exponential phase for reducing selenite due to activation of both cytoplasmic and periplasmic enzymes. Probably also the cell content of glutathione is more in exponential phase since BSO had the most effect on inhibition of reduction during exponential phase and NADPH acted as the best electron donor in this growth phase. For strain 95, enzymes which accept electron from NADH rather than NADPH, are more important for reduction of selenite and tellurite. Glutathione is important for selenite reduction but not tellurite reduction. For both strains selenite reduction occurs only in the presence of electron donors and tellurite is reduced better with NADH and NADPH. There are multi mechanism for reduction in both strains.

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Bacterial Cell B.95 (no BSO) B.95 (BSO 0.5 mM) B.95 (BSO 1 mM) B.95 (BSO 3 mM) B.DBT1 (no BSO) B.DBT1 (BSO 0.5 mM) B.DBT1 (BSO 1 mM) B.DBT1 (BSO 3 mM)

0h 1.00± 0.05 0.99± 0.03 0.99± 0 0.98± 0.01 0.50± 0 0.49± 0.01 0.49± 0 0.49± 0.01

3h 1.02± 0 0.99± 0.03 0.97± 0 0.99± 0 0.50± 0 0.49± 0.02 0.48± 0 0.49± 0

6h 0.96± 0.07 0.98± 0.01 0.99± 0 0.98± 0.01 0.48± 0.01 0.49± 0 0.48± 0.04 0.49± 0

24h 0.43± 0.01 0.39± 0.03 0.50± 0.01 0.57± 0.04 0.11± 0.02 0.23± 0 0.25± 0 0.22± 0

30h 0.13± 0.03 0.13± 0.01 0.33± 0.01 0.34± 0.02 0.06± 0.01 0.23± 0.03 0.16± 0.01 0.22± 0.03

48h 0.08± 0 0.06± 0.03 0.16± 0.01 0.22± 0.01 0.01± 0.01 0.20± 0 0.16± 0.01 0.21± 0.01

72h 0.06± 0.03 0.06± 0.03 0.19± 0.01 0.21± 0 0.01± 0 0.15± 0 0.16± 0.01 0.21± 0.02

Table 1: remaining concentration of selenite (in mM) in flasks with different concentrations of BSO in different time intervals. BSO 3 mM showed a significant difference with control after 24 hours for both strains (ANOVA; P< 0.05, n=3). Data is presented with standard deviation.

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Figure 20: Change of color in presence of selenite and BSO, this Picture is taken after 72 hours. Written concentrations on pictures belong to BSO. A shows strain 95 and B shows strain DBT1.

5 5.1

Assessment of Elemental Selenium and Nanoparticles (Se, Te) Measurement of Produced Elemental Selenium Amount

As shown in Figure 21, by reduction of selenite, Seº started to be formed and the color of medium started to turn to red in less than 24 hours. The control flasks (abiotic and without selenite) did not change color, indicating the formation of

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selenium by reduction of selenite by bacterial strains. As far as the dynamics of elemental selenium formation is concerned with respect to selenite reduction, a parallel trend was observed. To clarify, there was no delay between reduction of selenite and production of elemental selenium. The exception was the strain DBT1 in the presence of highest concentration of selenite in which there was a 24 hours delay

between

reduction

Rhodobacter sphaeroides

of

selenite

and

production

of

Seº.

In

(Van Fleet-Stalder et al. 2000) and Ralstonia

metallidurans CH34 (Sarret et al. 2005) a delay was observed which was due to the production of organic selenide (RSeR) intermediate before converting to elemental selenium. As it was explained in material and method part, spectroscopical method was used to detect elemental selenium in the cultures. The highest absorbance related to concentration of elemental selenium in the culture, was obtained from B. fungorum 95 culture grown with 2 mM selenite. 1 mM elemental selenium produced under these conditions by strain 95 by reducing 1.2 mM selenite with 85% yield of Seº production. B. fungorum DBT1 in the presence of 2 mM selenite had only 15% selenium production yield. However, both strains with 0.5 mM selenite had 65% selenium production yield. Therefore, in B. fungorum 95 by increasing the initial concentration of selenite in the flasks (from 0.5 mM to 2 mM), both amount of reduced selenite and produced selenium increased (from 0.5 mM to 1.2 mM for selenite and from 0.3 mM to 1 mM for selenium); while in B. fungorum DBT1 they are constant amounts (0.5 mM for selenite and 0.3 mM for selenium), regardless of initial selenite concentration. This can be another evidence for existence of negative feedback control mechanism in strain DBT1. Tam and colleagues (2010) found a similar behavior as B. fungorum DBT1 in Shewanella sp HN-41. This strain did not produced more selenium by adding more selenite in the medium. They used a range of initial concentrations of selenite from 0.01 to 1 mM. In this case, maximum selenite reduction and nanoparticle production were obtained with only 0.1 mM of selenite, due to the saturation of reductase enzyme.

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Furthermore, spectra of SeNPs produced after three different incubation times, 24, 48 and 72 hours, were analyzed. All spectra showed the max absorption at 270 nm (Figure 22). This means, as already reported (Debieux et al. 2011; Lenz et al. 2011) that some peptides and protein groups are likely to cover nanoparticles that have absorbance in this wavelength. In other words, this absorption can be due to the presence of amino acid groups around nanoparticles.

Figure 21: Selenite reduction and selenium production for strain 95 (A and B) and strain DBT1 (A and C), in the presence of 0.5 mM selenite (A) and 2 mM selenite (B and C). Selenite reduction of B.95 (star) and B.DBT1 (white circle) selenium formation of B.95 (white triangle) and B.DBT1 (black circle) (n=3).

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Figure 22: Spectrum of Se nanoparticles produced by B. fungorum 95 (left) and B. fungorum DBT1 (right) after 24 (dot line), 48 (line) and 72 (dash line) hours.

5.2

Morphology, Localization and characterization of Nanoparticles

SeNPs from cultures of both bacterial strains were spherical (Figure 23, Figure 24). Average hydrodynamic diameter of Se nanoparticles measured after 24, 48 and 72 hours was constant, 170 nm for B. fungorum 95 and 200 nm for B. fungorum DBT1. The zeta potential of SeNPs generated by both Burkholderia strains was zero (Figure 25). As for TeNPs, they were needle- like nanorodes (Figure 26, Figure 27). Average hydrodynamic diameter of Te nanoparticles measured after 24, 48 and 72 hours was constant, 120 nm for B. fungorum 95 and 170 nm for B. fungorum DBT1 (Figure 28). The zeta potential of TeNPs produced by Burkholderia strains was positive. Nanoparticles were seen only in the presence of metalloid salts. Bacillus sp. strain SF-l was able to produce SeNPs with size of 200 to 300 nm localized both inside and outside of cells (Kashiwa et al. 2000). Also Bacillus mycoides SeITE01 produced SeNPs with size of 50 to 100 nm both inside and outside cells with major portion outside of bacterial cells (Lampis et al. 2014).

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SeNPs were located inside Shewanella oneidensis MR-1, produced by intracellular reduction mechanism using periplasmic reductase (Li et al. 2014). Extracellular SeNPs and TeNPs were obtained using three strains by Bajaj and Winter (2014). Biological and non-biological selenium nanoparticles are usually negatively charged. However, Sonkusre and coworkers (2014) used Bacillus licheniformis JS2 to produce selenium nanoparticles and they obtained zero-charged nanoparticles. They proved that those nanoparticles were surrounded by some proteins and functional groups. It has been shown that functional groups surrounding nanoparticles, can change the zeta potential of nanoparticles according to their charge. Electrostatic, hydrophobic and chemical interactions exist between proteins and nanomaterials, especially electrostatic bound is proved to be the most important factor for selective absorption (lynch and Dawson, 2008). For instance, Rezwan and colleagues (2005) have proved that the negatively charged bovine serum albumin (BSA) and positively charged lysozyme can interact with metal oxide particles of alumina, silica, titania and zirconia and the existence of proteins changes the zeta potentials and the isoelectric points of the oxide particles. They also correlated the zeta potential of nanoparticle-protein complex with the nature of the main absorbed protein(s). Proteins associated to nanoparticles not only can alter the charge but also might be involved in selenite reduction (Lenz et al. 2011). Spectra and zeta potential of SeNPs produced by the Burkholderia strains of interest provide enough information to hypothesize the presence of some active proteins with functional groups or biomolecules surrounding the particles. Se0 nanoparticles mainly occurred extracellularly by 24-hours bacterial culture incubation (Figure 23). TEM showed both intracellular and extracellular SeNPs after 24 h of incubation (Figure 24). Selenite reduction activity assays showed cytoplasmic enzymatic activity, which suggests a possible intracellular formation of SeNPs. Necessarily, extracellular appearance of selenium nanoparticles is due to

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cell lysis or existence of some export mechanism(s). The transport should be further studied since nanoparticles are relatively large to pass through the membrane and vesicular excretion in bacteria is still controversial. However, there are some suggested secretion mechanisms for gram-negative bacteria which is called type 1– 6 secretion systems (TxSS). Another possibility to export both soluble and insoluble materials deals with outer membrane vesiculation (formation of spherical vesicles as a response to stresses). These vesicles, deriving from the periplasmic origin, can enclose different matrices within their lipid bilayer membrane and secrete them out (McBroom and Kuehn, 2007; Kulp and Kuehn, 2010). Debieux and colleagues (2011) proved that in Thauera selenatis a protein (SefA), which is exported directly from the cytoplasm, cause the secretion of selenium nanoparticles. However, the hypothesis of cell lysis is more possible since we can see some dead cells in TEM pictures after 24 hours of incubation (Figure 24). Granules of selenium accumulated inside cells can cause lower viability in cultures with selenite (Tomei et al. 1995). The lysis hypothesis is even more evident for tellurium nanoparticles. As we see from SEM pictures (Figure 26), after 24 and 48-hours of incubation bacterial cells are intact, while there is a tellurium peak in EDAX spectra, no nanoparticle can be seen, which provides evidence of intracellular formation of tellurium. However, after 72 hours, bacterial cells clearly lost their morphology and tellurium nanoparticles can be seen outside of cells. This fact indicates the probability of bursting cells in presence of toxic element and the consequent releasing cell materials including tellurium nanoparticles. 24 hours-TEM pictures confirmed the hypotheses provided by SEM results that i) tellurium is produced (section 4.4.1) and localized intracellularly (Figure 26), ii) few bacterial cells are starting to die even after 24 hours and tellurium nanoparticles realizing outside of cells (Figure 27). Enzymatic assays revealing involvement of cytoplasmic enzymes, is compatible with intracellular production of tellurium. Biogenic TeNPs generally are located intracellular with shape of needle crystals. However, there are also extracellular TeNPs. For example, marine tellurite resistant strains of Rhodotorula spp and Bacillaceae produce intracellular needle like TeNPs (Ollivier at al. 2008).

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Borghese et al. (2016) obtained needle like TeNPs produced in the cytoplasm of Rhodobacter capsulatus. Halococcus salifodinae BK3 also synthesizes intracellular crystals of TeNPs with needle shape (Srivastava et al. 2015b). The charge of produced Te nanoparticles is positive for both strains (Figure 28). Previously, antimicrobial effects for tellurium nanoparticles were described (Zonaro et al. 2015). Overall charge of bacterial cells is negative due to the presence of carboxylic groups. Therefore, the positive charge of tellurium nanoparticles increases their binding chance to bacterial cells due to electrostatic forces. Thus we can expect antimicrobial effects for produced tellurium nanoparticles (Jassim et al. 2015).

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Figure 23: SEM analysis for B. fungorum DBT1 (A-A3) and B. fungorum 95 (B-B3) showing spheric elemental SeNPs (shown by circles) and rod-shaped bacterial cells. A and B are controls without selenite. A1 and B1 are strains grown with 2 mM selenite after 24 hours, A2 and B2 are after 48 hours and A3 and B3 are after 72 hours. Graphs are EDAX spectra for both strains with selenium (I with Se peak shown by an arrow) and control (II without Se peak).

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Figure 24: TEM analysis for both strains with 2 mM selenite after 24 hours. B. fungorum DBT1 is A and C. fungorum 95 is B and D. Circle shows nanoparticles outside of bacterial cells and dead cell. Arrows show nanoparticles produced inside of cells.

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Figure 25: DLS analysis and Zeta potential: A is the size distribution of SeNPs from B. fungorum 95 and B is the size distribution of SeNPs from B. fungorum DBT1 cultures. C and D are Zeta potential of SeNPs generated by B. fungorum 95 (C) and B. fungorum DBT1 (D) cultures.

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B3

Figure 26: SEM analysis for B. fungorum DBT1 (A-A3) and B. fungorum 95 (B-B3). A and B are controls without tellurite. A1 and B1 are strains grown with 0.2 mM tellurite after 24 hours, A2 and B2 are after 48 hours and A3 and B3 are after 72 hours. Graphs are EDAX spectra for both strains with tellurium (I and II are with Te peak shown by arrows) and control (III without Te peak). I shows the EDAX spectra for area (figure A1-3, B1-3) and II shows the EDAX spectra for white spots shown with circles in figure A3 and B3.

95

Figure 27: TEM analysis for both strains with 0.2 mM tellurite after 24 hours. B. fungorum DBT1 is A and B. fungorum 95 is B. Arrows show needle like nanoparticles localized inside of cells and square shows dead cells along with nanoparticles released outside of bacterial cells by cell lysis.

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Figure 28: DLS analysis and Zeta potential: A is the size distribution of TeNPs from B. fungorum DBT1 and B is the size distribution of TeNPs from B. fungorum 95 cultures. C and D are Zeta potential of TeNPs generated by B. fungorum DBT1 (C) and B. fungorum 95 (D) cultures.

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CONCLUSION Several human activities such as combustion of fossil fuels, incineration of waste, industrial processes are causing the release of organic and non-organic pollution in the environment. Environmental matrices such as soils are co-contaminated with both types of pollutions. Therefore, finding a biotechnological alternative to chemical methods for their treatment, for example using bacterial strains tolerant to high concentrations of pollutants, is a priority. In the present study, 2 different DBT degrading isolates were studied for their aromatic hydrocarbon resistance in order to be used in bioremediation of soils contaminated by organic pollutants and metalloids. They were further studied for their reduction ability of selenite and tellurite as the sample oxyanions and production capability of nanoparticles. They are strain DBT1 and 95 of Burkhoderia genus, isolated respectively from oil drainage wastewater and plant tissue. They are promising candidates for bioremediation of soils polluted with PAHs and obtaining metalloid nanoparticles from oxyanions present in the same soil. Strain 95 can be also used in phytoremediation processes by using an integrated plant-microbe system for bioremediation. Both B. fungorum 95 and DBT1 formed good amount of biofilm. Moreover, they were tolerant to high concentrations of DBT and mixture of PAHs, grown either as a biofilm or as free-living cells, and biofilm formation increased the tolerance of B. fungorum 95 in the presence of PAHs mixture. Based on preliminary results of this study, these findings provide new perspectives on the effectiveness of using these strains as biofilm in bioremediation strategies of hydrocarbon contaminated sites. Furthermore, both B. fungorum 95 and B. fungorum DBT1 can efficiently reduce selenite and tellurite and produce SeNPs and TeNPs. With regards to selenite, in the presence of higher concentrations of selenite, B. fungorum 95 showed a better

98

performance than B. fungorum DBT1 either in selenite reduction or in production yield of nanoparticles. To clarify, in B. fungorum 95 culture, increasing the initial concentration of selenite from 0.5 mM to 2 mM resulted in the rise of SeO32reduction from 0.5 mM to 1.2 mM as well as the shift from the exponential phase to the stationary phase for main part of transformation. Formation of elemental selenium also was boosted from 0.3 mM to 1 mM with more initial selenite. On the contrary, in B. fungorum DBT1 both selenite reduction and Seº production remained constant (0.5 mM and 0.3 mM respectively) regardless of the initial selenite concentration and always occurring during the exponential phase. In B. fungorum DBT1 saturation of selenite reductive enzymes produced during exponential growth phase or depleting the glutathione content during the same phase and negative feedback control could be the reasons. With regards to tellurite, generally B. fungorum DBT1 showed a better performance than B. fungorum 95 in reduction. In the presence of lower concentration of tellurite (0.1 mM), B. fungorum 95 reduced half amount of initial tellurite to elemental tellurium, while B. fungorum DBT1 reduced the whole initial amount. Both strains with 0.2 mM of tellurite reduced only 15%. 0.2 mM is toxic for B. fungorum 95 but not severely toxic for B. fungorum DBT1. We also proved that the mechanism of selenite and tellurite reduction adopted by the two Burkholderia strains is mainly reduction due to the activity of cytoplasmic reductases which accept electron from NADH and NADPH. Slight tellurite reduction also occurred without used electron donors. Glutathione is also involved in selenite (but not tellurite) reduction since the inhibition of GSH synthesis by BSO had negative effects on SeO32- transformation to Se0. Moreover, glutathione appeared more crucial in B. fungorum DBT1 whereas the addition of BSO evidenced more consequences on selenite reduction than in B. fungorum 95. Moreover, other free thiols and thiol groups in proteins probably have a role in selenite and tellurite reduction as well as NADH dependent reductases.

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SeNPs synthesized by the strains 95 and DBT1 of Burkholderia fungorum have a diameter of 170 nm and 200 nm respectively and zero charge. A deeper research needs to be done in order to understand the cause of the neutral charge shown by SeNPs deriving from Burkholderia fungorum cultures. Furthermore, as SeNPs produced by both strains of B. fungorum mainly occurred outside the microbial cells, although the reactive protein fraction was cytoplasmic one, we hypothesized that they are initially formed enzymatically as a consequence of selenite reduction inside the cells and then likely transported outside by means of some secretion mechanisms or by cell lysis. TeNPs synthesized by the strains 95 and DBT1 of Burkholderia fungorum present a diameter of 120 nm and 170 nm respectively and positive charge. Finally, TeNPs produced by both strains of B. fungorum tested in this study were produced intracellularly and mainly localized intracellularly as well. However, after 72 hours of incubation we also saw extracellular tellurium, released due to cell burst caused by tellurite toxicity. All in all, we found out that B. fungorum DBT1 and B. fungorum 95 can be exploited for bioremediation of organic polluted sites containing also low to medium amount of selenite and tellurite and biogenesis of selenium and tellurium nanoparticles. The possible applications for these nanoparticles in medical (antibacterial and antifungal) or technological fields should be investigated especially for TeNPs with more toxicity to bacterial cells than SeNPs and with positive charge that have the better potential to be absorbed to pathogens as an antimicrobial agent. As future perspectives, their ability to reduce metalloids and degrade DBT and PAHs need to be done in situ or in bioreactor conditions. Also, additional experiments on nanoparticles external coating and transport mechanism are currently being performing.

Possible antimicrobial effects of NPs and

biodegradation capacity of biofilms are other aspects to consider.

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Insights into selenite reduction and biogenesis of elemental selenium nanoparticles by two environmental isolates of Burkholderia fungorum Nazanin Seyed Khoei¹, Silvia Lampis¹, Emanuele Zonaro¹, and Giovanni Vallini¹ 1

Department of Biotechnology, University of Verona, Verona, Italy

Background: Strains capable of transforming toxic selenium oxyanions to non-toxic elemental selenium provide not only the possibility of bioremediation of polluted sites but also green biogenesis of selenium nanoparticles with medical or technological applications. The present paper discusses the reduction of selenite by Burkholderia fungorum DBT1 (B.DBT1) and Burkholderia fungorum 95 (B.95) previously isolated from an oil refinery drainage and hybrid poplar grown in polycyclic aromatic hydrocarbon (PAH)-polluted soil, respectively. The aim of this study are to evaluate the growth and transformation ability and mechanism of these strains in presence of selenium oxyanion (selenite) and characterize obtained nanoparticles. Methods: Three parameters were measured during the assay of reduction efficacy: i) Bacterial growth by counting CFUs on Nutrient Agar Plates ii) Amount of SeO3 2- residue and iii) Formed elemental selenium [1]. Intercellular proteins were extracted and subjected to enzyme activity assay [1]. Extracted nanoparticles were characterized by using scanning electron microscopy and dynamic light scattering apparatus [2]. Results and Discussion: Selenite reduction capability was assessed by using two different initial concentrations of this selenium oxyanion: 0.5 and 2 mM. B.DBT1 was able to reduce 0.5 mM selenite, while B.95 reduced more than 1 mM selenite in 96 hours. Selenium nanoparticles produced as a result of selenite reduction were spherical and zero charged with size of 170 nm and 200 nm for B.95 and B.DBT1 cultures respectively. Nanoparticle production efficiency was always 0.3 mM for B.DBT1, while B.95 produced same amounts with 0.5 mM selenite but 1 mM nanoparticles with 2 mM selenite. A delay of more than 6 hours was observed between introducing selenite and appearance of selenium nanoparticles. Nanoparticles occurred extracellularly although their production was induced by cytoplasmic enzymatic activation with nicotinamide adenine dinucleotide (NADH) as an electron donor. There are some export mechanisms for the eventual release of Se0 nanoparticles outside of cells. Conclusion: The formation of elemental selenium nanoparticles under aerobic conditions in these strains might be ascribed to a reduction mechanism involving redox reaction. The obtained nanoparticles are in optimum size and can possibly have some applications which should be investigated. Keywords: bioremediation, nanoparticle biogenesis, polycyclic aromatic hydrocarbon, environmental isolates

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Does dibenzothiophene (DBT) and poly aromatic hydrocarbons effect biofilm versus planktonic growth of DBT degrading strains? Nazanin Seyed Khoei1, Silvia Lampis1, Giovanni Vallini1, Raymond J. Turner2.3. 1

Department of Biotechnology, University of Verona, Verona, Italy Department of Biological Sciences, University of Calgary, Calgary AB, Canada 3 Biofilm Research Group, University of Calgary, Calgary AB, Canada 2

Background: Tolerance of 2 dibenzothiophene (DBT) degrading strains, Burkholderia fungorum DBT1 (B.DBT1) isolated from an oil refinery wastewater drainage and Burkholderia fungorum 95 (B.95) from root of hybrid poplar grown in poly aromatic hydrocarbons (PAH) polluted soil in form of planktonic and biofilm is studied here. These strains have the capacity to degrade some poly aromatic hydrocarbons including DBT. The objective of study is to find differences in DBT and the mixture of PAH concentration levels that can be tolerated by these 2 soil isolates and note impacts of these pollutants on the biofilm structure. Methods: The growth as biofilm in 96 plate using Calgary Biofilm Device for both strains was established with the method described by Ceri et al., 1999 [1]. Growth curves were drawn based on colony counting units (CFU) counting on YMA (yeast, mannitol, and agar) plates. Biofilm photos were taken by Confocal Microscopy. Results and Discussion: The total count of B.95 is almost 20 fold of those for B.DBT1 after 72 hours. Growth as planktonic state have good survivability to DBT to concentrations up to 2048 ppm for both strains. Both strains as biofilm have good tolerance to DBT but generally B.95 has a slightly better tolerance to DBT. As for preexposed cells on petri dishes with DBT crystals on the lid, after 72 h biofilm of B.DBT1 was better than biofilm of B.95. Both strains as planktonic form tolerated the PAH mixture. B.DBT1 grew with 2000 ppm of naphthalene plus 800 ppm of phenanthrene and 400 ppm of pyrene. While the highest concentration that B.95 could tolerate was the mixture of 500 ppm of naphthalene plus 200 ppm of phenanthrene and 100 ppm of pyrene. Although in biofilm state both strains showed good tolerance, the number of CFUs and survivability were less than planktonic state. Both strains form biofilm in the form of web in presence of DBT. Conclusion: We conclude that both strains are tolerant to DBT and PAH mixture in high concentration in form of planktonic and biofilm. Keywords: dibenzothiophene, biofilm, planktonic, tolerance

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How DBT Degrading Strains Behave in Presence of Organic and Metalloid Pollutants Nazanin Seyed Khoei ¹, Silvia Lampis ¹, Raymond J. Turner ²´³, *Giovanni Vallini ¹ 1 Department of Biotechnology, University of Verona, Verona – Italy; 2 Department of Biological Sciences, University of Calgary, Calgary AB, Canada; 3 Biofilm Research Group, University of Calgary, Calgary AB, Canada; *Corresponding author: [email protected]it Abstract In an oil polluted site, several components of environmental concern can be found including Polycyclic Aromatic Hydrocarbons (PAH), Dibenzothiophene (DBT) and metals. Studies showed that the naturally occurring soil bacteria are able to transform them to safe end products (bioremediation). The aims of this research are 1- To find tolerance of Burkholderia fungorum DBT1 (B.DBT1) and Burkholderia fungorum 95 (B.95) to DBT and PAH mixture and in planktonic and biofilm forms 2- Evaluating the growth and transformation ability of these strains in presence of selenium and tellurium as sample inorganic pollutants. The growth as biofilm in 96 well plate was established for both strains using Calgary Biofilm Device (CBD) with the method described by Ceri et al., 1999 (1). Growth curves were drawn based on CFU counting on YMA plates. Biofilm photos were taken by Confocal Microscopy. Selenite and tellurite reduction efficacy assessment was performed based on protocols described by Kessi et al, 1999 (2) and Turner et al, 1992 (3) respectively. Both strains can tolerate high concentrations of DBT (2048 ppm) in both planktonic and biofilm forms while the mixture of PAH can only be tolerated by planktonic form for both strains. B.95 is able to tolerate the maximum used concentration of selenite (2 mM) and transform more than half amount to elemental selenium. On the other hand B.DBT1 can only convert 0.5 mM of selenite, while is able to transform completely initial 0.1 mM concentration of tellurite to elemental tellurium. References 1. Ceri H, Olson ME, Stremick C, Read RR, Morck D, Buret A. (1999). The Calgary Biofilm Device: new technology for rapid determination of antibioticsuscepti bilities of bacterial biofilms. J Clin Microbiol, 37(6):1771-6. 2. Kessi J, Ramuz M, Wehrli E, Spycher M, Bachofen R. (1999). Reduction of selenite and detoxification of elemental selenium by the phototrophic bacterium Rhodospirillum rubrum. Appl Environ Microbiol, 65(11):4734–4740. 3. Turner RJ, Weiner JH, Taylor DE (1992). Use of diethyldithiocarbamate for quantitative determination of tellurite uptake by bacteria. Anal Biochem, 204(2):292-5. Keyword Bioremediation, Dibenzothiophene, Polycyclic Aromatic Hydrocarbons, metalloid, biofilm

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