AcknowledgementFirst of all I would like to express my sincere gratitude and

thanks to ALLAH SWT for his guidance through this work and forall the blessings He has bestowed upon me.

I am thankful to the Academy of Scientific Research andTechnology that give me the scholarship for doing this work and giveme the financial support for doing this practical work.

My deepest gratitude and thanks go to my supervisor, Prof.Dr. Tarek A. A. Moussa, Botany Department, Faculty of Science,Cairo University for his tireless guidance and enthusiasm during thecourse of this work. His excellence in academic standard, dedicationtoward research and teaching will always be a source of inspirationand encouragement to me. His support, encouragement and hisnumerous insightful suggestions made this experience an enjoyableone.

I would like to wholeheartedly acknowledge and express mydeepest gratitude and thanks to my supervisor Assoc. Prof. Dr.Mervat El-Sayed, Biochemistry section, Chemistry Department,Faculty of Science, Cairo University for her guidance, enthusiasmand words of encouragement during this research work.

I would like to thank Prof. Dr. Ahmed Al- Sabbagh, the headof Egyptian Petroleum Research Institute, for encouragement andsupporting me with different facilities.

I am sincerely grateful to Dr. Rehab M. Hafez, BotanyDepartment, Faculty of Science, Cairo University for her help. Iwant to appreciate my colleagues Omaima Sharaf, Mustafa, HananOsman, Akram Hassan, Mohammed Salah and Sara El- Tanany.

My deep thanks to my Parents for their understanding duringmy absence from them and for their support and encouragement tome.

LIST OF ABBREVIATIONS

Atmospheric Pressure Chemical IonizationAPCIBiotic purified starchBiosurfactants

B-PSBS

Cashew Apple JuiceCollision induced dissociationCritical micelle concentration

CAJCIDCMC

Critical Micelle DilutionCMDDiaminobutyric acidDABElectrospray ionisation mass spectrometryESI MSFast Atom BombardmentFABFourier transform infrared spectroscopyFTIRHydrophilic-lipophilic balanceHLBHigh Performance Liquid ChromatographyHPLCHigh solidsHigh TPH-contaminatedLiquid BrothLow solids

HSHTCLBLS

Low TPH-contaminatedLTCMercury cadmium tellurideMicrobial Enhanced Oil RecoveryMicrobial surfactants

MCTMEORMS

Mineral salts mediumMS mediumMethyl tertiary-butyl etherIon mass/ ion chargeNuclear Magnetic Resonance

MTBEm/zNMR

Olive oil mill waste effluentOMWE0-nitrophenyl- -D-galactosidaseONPGPolyaromatic hydrocarbonsPolymeric microbial surfactants

PAHPMS

RhamnolipidRLResponse surface methodologyRSMSodium dodecyl sulfateSophorolipids

SDSSLs

Thin Layer ChromatographyTLCTotal petroleum hydrocarbonsTPHTriple Sugar Iron Agar Wood-based activated carbon

TSI AgarWAC

TABLE OF CONTENTS

PageList of abbreviations List of figuresList of tables .................................................................................. Aim of workIntroduction .. 1Literature Review 32.1 Nature of surfactant 3

..... 42.3 Classification and chemical nature of biosurfactants .. 6

72 .. 7

... 7.. 7

2.3.2. Fatty Acids, Phospholipids, and Neutral Lipids 82.3.3. Polymeric microbial surfactants 82.3.4. Surface active antibiotics .. 92.3.4.1. Surfactin ... 9

.. 9

. 112.4 Economic factors of Biosurfactant production .. 112.5 Substrates for commercial microbial production .. 122.5.1. Agro-industrial wastes .. 122.5.1.1. Use of raw substrates . 132.5.1.1.1. Vegetable oils and oil wastes .. 132.5.1.1.2. Olive oil mill waste effluent (OMWE) 142.5.1.1.3. Starchy substrates ... 142.5.1.1.4. Enzymatic production of biosurfactants 152.5.2. Industrial wastes from animal and plant origin 152.5.2.1. Animal fat . 152.5.2.2. Dairy Industry Whey . 162.5.2.3. Molasses 162.5.2.4. Soy Molasses ... 16

2.5.3. Other industrial wastes . 172.5.3.1. Soap stock . 172.6 Factors affecting biosurfactant production . 172.6.1. Bio-process optimization . 172.6.2. Influence of the Culture Medium Composition onBiosurfactant production ..

18

2.6.2.1. Carbon Source . 192.6.2.2. Nitrogen Source .. 202.6.3. Environmental Factors Affecting the Production 212.6.3.1. pH 212.6.3.2. Temperature . 212.6.3.3. Metal ion concentration 212.6.3.4. Aeration and Agitation .. 222.6.4. Product recovery . 222.6. 5. Metabolic characterization of biosurfactants 232.7 Improved Strains for Biosurfactant Production . 242.8 Methods of Detection and Analysis of rhamnolipids .. 242.8.1. Qualitative Methods .. 242.8.2. Quantitative Methods 252.8.2.1. Thin Layer Chromatography (TLC) 252.8.2.2. Liquid Chromatography 252.8.2.3. Liquid Chromatography Coupled to Mass Spectrometry .. 262.8.2.4. Other Spectroscopic Method ... 262.9 Environmental Applications of Biosurfactants .. 262.10 Biosurfactants and Hydrocarbons Degradation . 28

2.10.1. Role of Biosurfactants in Biodegradation Processes 29

2.10.2. Biodegradation Studies .. 302.10.3. Soil Washing Technology 322.10.4. Microbial Enhanced Oil Recovery (MEOR) . 32

2.11 Industrial applications of biosurfactants . 33

2.12 Biomedical applications . 342.12.1. Antimicrobial activity of biosurfactants .. 362.12.2. Biomedical and therapeutic applications of biosurfactants .. 37Material and methods 39

3.1. Microorganisms and their isolation . 393.2. Identification of micro-organisms . 393.2.1. Gram Stain 393.2.2. Biochemical Tests .. 403.2.2.1 Motility test ... 40

3.2.2.2. Catalase test .. 40

3.2.2.3. Oxidase production test 41

3.2.2.4. Growth at 42oC .. 41

3.2.2.5. Reaction in Triple Sugar Iron Agar . 41

3.2.3. API 20E Identification for selected isolate 413.2.4. 16S rRNA gene sequence analysis for selected isolate 423.3. Production of biosurfactant . 433.3.1. Preparation of Culture medium . 433.3.2. Fermentation medium and condition 433.4. Dried weight cell determination ... 443.5. Extraction of the produced biosurfactants . 443.6. Optimization of Biosurfactant Production . 443.6.1. Optimum Carbon Substrate . 443.6.2. Optimum Nitrogen Source .. 453.6.3. Optimum pH 453.6.4. Optimum Temperature . 453.6.5. Optimum Agitation Rate 453.7. Purification of biosurfactant .. 453.8. Structural characterization of biosurfactant .. 463.8.1. Fourier transform infrared spectroscopy (FTIR) . 463.8.2. Nuclear magnetic resonance analysis .. 463.8.3. Electrospray ionisation mass spectrometry (ESI MS) 473.9. Physicochemical characterization 473.9.1. Oil displacement test . 473.9.2. Surface tension measurement . 473.9.3. Measurement of critical micelle concentration (CMC) 483.9.4. Determination of emulsification index 48Experimental Results 49

4.1. Isolation and identifiction of selected Biosurfactant-producingbacterial strain

49

4.2. Optimization of Biosurfactant Production 534.2.1. Effects of carbon sources on rhamnolipid production 534.2.2. Effects of nitrogen sources on rhamnolipid production 534.2.3. Effect of pH on rhamnolipid production 544.2.4. Effect of temperature on rhamnolipid production 544.2.5. Effect of agitation rate on rhamnolipid production 574.3. Purification of biosurfactant 584.4. Structural characterization 584.4.1. Thin Layer Chromatography 584.4.2. Fourier transform infrared spectroscopy (FTIR) 604.4.3. Nuclear magnetic resonance NMR 604.4.4. Electrospray ionisation mass spectrometry (ESI MS) 604.5. Oil displacement test for par 654.6. Critical micelle concentration of partially purified biosurfactants 654.7. Estimation of Emulsification Activity 70Discussion 72Summary 78References 80Appendix 113

LIST OF FIGURES

No. Subject Page

2.1 Types of biosurfactants produced by differentmicroorganisms. 10

2.2 Accumulation of biosurfactants at the interface betweenliquid and air. 27

2.3 The relationship between biosurfactant concentration,surface tension and formation of micelles. 28

2.4 Mechanisms of hydrocarbon removal by biosurfactantsdepending on their molecular mass and concentration. 29

2.5 Mechanism of enhanced oil recovery by biosurfactants. 33

4.1 Growth of Pseudomonas aeruginosa on blood agar,MacConkey agar, Cetrimide agar, glucose-containing MSmedium and its gram stain. 51

4.2 The neighbor-joining tree based on the 16S rRNAsequence, demonstrating the phylogenetic position ofstrain TMN within Pseudomonas species. 52

4.3 Effect of different carbon sources on the yield and surfacetension. 55

4.4 Effect of different nitrogen sources on the yield andsurface tension. 55

4.5 Effect of different pH on rhamnolipid yield and dry cellweight. 56

4.6 Effect of different temperatures on rhamnolipid yield anddry cell weight. 56

4.7 Effect of different agitation rates on rhamnolipid yieldand dry cell weight. 57

4.8 Thin layer chromatography of crude extract ofbiosurfactant produced by P. aeruginosa TMN. 59

4.9 FTIR spectra of rhamnolipids produced by P. aeruginosaTMN. 61

No. Subject Page

4.10 1H-NMR spectrum of purified biosurfactant produced byP. aeruginosa TMN. 62

4.11 ESI mass spectrum and chemical structure of Rha-Rha-C10 C12 rhamnolipid and its fragments. 64

4.12 Diameters of the clear zones on the oil surface obtainedfrom oil displacement testing with the crude biosurfactantproduced from different carbon and nitrogen sources byPseudomonas aeruginosa TMN. 66

4.13 CMC of partially purified biosurfactant produced usingglucose as carbon source and NaNO3 as nitrogen source. 67

4.14 CMC of biosurfactants produced from different carbonsources. 68

4.15 CMC of biosurfactants produced from different nitrogensources. 69

4.16 Emulsifying activity (E24%) of biosurfactants obtainedfrom different carbon sources against kerosene. 71

4.17 Emulsifying activity (E24%) of biosurfactants obtainedfrom different nitrogen sources against kerosene. 71

LIST OF TABLES

No. Subject Page

2.1 List of biosurfactant producing organisms. 4

2.2 Methods for the recovery of biosurfactants. 23

2.3 Examples of biosurfactant applications in the medical field. 35

3.1 List of primers used in this study. 43

4.1 Biochemical characteristics of the 6 isolates of gramnegative bacteria grown on the API 20E strip and identifiedas Pseudomonas aeruginosa. 50

4.2 Purification of biosurfactant using different mobile phasesand oil displacement test for each fraction. 59

4.3 1H-NMR chemical shift data for rhamnolipid components 62

4.4 Homologues of rhamnolipid produced by Pseudomonasaeruginosa, ausing glucose as carbon source. 63

4.5 CMC of biosurfactants produced from different carbon andnitrogen sources. 67

Appendix Tables 113

A.1 Effect of different carbon sources on the yield and surfacetension. 113

A.2 Effect of different carbon sources on dry cell weight andyield. 113

A.3 Effect of different nitrogen sources on the yield and surfacetension. 114

A.4 Effect of different nitrogen sources on the dry cell weightand yield. 114

A.5 Effect of different PH on rhamnolipid yield and dry cellweight. 115

No. Subject Page

A.6 Effect of different temperatures on rhamnolipid yield anddry cell weight. 115

A.7 Effect of different agitation rates on rhamnolipid yield anddry cell weight. 116

A.8 Effect of different carbon sources on yield and Oildisplacement test 116

A.9 Effect of different nitrogen sources on the yield and Oildisplacement test. 117

A.10 Emulsification index of the carbon sources. 117

A.11 Emulsification index of the nitrogen sources. 118

The present study is designed aiming to use cheap materials could be introduced

from natural sources based on scientific and traditional merits for

bioremediation and oil recovery.

Aim of work

Introduction

CHAPTER I

INTRODUCTION

Microbial surface-active compounds are a group of structurally diverse

molecules produced by different microorganisms and are mainly classified by their

chemical structure and their microbial origin. All biosurfactants (BS) are amphiphiles,

they consist of two parts a polar (hydrophilic) moiety and non polar (hydrophobic)

group. A hydrophilic group consists of mono-, oligo- or polysaccharides, peptides or

proteins and a hydrophobic moiety usually contains saturated, unsaturated and

hydroxylated fatty acids or fatty alcohols (Lang, 2002). A characteristic feature of

biosurfactants is a hydrophilic-lipophilic balance (HLB) which specifies the portion of

hydrophilic and hydrophobic constituents in surface-active substances. These

structures confer a wide range of properties, including the ability to lower surface and

interfacial tension of liquids and to form micelles and microemulsions between two

different phases.

These compounds can be roughly divided into two main classes (Neu, 1996):

low molecular-weight compounds called biosurfactants, such as lipopeptides,

glycolipids, proteins and high-molecular-weight polymers of polysaccharides,

lipopolysaccharides proteins or lipoproteins that are collectively called bioemulsans

(Rosenberg and Ron, 1997) or bioemulsifiers (Smyth et al., 2010a). The former group

includes molecules which can efficiently reduce surface and interfacial tension, while

the latter are amphiphilic and polyphilic polymers which are usually more effective in

stabilising emulsions of oil-in-water but do not lower the surface tension as much

(Smyth et al., 2010b).

The best-studied microbial surfactants are glycolipids. Among these, the best-

known compounds are rhamnolipids (RL), trehalolipids, sophorolipids and

mannosylerythritol lipids, which contain mono- or disaccharides, combined with

long-chain aliphatic acids or hydroxyaliphatic acids. Rhamnolipid production by

Pseudomonas species has been extensively studied, and potential applications have

been proposed (Maier and Soberón-Chávez, 2000).

Introduction

Rhamnolipids from Pseudomonas aeruginosa are currently commercialised by

Jeneil Biosurfactant, USA, mainly as a fungicide for agricultural purposes or an

additive to enhance bioremediation activities. Biosurfactants have many

environmental applications such as bioremediation and dispersion of oil spills,

enhanced oil recovery and transfer of crude oil. Other potential applications of

biosurfactants relate to food, cosmetic, health care industries and cleaning toxic

chemicals of industrial and agricultural origin.

Literature Review

CHAPTER II

LITERATURE REVIEW

2.1 Nature of surfactant

Surfactants are SURFace ACTive AgeNTS with wide ranging properties

including the lowering of surface and interfacial tensions of liquids. Surfactants have

been used industrially as adhesives, flocculating, wetting and foaming agents, de

emulsifiers and penetrants (Mulligan and Gibbs, 1993). They are used for these

applications based on their abilities to lower surface tensions, increase solubility,

detergency power, wetting ability and foaming capacity. The petroleum industry has

traditionally been the major users, as in enhanced oil removal applications. In this

application, surfactants increase the solubility of petroleum components (Falatko,

1991). The effectiveness of a surfactant is determined by its ability to lower the

surface tension, which is a measure of the surface free energy per unit area required to

bring a molecule from the bulk phase to the surface (Rosen, 1978). Efficient

surfactants have a low critical micelle concentration (CMC) (i.e. less surfactant is

necessary to decrease the surface tension). The CMC is defined as the minimum

concentration necessary to initiate micelle formation (Becher, 1965).

In practice, the CMC is also the maximum concentration of surfactant

monomers in water and is influenced by pH, temperature and ionic strength. The

choice of surfactant is primarily based on product cost (Mulligan and Gibbs, 1993). In

general, surfactants are used to save energy and consequently energy costs (such as

the energy required for pumping in pump and treat techniques). Charge-type,

physicochemical behaviour, solubility and adsorption behaviour are some of the most

important selection criteria for surfactants. New markets are currently being

developed for use in the bioremediation of contaminated land sites (Oberbremer et al.,

1990; Samson et al., 1990).

Microbial-derived surfactants or biosurfactants are produced by a wide variety

of microbes and are amphipathic molecules with a hydrophilic and a hydrophobic

domain seem to facilitate the uptake of hydrocarbons into cells. Because of these

traits, biosurfactants accumulate at interfaces, can form micelles, lower the surface

tension and thereby enhance the solubility of poorly soluble compounds in water

Literature Review

(Kuiper et al., 2004). Wide spectra of microbial compounds, including glycolipids,

lipopeptides, fatty acids, and polymeric biosurfactants, have been found to have

surface activity (Morikawa et al., 2000).

Most of these compounds are either anionic or neutral. Only a few are cationic

such as those containing amine groups. The hydrophobic part of the molecule is based

on long- -alkyl- -hydroxy fatty acids. The

hydrophilic portion can be a carbohydrate, amino acid, cyclic peptide, phosphate,

carboxylic acid or alcohol. A wide variety of microorganisms can produce these

compounds.

The CMCs of the biosurfactants generally range from 1 to 200 mg/l and their

molecular mass is from 500 to 1500 Da (Lang and Wagner, 1987). Biosurfactants

have important advantages, such as biodegradability, low toxicity, and various

possible structures, relative to chemically synthesized surfactants (Benincasa et al.,

2002). For example, glycolipids from Rhodococcus species 413A were 50% less toxic

than Tween 80 in naphthalene solubilization tests (Kanga et al., 1997).

2.2 Microorganisms producing biosurfactants

Biosurfactants produced by a variety of microorganisms mainly bacteria, fungi

and yeasts are diverse in chemical composition and their nature and the amount

depend on the type of microorganism producing a particular biosurfactant as shown in

Table (2.1) (Saharan et al., 2011). Many microorganisms for industrial utilization for

waste products have been isolated from contaminated soils, effluents and waste water

sources. Thus, these have an ability to grow on substrates considered potentially

noxious for other non-producing microorganisms.

Table (2.1) List of biosurfactant producing organisms (Saharan et al., 2011)

Sr.No.

Biosurfactant Microorganism(s) Current economicImportance

Reference(s)

1. Cellobiose lipids Ustilago maydis AntifungalCompounds

(Teichmannet al., 2007)

2. Serrawettin Serratia marcescens Emulsification ofHydrocarbons

(Lai et al.,2009)

3. Polyol lipids Rhodotorulaglutinis, R. graminis

Anti-proliferativeActivity

(Amaral etal., 2006)

4. Trehalose lipids Rhodococcus Dissolution of (Muthusamy

Literature Review

erythropolis,Arthrobacter sp.,Nocardiaerythropolis,Corynebacteriumsp.,Mycobacterium sp

Hydrocarbons et al., 2008)

5. Ornithine lipids Pseudomonas sp.,Thiobacillusthiooxidans,Agrobacterium sp.

Bio-emulsifiers (Desai andBanat, 1997)

6. Viscosin Pseudomonasfluorescens,Leuconostocmesenteriods

Surface activeLipopeptides

(Banat et al.,2010)

7. Rhamnolipids Pseudomonasaeruginosa,Pseudomonaschlororaphis,SerratiaRubidea

Bioremediation,Antimicrobial andbiocontrol properties

(Jadhav etal., 2011)

8. Carbohydrate-lipid

P.fluorescens,DebaryomycesPolmorphus

Bio-emulsifiers (Nerurkar etal., 2009)

9. Protein PA P.aeruginosa Bio-emulsifiers (Hisatsuka etal., 1971)

10. DiglycosylDiglycerides

Lactobacillusfermentum

Bio-remediation (Mulligan etal., 2001)

11. Whole cell Cyanobacteria Bio-flocculent (Levy et al.,1990)

12. Fatty acids/neutralLipids

Clavibactermichiganensis subsp.Insidiosus

Bio-emulsifiers (Herman andMaier, 2002)

13. Sophorolipids Candida bombicola,C. antartica,Torulopsispetrophilum C.botistae,C. apicola, C.riodocensis, C.stellata, C. bogoriensis

Antimicrobial,Antiviral,Spermicidal

(Felse et al.,2007)

14. Liposan C. tropicalis Bio-emulsan (Ciriglianoand Carman,1984)

15. MonnosylerythritolLipids

C.antartica,Kurtzmanomyces sp.,Pseudozymasiamensis

Antifungalcompounds

(Kitamoto etal., 1993)

16. Surfactin/Iturin B. subtilis, B. Antimicrobial (Arguelles-

Literature Review

amyloliquefaciens Properties Arias et al.,2009)

17. Subtilisin B. subtilis AntimicrobialProperties

(Sutyak etal., 2008)

18. Aminoacidslipids

Bacillus sp. AntimicrobialProperties

(Cotter et al.,2005)

19. Lichenysin Bacilluslicheniformis, B.subtilis

Microbiallyenhancedoil recovery(MEOR)

(Yakimov etal., 1997)

20. Peptide lipids B. licheniformis AntimicrobialProperties

(Begley etal., 2009)

21. Phospholipids Acinetobacter sp. Bioremediation (Kosaric,2001)

22. Vesicles &fimbriae

Acinetobactercalcoaceticus, P.marginilis, P.Maltophila

Bioremediation (Desai andBanat, 1997)

23. Emulsan A. calcoaceticus Microbiallyenhancedoil recovery(MEOR)

(Choi et al.,1996)

24. Alasan A. radioresistens Biodegradation ofpolyaromaticcompounds

(Barkay etal., 1999)

2.3 Classification and chemical nature of biosurfactants

The microbial surfactants (MS) are complex molecules covering a wide range

of chemical types including peptides, fatty acids, phospholipids, glycolipids,

antibiotics, lipopeptides, etc. Microorganisms also produce surfactants that are in

some cases combination of many chemical types: referred to as the polymeric

microbial surfactants (PMS). Many MS have been purified and their structures

elucidated. While the high molecular weight MS are generally polyanionic

heteropolysaccharides containing both polysaccharides and proteins, the low

molecular weight MS are often glycolipids. Various types of biosurfactant produced

from different organisms are shown in Figure (2.1).

Literature Review

2.3.1. Glycolipids

Glycolipids are the most common types of BS (Karanth et al., 1999). The

constituent mono, di-, triandtetrasaccharides include glucose, mannose, galactose,

glucuronic acid, rhamnose, and galactose sulphate. The fatty acid component usually

has a composition similar to that of the phospholipids of the same microorganism.

The glycolipids can be categorized as:

2.3.1.1. Trehalose lipids

Several structural types of microbial trehalolipids biosurfactants have been

reported. Disaccharide trehalose linked at C-6 and C-6' to mycolic acids is associated

with most species of Mycobacterium, Nocardia, and Corynebacterium. Mycolic acids

-branched- -hydroxy fatty acids. Trehalolipids from different

organisms differ in the size and structure of mycolic acid, the number of carbon

atoms, and the degree of unsaturation (Cooper et al., 1989). Trehalose dimycolate

produced by rhodococcus erythropolis has been extensively studied. R.erythropolis

also synthesizes a novel anionic trehalose lipid. Trehalose lipids from R.erythropolis

and Arthrobacter sp. lowered the surface and interfacial tensions in the culture broth

to 25 to 40 and 1 to 5 mN/m, respectively (Rapp et al., 1979; Kretschmer et al., 1982;

Li et al., 1984).

2.3.1.2. Sophorolipids

These are produced by different strains of the yeast, Torulopsis. The sugar unit

is the disaccharide sophorose which consists of two b -1,2-linked glucose units. The 6

and 6¢ hydroxy groups are generally acetylated. The sophorolipids reduce surface

tensions between individual molecules at the surface, although they are effective

emulsifying agents. The sophorolipids of Torulopsis have been reported to stimulate,

inhibit, and have no effect on growth of yeast on water-insoluble substrates (Karanth

et al., 1999).

2.3.1.3. Rhamnolipids

A group of biosurfactants that has been studied extensively is the rhamnolipids

from Pseudomonas aeruginosa (Hitsatsuka et al., 1971, Guerra-santos et al., 1984).

Up to seven homologues have now been identified (Abalos et al., 2001). Surface

tensions of 29 mN/m are characteristic of these compounds. Two types of

Literature Review

-hydroxy decanoic acid

or one rhamnose connected to the identical fatty acid.

The hydroxyl group of one of the acids is involved in glycosidic linkage with

the reducing end of the rhamnose disaccharide, whereas the hydroxyl group of the

second acid is involved in ester formation. Since one of the carboxylic groups is free,

the rhamnolipids are anions above pH 4. Rhamnolipid was reported (Hitsatsuka et al.,

1971) to lower surface tension, emulsify hydrocarbons and stimulate growth on n-

hexadecane of P. aeruginosa strains but not other hydrocarbon-degrading bacteria.

The pure rhamnolipid lowered the interfacial tension against n-hexadecane to

about 1 mN/m and had a CMC of 10 to 30 mg/l, depending on the pH and salt

conditions (Lang and Wagner, 1987; Parra et al., 1989). Zhang and Miller, (1995)

reported that the interfacial tension between hexadecane and water was decreased to

less than 0.1 mN/m by a methyl ester of rhamnolipid, whereas the free acid decreased

the interfacial tension only to 5 mN/m.

2.3.2. Fatty Acids, Phospholipids, and Neutral Lipids

Several bacterial and yeast produce large quantities of fatty acid and

phospholipid surfactants during growth on n-alkanes (Ciriglino and Garman, 1985).

The HLB is directly related to the length of the hydrocarbon chain in their structures.

In Acinetobacter sp. Strain HO1-N phosphatidulethanolamine, rich vesicles are

produced (Gautam and Tyagi, 2006), which form optically clear microemulsions of

alkanes in water. Phosphatidulethanolamine produced by R. erythropolis grown on n-

alkane caused a lowering of interfacial tension between water and hexadecane to less

than 1 mN/m and a CMC of 30 mg/l (Kretschmer et al., 1982).

2.3.3. Polymeric microbial surfactants

The high molecular weight microbial surfactants are generally polyanionic

heteropolysaccharides containing both polysaccharides and proteins. The best studied

polymeric biosurfactants are emulsan, liposan, mannoprotein, and other

polysaccharide protein complexes. Acinetobacter calcoaceticus RAG-1 produces a

potent polyanionic amphipathic heteropolysaccharide bioemulsifiers (Rosenberg et

al., 1979).

Literature Review

Emulsan is a very effective emulsifying agent for hydrocarbons in water even at a

concentration as low as 0.001 to 0.01%. It is one of the most powerful emulsion

stabilizers known today and resists inversion even at a water to-oil ratio of 1:4.

Liposan is an extracellular water-soluble emulsifier synthesized by Candida

lipolytica and is composed of 83% carbohydrate and 17% protein (Cirigliano and

Carman, 1984). Cameron et al. (1988) reported the production of large amounts of

mannoprotein by Saccharomyces cerevisiae; this protein showed excellent emulsifier

activity toward several oils, alkanes, and organic solvents.

2.3.4. Surface active antibiotics

2.3.4.1. Surfactin

Bacillus subtilis produces a cyclic lipopeptide called surfactin or subtilysin

(Arima et al., 1968; Bernheimer and Avigad, 1970; Peypoux et al., 1999), which is

reputedly the most active biosurfactant known to date (Cooper and Zajic, 1980).

Surfactin has a CMC in water of 25 mg/l and lowers the surface tension to 27 mN/m.

The minimum interfacial tension against hexadecane was 1 mN/m. The yield of

surfactin produced by B. subtilis can be improved to around 0.8 g/l by continuously

removing the surfactant by foam fractionation and addition of either iron or

manganese salts to the growth medium (Cooper et al., 1981).

A recombinant B. subtilis, carrying a gene for surfactin production, produced

new surfactin variants (Nakayama et al., 1997). The amphipathic nature of surfactin

may contribute to some of its interesting biological properties, such as the formation

of ion-conducting pores in membranes (Grau et al., 1999).

2.3.4.2. Polymixins

These are a group of antibiotics produced by Brevibacterium polymyxa and

related bacilli. Polymixin B is a decapeptide in which amino acids 3 through 10 form

a cyclic octapeptide. A branched chain fatty acid is connected to the terminal 2, 4-

diaminobutyric acid (DAB). Polymixins are able to solubilize certain membrane

enzymes (Rosenberg et al., 1979).

Literature Review

A. Trehalolipid B. Sophorolipid

C. Structure of 4 different rhamnolipids produced by P. aeruginosa.

D. phosphatidulethanolamine. F. Emulsan.

Figure (2.1) Types of biosurfactants produced by different microorganisms(Desai and Banat, 1997).

Literature Review

2.3.4.3. Antibiotic TA

Myxococcus xanthus produces antibiotic TA which inhibits peptidoglycan

synthesis by interfering with polymerization of the lipid disaccharide pentapeptide

(Cooper et al., 1981). Antibiotic TA has interesting chemotherapeutic applications

(Karanth et al., 1999).

2.4 Economic factors of Biosurfactant production

Economical large scale production for established and new applications of

biosurfactants remains a challenge (Bognolo, 1999). Researchers have emphasized the

key parameters affecting the efficiency of biosurfactant production in terms of higher

yields and lower production costs (Kosaric, 1992; Bognolo, 1999; Mukherjee et al.,

2006). According to them the biosurfactant formation and accumulation follows the

basic facts of metabolic process and need to be studied comprehensively. According

to Syldatk and Hausmann, (2010) the reasons for limited use of microbial surfactants

in industry are the use of expensive substrates, limited product concentrations, low

yields and formation of product mixtures rather than pure compounds. All these

factors and other growth and upscale problems like use of antifoaming agents add on

to the high costs of the downstream processing.

To overcome the expensive cost constraints associated with biosurfactant

production, two basic strategies are generally adopted worldwide to make it cost-

effective:

(i) The use of inexpensive and waste substrates for the formulation of

fermentation media which lower the initial raw material costs involved in

the process.

(ii) Development of efficient and successfully optimized bioprocesses,

including optimization of the culture conditions and cost-effective

recovery processes for maximum biosurfactant production and recovery.

As millions of tons of hazardous and non-hazardous wastes are generated each

year throughout the world, a great need exists for their proper management and

utilization. It has been suggested that successful approaches to more economical

production technologies of biosurfactant will be a collaborative approach involving

process development and sustainable raw materials supplies. According to Smyth et

Literature Review

al. (Smyth et al., 2010a, b) emphasis should be on cost effective management of

downstream processing. The potential to obtain pure biosurfactants is dependent on

several complex extraction and purification steps. Use of simple substrates with less

downstream processing will economize the process and the use of agricultural

substrates and their wastes represents a positive step towards achieving that goal.

Some of the prevalent downstream processing process uses solvent extraction

(e.g. chloroform-methanol, dichloromethane-methanol, butanol, ethyl acetate,

pentane, hexane, acetic acid, ether) or acid precipitation, use of ammonium sulfate

precipitation, crystallization, centrifugation, adsorption and foam fractionation

(Martins et al., 2006; Mukherjee et al., 2006; Chen et al., 2008; Kaar et al., 2009;

Neto et al., 2009). More studies are needed on these processes for efficient production

of biosurfactants. The availability of processes with limited downstream processing

will give significant economical advantages and have been sought after.

2.5 Substrates for commercial microbial production

Despite possessing many industrially attractive properties and advantages

compared with synthetic ones, the production of biosurfactants on industrial scale has

not been undertaken due to high investment costs. This necessitates their profitable

production and recovery on a large scale. Various aspects of biosurfactants, such as

their biomedical and therapeutic properties (Cameotra and Makkar, 2004) their

natural roles (Ron and Rosenberg, 2001), their production on inexpensive alternative

substrates and their industrial potential, have been reviewed (Makkar and Cameotra,

2002). However their cost of production continues to remain very high. Using low-

cost raw materials is a possible solution for this obstacle (Makkar and Cameotra,

2002). Another approach is to use renewable low cost starting materials from various

sources including industrial wastes from frying oils, oil refinery wastes, molasses,

starch rich wastes, cassava waste water and distilled grape marc (Nitschke and

Pastore, 2006; Rivera et al., 2007; Sobrinho et al., 2008) .

2.5.1. Agro-industrial wastes

These wastes are obtained at low cost from the respective processing

industries and are as potent as low-cost substrates for industrial level biosurfactant

production. Agricultural wastes such as rice water and water from the processing of

Literature Review

cereals, pulse and molasses have potential to be used as excellent substrates for the

production of biosurfactants. P. aeruginosa can be cultivated in Cashew Apple Juice

(CAJ) supplemented with peptone (5.0 g/l) and nutritive broth to obtain surfactants.

Surface tension during the fermentation can be reduced by 41% when P. aeruginosa

is cultivated in CAJ supplemented with peptone (Maria et al., 2007) compared to

other amino acid sources.

2.5.1.1. Use of raw substrates

2.5.1.1.1. Vegetable oils and oil wastes:

Several studies with plant-derived oils have shown that they can act as

effective and cheap raw materials for biosurfactant production, for example, rapeseed

oil (Trummler et al., 2003), Babassu oil and corn oil (Vance-Harrop et al., 2003;

Pekin et al., 2005). Similarly, vegetable oils such as sunflower and soybean oils

(Ferraz et al., 2002; Rahman et al., 2002; Kim et al., 2006) were used for the

production of rhamnolipid, sophorolipid and mannosylerythritol lipid biosurfactants

by various microorganisms. Apart from various vegetable oils, oil wastes from

vegetable oil refineries and the food industry were also reported as good substrates for

biosurfactant production. In addition, industrial oil wastes such as tallow, soapstock,

marine oils, lard and free fatty acids can potentially induce microbial growth and

metabolite production owing to their typical fatty acid composition. Furthermore,

various waste oils with their origins at the domestic level, in vegetable oil refineries or

the soap industries were found to be suitable for microbial growth and biosurfactant

production (Haba et al., 2000; Abalos et al., 2001; Benincasa et al., 2002; Bednarski

et al., 2004; Benincasa et al., 2004; Nitschke et al., 2005).

These oils and oil wastes are readily available in good amounts throughout the

world. However, the oils used to date for biosurfactant production are mostly edible

oils and are not cheap. Several plant-derived oils, for example, jatropha oil, mesua oil,

castor oils, ramtil oil and jojoba oil, are not suitable for human consumption due to

their unfavourable odour, colour and composition and are, therefore, available at

much cheaper rates. Incorporation of these cheaper oils and oil wastes in the industrial

production media might potentially reduce the overall costs of biosurfactant

production.

Literature Review

2.5.1.1.2. Olive oil mill waste effluent (OMWE):

which is estimated at over 2.5 million metric tons per year with about 75% being

produced in the European Union. The process of olive oil extraction results in a large

amount of liquid waste. OMWE are characterized by an intensive dark brown color, a

strong acidic smell and a high organic content (COD 220 g/l). OMWE is a black

liquor and consists of a high content of organic matter (20-60 kg COD/m3), depending

on the olive oil extraction procedure (Marques, 2001). OMWE contains toxic

substances such as polyphenols (Hamman et al., 1999) making it unsuitable for

human consumption in raw or processed form but has valuable organic substances

such as sugars, nitrogen compounds, organic acids and residual oils which aid

microbial growth. The large diversity of components found in OMWE (carbohydrates,

polysaccharides, sugars, lipids and phenolic compounds) makes their treatment

difficult, and their disposal becomes a critical environmental problem (Niaounakis

and Halvadakis, 2004).

Thus, utilization of these materials is important from both environmental and

economical points of view and can be considered not only as a waste to be treated but

also a resource to be recovered. The use of fungi can lead to OMWE valorization

through the enzymes production (Crognale et al., 2006). The anaerobic treatment of

OMWE can also represent an effluent recovery and methane production. Yeasts can

also be used to degrade the phenolic compounds in OMWE. Specifically, Yarrowia

lipolytica strains are good candidates for the OMWE treatment and recovery

(Papanikolaou et al., 2008).

2.5.1.1.3. Starchy substrates:

Starchy waste materials are inexpensive raw materials suitable for the

production of surface active agents. A major source of inexpensive starchy substrate

is the potato processing industry. Potatoes are generally composed of 80% water, 17%

carbohydrates, 2% protein, 0.1% fat and 0.9% vitamins, inorganic minerals and trace

elements. They are a rich source of carbon (in the form of starch and sugars), nitrogen

and sulfur (from protein), inorganic minerals, trace elements and vitamins. Thompson,

(2000) reported the use of high solids (HS) and low solids (LS) potato effluents as

substrates for Surfactin production. Growth rate was higher in all HS- and LS-based

Literature Review

media than in the B-PS (biotic purified starch) control. Potato process effluents

(wastes from potato processing industries) can also be used to produce biosurfactant

by B. subtilis (Noah, 2005).

Cassava wastewater is another carbohydrate-rich residue generated in large

amounts during the preparation of cassava flour and is an attractive alternative

substrate in fermentation processes. It has been used for surfactin production by B.

subtilis (Nitschke and Pastore, 2006). Siddhartha et al. (2009) used Cassava

wastewater as a substrate for the simultaneous production of rhamnolipids and

polyhydroxyalkanoates by P. aeruginosa.

2.5.1.1.4. Enzymatic production of biosurfactants:

Polyglycerol and carbohydrate fatty acid esters are broadly used as industrial

detergents and as emulsifiers in a huge range of food formulations. Adelhorst et al.

(1990) have carried solvent-free esterification of simple alkyl-glycosides by means of

molten fatty acids and immobilized C. antarctica lipase. Fregapane et al. (1991)

obtained mono- and diesters of monosaccharides in elevated yields, with sugar acetyls

as preliminary resource.

Lipase from A. terreus synthesizes a biosurfactant by transesterification

involving natural oils and sugar alcohols (Yadav et al., 1997). Lipases may

furthermore substitute phospholipases in the production of lysophospholipids. Mucor

miehei lipase has been used for the transesterification of phospholipid in a variety of

primary- and secondary alcohols (Montet et al., 1990). Lipases may also be valuable

in the synthesis of an entire range of amphoteric biodegradable surfactants,

specifically amino acid-based esters and amides.

2.5.2. Industrial wastes from animal and plant origin

2.5.2.1. Animal fat:

Animal fat and tallow can be obtained in large quantities from meat processing

industries and have been used as a cooking medium for foods. Deshpande and

Daniels, (1995) used animal fat for the production of sophorolipids biosurfactant

production using the yeast, C. bombicola. When fat was provided as the sole carbon

source, the growth was poor. However, a mixture of 10% glucose and 10% fat gave

Literature Review

the highest level of growth indicating the requirement of an additional carbon source

in the medium.

2.5.2.2. Dairy Industry Whey:

The whey from dairy industries is also a cheap and viable substrate for

biosurfactant fermentation. About 6 liters of whey is produced per kg of paneer

(cheese). A large portion of whey from the organized dairy sector is not utilized and is

being disposed through the effluent treatment systems though it contains valuable

nutrients (proteins, peptides, amino acids, lipids, minerals and vitamins).Thus, the

effluent from the dairy industry, known as dairy wastewater supports good microbial

growth and can be used for biosurfactant production (Dubey and Juwarkar, 2004;

Siddhartha et al., 2009). Daniel et al. (1998) used dairy wastes as substrates and

achieved production of high concentrations of sophorolipids using two-stage

cultivation process for the yeast Cryptococcus curvatus ATCC 20509.

2.5.2.3. Molasses:

This is a co-product of sugar industry generated during sugar manufacturing

from either sugarcane or sugar beet and is a rich source of available carbon. Average

values for the constituents of cane molasses (75% dry matter) are: 48-56% (total

sugar), 9-12% (organic matter excluding sugar), 2.5% (protein), 1.5-5.0% (Potassium)

0.4-0.8% (Calcium), 0.06% (Magnesium), 0.06-2.0% (Phosphorus), 1.0-3.0 mg/kg

(biotin), 15-55 mg/kg (pantothenic acid), 2,500-6,000 mg/kg (inositol) and 1.8 mg/kg

(thiamine). Patel and Desai, (1997) used molasses and corn-steep liquor as the

primary carbon and nitrogen source to produce rhamnolipid biosurfactant using P.

aeruginosa (Strain GS3). The biosurfactant production reached a maximum when a

combination of 7% (v/v) molasses and 0.5% (v/v) corn-steep liquor waste used.

2.5.2.4. Soy Molasses:

Soy molasses is a cheap feedstock as it is a low-value co-product of soybean

processing and also rich in potentially fermentable sugar content and other growth

factors useful for sustaining microbial growth. As health-conscious consumers

continue to drive up the demands for soy protein-based foods and drinks, the soy

protein industry has experienced a sustained 10% annual growth for the past several

years with a market value of nearly $4 billion (Deak and Johnson, 2006). Thus, an

increasing amount of agricultural wastes from soy cultivation is becoming available

Literature Review

as a raw material for utilization in biosurfactant production. The major components of

the soluble carbohydrates in soy molasses are sucrose, raffinose and stachyose.

Investigators have demonstrated that soy molasses could be used in fermentation

processes to produce industrial chemicals such as lactic acid, butanol (Qureshi et al.,

2001), sophorolipids biosurfactant (Solaiman et al., 2004) and poly-

hydroxyalkanoates (Solaiman et al., 2006). Daniel et al. (1998) formulated a soy

molasses-based medium for reduced cost production of sophorolipids biosurfactant by

C . bombicola.

2.5.3. Other industrial wastes

2.5.3.1. Soap stock:

Soap stock is a gummy, amber colored by-product of oilseed processing

produced when hexane and other chemicals are used to extract and refine edible oil

from the oilseeds. It has been used to produce emulsan and bio-dispersan. Emulsan

forms and stabilizes oil-inwater emulsion, whereas bio-dispersan disperses the large

solid limestone granules, forming micrometer-size water suspension (Rosenberg et

al., 1988). P. aeruginosa strain LBI, isolated from petroleum contaminated soil, could

produce surface-active rhamnolipids biosurfactant (RLLBI) by batch fermentation in

a mineral salts medium with soapstock as the sole carbon source (Benincasa et al.,

2002). Biosurfactant production increased after nitrogen depletion and the maximum

rhamnolipids concentration was 15.9 g/l. RLLBI produced stable emulsions with

hydrocarbons (crude oil, kerosene, toluene, n-alkanes (C12-C14) and mineral oil) and

vegetable oils (linseed oil, almond oil).

2.6 Factors affecting biosurfactant production

2.6.1. Bio-process optimization

Type, quality and quantity of biosurfactant production is dependent on the

cultural conditions i.e. pH, temperature, agitation, aeration, dilution rate, the

concentration of metal ions, the nature of the carbon and nitrogen sources. There are

lots of studies regarding biosurfactant production relating the optimization of their

physicochemical properties (Sarubbo et al., 2001). Environmental factors are

exceptionally significant in the yield and characteristics of the biosurfactant produced.

Literature Review

In order to acquire large quantities of biosurfactant, it is essential to optimize the

process conditions.

Several elements, media compositions and precursors affect the process of

biosurfactant production. Different elements such as nitrogen, iron, and manganese

affect the production of biosurfactants. Limitation of nitrogen enhances biosurfactant

production in P. aeruginosa strain BS-2 (Dubey and Juwarkar, 2001) and U. maydis

(Hewald, 2005). Addition of iron and manganese to the culture medium increased the

production of biosurfactant by B. subtilis. The classical method of medium

optimization involves changing one variable at a time, while keeping the others at

fixed levels; however, this method is time consuming and does not guarantee the

optimal metabolite production.

A statistical optimization strategy response surface methodology (RSM) has

been developed for the optimization of process. RSM explores the relationships

between several explanatory variables and one or more response variables. This

method could be used to determine the optimum media, inoculum and environmental

conditions for the enhanced production of surfactin by B. subtilis (Sen and

Swaminathan, 2005). RSM has also been applied to enhance biosurfactant production

by P. aeruginosa AT10 (Abalos et al., 2002).

Such optimization methods would help the industry to design the best

combination of cheaper substrates for media production and to use the most favorable

environmental conditions for improved biosurfactant production. Current

developments in the area of optimization of fermentation conditions have resulted in a

considerable enhancement in production yields, making them more commercially

attractive.

2.6.2. Influence of the Culture Medium Composition on

Biosurfactant production

The use of different carbon sources alters the structure of the biosurfactant

produced and its properties and can be exploited to get products with desired

properties for particular applications. There are a number of studies in biosurfactant

production involving the optimization of their physicochemical properties (Sarubbo et

Literature Review

al., 2001; Sarubbo et al., 2006). The composition and characteristics of biosurfactants

are influenced by the nature of the nitrogen source as well as the presence of iron,

magnesium, manganese, phosphorus and sulphur in the media.

2.6.2.1. Carbon Source

Till date, biosurfactants are unable to compete inexpensively with chemically

synthesized compounds due to their high production costs and recovery system. These

costs may be significantly reduced by the use of alternative sources of nutrients.

Water-soluble carbon sources such as glycerol, glucose, mannitol, and ethanol were

all used for rhamnolipid production by Pseudomonas sp.

Biosurfactant product, however, was inferior to that obtained with water-

immiscible compounds such as n-alkanes and olive oil (Robert et al., 1989). Syldatk

et al., (1985a) demonstrated that although different carbon sources in the medium

affected the composition of biosurfactant production in Pseudomonas sp., substrates

with different chain lengths exhibited no effect on the chain length of fatty acid

moieties in glycolipids. On the other hand, Neidleman and Geigert, (1984) showed

evidence for qualitative variation, reflecting the carbon number of alkane for

biosurfactant production in Acinetobacter sp. strains H13-A and HO1-N, respectively.

When Arthrobacter paraffineus ATCC 19558 was grown on D-glucose,

supplementation with hexadecane in the medium during the stationary growth phase

resulted in a significant increase in biosurfactant yield (Duvnjak et al., 1982).

Others observed a little biosurfactant production, when cells were growing on

a readily available carbon source, only when all the soluble carbon was consumed and

when water-immiscible hydrocarbon was available was biosurfactant production

triggered (Banat et al., 1991; Banat, 1995).

Cavalero and Cooper, (2003) have shown that the sophorolipid yield from C.

bombicola ATCC 22214 increases with the n-alkane chain length (from C12 to C15).

This indicated that different microbes respond differently to the carbon sources. The

soy molasses, a byproduct from the production of soybean oil, plus oleic acid were

tested as carbon sources for the production of sophorolipids (SLs) by the yeast C.

bombicola (Solaiman et al., 2004). The purified SLs were obtained at 21 g/l and were

97% in lactone form. The surface properties of the SLs obtained from the soy

Literature Review

molasses/oleic acid fermentation had minimum surface-tension values of 37 mN/m

(pH 6) and 38 mN/m (pH 9), and critical micelle concentration values of 6 mg/l (pH

6) and 13 mg/l (pH 9).

The described C-sources, such as glucose, glycerol, acetates and other organic

acids, as well as pure n-alkanes are quite expensive and cannot reduce the cost of

biosurfactant production. An approach to lessen the cost is partial or complete

replacement of pure reagents with industrial/agricultural mixtures.

2.6.2.2. Nitrogen Source

Medium constituents other than carbon source also affect the production of

biosurfactants. Among the inorganic salts tested, ammonium salts and urea were

preferred nitrogen sources for biosurfactant production by Arthrobacter paraffineus,

whereas nitrate supported maximum surfactant production by Pseudomonas

aeruginosa (Guerra-Santos et al., 1986) and Rhodococcus sp. (Abu-Rawaida et al.,

1991a). Biosurfactant production by Arthrobacter paraffineus is increased by the

addition of amino acid such as aspartic acid, glutamic acid, asparagine, and glycine to

the medium.

Robert et al. (1989) and Abu-Ruwaida et al. (1991a), observed nitrate to be

the best source of nitrogen for biosurfactant production by Pseudomonas strain 44T1

and Rhodococcus strain ST-5 growing on olive oil and paraffin, respectivly. Similarly,

nitrogen limitation caused increased biosurfactant production in Pseudomonas

aeruginosa (Ramana and Karanth, 1989), Candida tropicalis IIP-4 (Singh et al.,

1990), and Nocardia strain SFC-D (Kosaric et al., 1990).

Syldatk et al. (1985b) showed that nitrogen limitation not only caused

overproduction of biosurfactant but also changed the composition of the biosurfactant

produced. Guerra-Santos et al. (1986), showed maximum rhamnolipid production

after nitrogen limitation at a C: N ratio of 16:1 to 18:1 and no surfactant production

below a C: N ratio of 11:1, where the culture was not nitrogen limited. According to

Hommel et al. (1987) it was the absolute quantity of nitrogen and not its relative

concentration that appeared to be important for optimum biomass yield, while

concentration of hydrophobic carbon source determines the conversion of carbon

available to the biosurfactant.

Literature Review

2.6.3. Environmental Factors Affecting the Production

Environmental factors are extremely important in the yield and characteristics

of the biosurfactant produced. In order to obtain large quantities of biosurfactant it is

necessary to optimize the process conditions because the production of a biosurfactant

is affected by variables such as pH, temperature, aeration and agitation speed.

2.6.3.1. pH

The pH of the medium plays an important role in sophorolipid production by

Torulopsis bombicola (Gobbert et al., 1984). Rhamnolipid production in

Pseudomonas sp. was at its maximum at a pH range from 6 to 6.5 and decrease

sharply above pH 7(Guerra-Santos et al., 1984). In contrast, Powalla et al. (1989)

showed that penta- and disaccharide lipid production in Nocardia corynbacteroides is

unaffected in the pH range of 6.5 to 8. In addition, surface tension and CMC of a

biosurfactant product remained stable over a wide range of pH values, whereas

emulsification had a narrower pH range (Abu-Rawaida et al., 1991b).

2.6.3.2. Temperature

In Arthrobacter paraffineus and Pseudomonas sp. strain DSM-2874 (Syldatk

et al., 1985b) temperature caused alteration in the composition of the biosurfactant

produced. A thermophilic Bacillus sp. grew and produced biosurfactant at temperature

above 40oC. Heat treatment of some biosurfactant caused no appreciable change in

biosurfactant properties such as the lowering of surface tension and interfacial tension

and the emulsification efficiency, all of which remained stable after autoclaving at

120 oC for 15 min (Abu Rawaida et al., 1991b).

2.6.3.3. Metal ion concentration

Metal ions concentrations play a very important role in the production of some

biosurfactants as they form important cofactors of many enzymes. The

overproduction of surfactin biosurfactant occurs in presence of Fe2+ in mineral salt

medium. The properties of surfactin are modified in the presence of inorganic cations

such as overproduction (Thimon et al., 1992).

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2.6.3.4. Aeration and Agitation

An increase in agitation speed results in the reduction of biosurfactant yield

due to the effect of shear in Nocardia erythropolis (Margaritis et al., 1979). While

studying the mechanism of biosurfactant production in Acinetobacter calcoaceticus

RAG-1, Wang and Wang, (1990), revealed that the cell-bound polymer/dry-cell ratio

decrease as the shear stress increase. On the other hand, in yeast, biosurfactant

production increases when the agitation and aeration rates increased. Sheppard and

Cooper, (1990) had concluded that oxygen transfer was one of the Key parameters for

the process optimization and scale-up of surfactin production in Bacillus subtilis.

2.6.4. Product recovery

Even if optimum production is obtained using optimal media and cultural

conditions, the production process is still incomplete without an efficient and

economical means for the downstream processing. For many microbiological

products, the downstream processing costs account for 60% of the total production

costs. Several methods have been developed for improving the recovery of

Biosurfactants Table (2.2) (Saharan et al., 2011). These procedures take advantage of

some of the properties of biosurfactants such as their surface activity or their ability to

form micelles and are particularly applicable for large-scale continuous recovery of

extracellular biosurfactants from culture broth.

A few examples of such biosurfactant recovery strategies include foam

fractionation (Sarachat et al., 2010), ultra-filtration (Sen et al., 2010), adsorption-

desorption on polystyrene resins and ion exchange chromatography (Reiling, 1986),

and adsorption-desorption on wood-based activated carbon (WAC) (Dubey, 2005)

Ammonium sulphate precipitation and selective crystallization (Satpute et al., 2010).

Cheap and less toxic solvents such as methyl tertiary-butyl ether (MTBE) have been

successfully used in recent years to recover biosurfactants produced by Rhodococcus

(Kuyukina, 2001). These types of low cost, less toxic and highly available solvents

can be used to cut the recovery expenses substantially and minimize the

environmental hazards.

Literature Review

Table (2.2) Methods for the recovery of biosurfactants (Saharan et al., 2011)

Sr. No

Method(s) Mechanism(s) Reference(s)

1. Adsorption on wood Adsorption (Dubey, 2005;Heyd et al., 2008)

2. Adsorption on Polystyrene Adsorption (Reiling, 1986)3. Ion exchange Chromatography Charge separation (Reiling, 1986)4. Solvent extraction Dissolves in organic

solvents(Kuyukina,2001)

5. Centrifugation Due to Centrifugal force (Nitschke et al.,2003)

6. Acid Precipitation Insoluble at low Ph (Sen andSwaminathan,2004)

7. Membrane Ultra filtration Micelles formation (Sen andSwaminathan,2005)

8. Selective Crystallization Redissolution in organicSolvents

(Satpute et al.,2010)

9. Ammonium Sulphateprecipitation

Salting out of protein (Satpute et al.,2010)

10. Organic Solvent extraction Solubility in organicsolvents

(Dubey, 2005)

11. Foam fractionation Surface activity (Sarachat et al.,2010)

12. Thin layer chromatography Difference in relative flowagainst Solvent

(Priya andUsharani, 2009)

13. Dialysis Difference in soluteconcentration

(Satpute et al.,2010)

14. Lyophilization Cryodesiccation (Satpute et al.,2010)

15. Iso-electric focusing Electric charge difference (Satpute et al.,2010)

2.6. 5. Metabolic characterization of biosurfactants

Microbial production of biosurfactant is growth associated. Growth-associated

biosurfactant production has been reported for the release of biodispersan by A.

calcoaceticus (Desai and Desai, 1993). In addition, biosurfactant production may

possibly occur (or be stimulated) by growing the microbial cells below growth

restrictive conditions. P. aeruginosa shows an over production of rhamnolipid when

the culture reaches the stationary growth phase due to limitation of the nitrogen

Literature Review

source. Additionally, a direct relation exists between biosurfactant production and cell

growth along the fermentation process.

2.7 Improved Strains for Biosurfactant Production

One of the most important factors for economical production of biosurfactant

is use of mutant strain which may be hyper secretary or recombinant which can grow

on cheap raw material supported with efficient recovery process. The Pseudomonas

putida PCL1445 produces two cyclic lipopeptide biosurfactants, putisolvins I and II.

Studies on the regulation of putisolvin production indicate that dnak, together with the

dnaj and grpE heat shock genes were involved in the possible regulation (directly or

indirectly) of putisolvin biosynthesis at the transcription level. A gamma ray induced

mutant viz. B. subtilis AB01335-1M4 and B. subtilis AB02238-1R2 showed 5 and 3

times more Surfactin production, respectively, compared to parent strains when grown

on minimal medium (Bashandy et al., 2005).

Isolation of facultative anaerobic strain which could produce biosurfactant

with crude oil as carbon source and reduce surface tension from 16.36 mN/m to 6.49

mN/m has been reported (Liu and Li, 2005). They isolated the mutant of this strain by

both UV and EMS which could further reduce the surface tension by 32.8%.

2.8 Methods of Detection and Analysis of rhamnolipids

2.8.1. Qualitative Methods

Oil spreading test is a sensitive method for the rapid screening of RL

production by various isolates. In this assay a drop of bacterial supernatant is added

on top of an oil/ water interface (Morikawa et al., 2000). The presence of a surface-

active molecule will cause the oil to be repelled, forming a clearing zone whose

diameter can be correlated with the activity of the tensioactive compounds in the

supernatant.

A more precise approach based on the tensioactive properties of RLs is the

direct measurement of surface tension of culture broths. This method is typically

performed with a du Nouy-type tensiometer, which measures the force required to

pull a thin metal ring out of the surface of the solution. The measurement of the

surface tension after sequential dilution of the solution gives the concentration at

Literature Review

which the surface tension starts to increase and provides the CMC, which is specific

to each surfactant.

Thus, the degree of dilution required to attain the CMC allows for the

quantification of the surfactant in the initial solution (CMD Critical Micelle

Dilution). However, this method suffers from some drawbacks, as it is time-

consuming and not applicable to high-throughput screenings. In addition, as for all the

previous indirect tests based on surface tension, it will be affected by the presence of

tensioactive compounds other than RLs.

2.8.2. Quantitative Methods

2.8.2.1. Thin Layer Chromatography (TLC)

One of the problems of RL quantification is that these compounds are

produced as complex mixtures of congeners, in a medium that may contain many

other interfering compounds. RLs can be somewhat purified by simple extraction

methods, taking advantage of the fact that they are acidic and thus that they will

remain in the aqueous phase in basic medium, while being extractable by relatively

non polar solvents such as ethyl acetate or ethyl ether after acidification of the

aqueous solution. Nevertheless, such crude extracts are seldom pure enough to

gravimetrically quantify only RLs present in the broth. Thus, this requires a

preliminary separation step prior to quantification. TLC is not very sensitive

compared to those mentioned below. As an alternative, direct mass spectrometric

analysis of the eluted TLC plates can be performed using Fast Atom Bombardment

(FAB) to ionize the RLs prior to mass analysis (deKoster et al., 1994). Although this

method provides good structural information, it is not suitable for quantification

purposes.

2.8.2.2. Liquid Chromatography

High Performance Liquid Chromatography (HPLC) is especially well-suited

for RL analysis. It is generally performed using C8 or C18 reverse-phase columns with

a water/acetonitrile gradient. The main problem is the lack of standards to identify

each of the numerous RL congeners present in the culture medium. This can be

overcome by using a mass spectrometer as detector.

Literature Review

2.8.2.3. Liquid Chromatography Coupled to Mass Spectrometry

Direct coupling of reverse phase liquid chromatography to a mass

spectrometer provides the advantages of characterizing a given RL congener by its

retention time along with its mass spectral signature. This is normally done by

splitting the flow coming from the HPLC using a splitter that conveys only a fraction

of the eluent into the mass spectrometer. Electrospray Ionization (ESI), and

sometimes Atmospheric Pressure Chemical Ionization (APCI), has been mostly used

to ionize RLs prior to mass analysis (De´ziel et al., 1999, 2000; Haba et al., 2003a;

Benincasa et al., 2004; Monteiro et al., 2007). In negative ESI, the molecular weight

of the pseudomolecular ion [M-H]_ can be directly obtained. This provides some

information on the nature of the RL congener eluting from the column at that

retention time. In order to improve ionization, ammonium acetate is added to both

solvents of the water/acetonitrile gradient (De´ziel et al., 1999, 2000).

2.8.2.4. Other Spectroscopic Method

Infrared (IR) has been used mostly to quantify complex mixtures of congeners

(Gartshore et al., 2000). This approach is based on the relatively broad IR absorption

bands corresponding to various hydroxyl, ester, and carboxylic groups present in RLs.

This method has been used for the quantification of complex RL mixtures, but it

suffers from interferences by other constituents in the medium and of changes in pH.

Nuclear Magnetic Resonance (NMR) measures the absorption of radio frequencies for

various atoms exposed to a magnetic field. It provides very detailed information on

the chemical environment of atoms (the proton and 13C) within a molecule. This tool

has been used mostly for the structural analysis of purified congeners (Haba et al.,

2003; Monteiro et al., 2007) rather than for quantification of complex RL congener

mixtures.

2.9 Environmental Applications of Biosurfactants

As mentioned before biosurfactant are divided into low-molecular-mass

biosurfactants including glycolipids, phospholipids and lipopeptides and into high-

molecular-mass biosurfactants/bioemulsifiers containing amphipathic

polysaccharides, proteins, lipopolysaccharides, lipoproteins or complex mixtures of

these biopolymers. Low-molecular-mass biosurfactants are efficient in lowering

Literature Review

surface and interfacial tensions, whereas high-molecular-mass biosurfactants are more

effective at stabilizing oil-in-water emulsions (Rosenberg and Ron, 1999; Calvo et al.,

2009).

The biosurfactants accumulate at the interface between two immiscible fluids

or between a fluid and a solid. By reducing surface (liquid-air) and interfacial (liquid-

liquid) tension they reduce the repulsive forces between two dissimilar phases and

allow these two phases to mix and interact more easily (Soberón-Chávez and Maier,

2011) Figure (2.2).

Figure (2.2) Accumulation of biosurfactants at the interface between liquid and

air (Pacwa- et al., 2011).

The most active biosurfactants can lower the surface tension of water from 72

to 30 mN/m and the interfacial tension between water and n-hexadecane from 40 to 1

mN/m (Desai and Banat, 1997; Soberón-Chávez and Maier, 2011). Biosurfactant

activities depend on the concentration of the surface-active compounds until the CMC

is obtained. At concentrations above the CMC, biosurfactant molecules associate to

form micelles, bilayers and vesicles Figure (2.3). Micelle formation enables

biosurfactants to reduce the surface and interfacial tension and increase the solubility

and bioavailability of hydrophobic organic compounds (Whang et al., 2008).

The CMC is commonly used to measure the efficiency of surfactant. Efficient

biosurfactants have a low CMC, which means that less biosurfactant is required to

decrease the surface tension (Desai and Banat, 1997). Micelle formation has a

significant role in microemulsion formation (Nguyen et al., 2008). Microemulsions

are clear and stable liquid mixtures of water and oil domains separated by monolayer

Literature Review

or aggregates of biosurfactants. Microemulsions are formed when one liquid phase is

dispersed as droplets in another liquid phase, for example oil dispersed in water

(direct microemulsion) or water dispersed in oil (reversed microemulsion) (Desai and

Banat, 1997).

Figure (2.3) The relationship between biosurfactant concentration, surfacetension and formation of micelles (Whang et al., 2008).

The biosurfactant effectiveness is determined by measuring its ability to

change surface and interfacial tensions, stabilization of emulsions and by studying its

HLB. The HLB value is a measure to indicate whether a biosurfactant is related to

water-in-oil or oil-in-water emulsion. This factor can be used to determine the suitable

applicability of biosurfactants. Emulsifiers with low HLB are lipophilic and stabilize

water-in-oil emulsification, whereas emulsifiers with high HLB have the opposite

effect and confer better water solubility (Desai and Banat, 1997; Christofi and

Ivshina, 2002).

2.10 Biosurfactants and Hydrocarbons Degradation

The extensive production and use of hydrocarbons has resulted in widespread

environmental contamination by these chemicals. Due to their toxicity, persistent and

negative influence on living organisms it is important to clean-up the polluted sites.

Hydrocarbons, as the hydrophobic organic chemicals, exhibit limited solubility in

groundwater and tend to partition to the soil matrix. This partitioning can account for

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as much as 90 95% or more of the total contaminant mass. As a consequence, the

hydrocarbon contaminants exhibit moderate to poor recovery by physico-chemical

treatments; limited bioavailability to microorganisms; and limited availability to

oxidative and reductive chemicals when applied to in-situ and/or ex-situ applications.

2.10.1. Role of Biosurfactants in Biodegradation Processes

A promising method that can improve bioremediation effectiveness of

hydrocarbon contaminated environments is the use of biosurfactants. They can

enhance hydrocarbon bioremediation by two mechanisms. The first includes the

increase of substrate bioavailability for microorganisms, while the other involves

interaction with the cell surface which increases the hydrophobicity of the surface

allowing hydrophobic substrates to associate more easily with bacterial cells

(Mulligan and Gibbs, 2004).

By reducing surface and interfacial tensions, biosurfactants increase the

surface areas of insoluble compounds leading to increased mobility and

bioavailability of hydrocarbons. In consequence, biosurfactants enhance

biodegradation and removal of hydrocarbons. Addition of biosurfactants can be

expected to enhance hydrocarbon biodegradation by mobilization, solubilization or

emulsification Figure (2.4) (Déziel et al., 1996; Bai et al., 1997; Rahman et al., 2003;

Urum and Pekdemir, 2004; Nguyen et al., 2008; Nievas et al., 2008).

Figure (2.4) Mechanisms of hydrocarbon removal by biosurfactants dependingon their molecular mass and concentration (Rosenberg and Ron, 1999; Urum andPekdemir, 2004).

Literature Review

The mobilization mechanism occurs at concentrations below the biosurfactant

CMC. At such concentrations, biosurfactants reduce the surface and interfacial

tension between air/water and soil/water systems. Due to the reduction of the

interfacial force, contact of biosurfactants with soil/oil system increases the contact

angle and reduces the capillary force holding oil and soil together. In turn, above the

biosurfactant CMC the solubilization process takes place. At these concentrations

biosurfactant molecules associate to form micelles, which dramatically increase the

solubility of oil. The hydrophobic ends of biosurfactant molecules connect together

inside the micelle while the hydrophilic ends are exposed to the aqueous phase on the

exterior. Consequently, the interior of a micelle creates an environment compatible

for hydrophobic organic molecules. The process of incorporation of these molecules

into a micelle is known as solubilization (Urum and Pekdemir, 2004).

Emulsification is a process that forms a liquid, known as an emulsion,

containing very small droplets of fat or oil suspended in a fluid, usually water. The

high molecular weight biosurfactants are efficient emulsifying agents. They are often

applied as an additive to stimulate bioremediation and removal of oil substances from

environments.

2.10.2. Biodegradation Studies

The capability of biosurfactants and biosurfactant-producing bacterial strains

by many authors (Déziel et al., 1996; Rahman et al., 2003; Inakollu et al., 2004).

Obayori et al. (2009) investigated the biodegradative properties of biosurfactant

produced by Pseudomonas sp. LP1 strain on crude oil and diesel. The results obtained

confirmed the ability of strain LP1 to metabolize the hydrocarbon components of

crude and diesel oil. They reported 92.34% degradation of crude oil and 95.29%

removal of diesel oil.

The effective microbiological method in bioremediation of hydrocarbon

polluted sites is the use of biosurfactant producing bacteria without necessarily

characterizing the chemical structure of the surface active compounds. The cell free

culture broth containing the biosurfactants can be applied directly or by diluting it

appropriately to the contaminated site. The other benefit of this approach is that the

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biosurfactants are very stable and effective in the culture medium that was used for

their synthesis.

The usefulness of biosurfactant producing strains in bioremediation of sites

highly contaminated with crude petroleum-oil hydrocarbons was confirmed by Das

and Mukherjee, (2007). The ability of three biosurfactant producing strains: Bacillus

subtilis DM-04, Pseudomonas aeruginosa M and Pseudomonas aeruginosa NM to

remediate petroleum crude-oil contaminated soil samples was investigated by treating

the soil samples with aqueous solutions of biosurfactants obtained from the respective

bacteria strains. Additionally, the tested soil was inoculated with mineral-salts media

containing a specified amount of Bacillus subtilis DM-04 or Pseudomonas aeruginosa

M and NM strains. To determine the extent of biodegradation, the soil-phase total

petroleum hydrocarbons (TPH) concentrations were analyzed after 120 days and

compared to a control where the soil was treated with un-inoculated medium.

Biosurfactants have often been used to enhance bioavailability and

biodegradation of hydrophobic compounds but there is little knowledge available

about the effect of simultaneous emulsifier production on biodegradation of complex

hydrocarbon mixtures. Nievas et al. (2008) studied the biodegradation of a bilge

waste which is a fuel oil-type complex residue produced in normal ship operations.

Bilge waste is a hazardous waste composed of a mixture of sea-water and

hydrocarbon residue, where n-alkanes, resolvent total hydrocarbons and unsolvent

complex mixture are the main constituents. Unsolvent complex mixture principally is

composed by branched and cyclic aliphatic hydrocarbons and aromatic hydrocarbons,

which usually show the greatest resistance to biodegradation. In their studies, they

investigated the biodegradation of an oily bilge wastes by an emulsifier-producing

microbial consortium. As the result for both levels of oily wastes, 136 g/kg of

resolvent hydrocarbons and 406 g/kg of unsolvent mixture, they found that all of the

hydrocarbon types showed an important concentration reduction from their initial

values. They observed that the extent of biodegradation followed the order n-alkanes

> resolved total hydrocarbon > unsolvent complex mixture. An emulsifier-producing

microbial consortium used for biodegradation of bilge wastes showed reduction of n-

alkanes, resolvent hydrocarbons and unsolvent mixture around by 85%, 75% and

58%, respectively.

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2.10.3. Soil Washing Technology

Soil washing technology is characterized by chemico-physical properties of

the biosurfactant and not by their effect on metabolic activities or changes in cell-

surface properties of bacteria (Banat et al., 2010). However, the processes may

enhance the bioavailability for bioremediation. Aqueous solutions of biosurfactants

can be also used to release compounds characterized by low solubility from soil and

other media in process called washing.

Lai et al. (2009) studied the ability of removing TPH from soil by two

biosurfactants: rhamnolipid and surfactin, and two synthetic surfactants: Tween 80

and Triton X-100. The TPH removal efficiency was examined for low TPH-

contaminated (LTC) and high TPH-contaminated (HTC) soils (containing 3000 and

9000 mg/kg dry soil of TPH, respectively) by washing them with (bio) surfactant

solutions. As a result, they observed that addition of 0.2 mass% of rhamnolipid,

surfactin, Triton X-100 and Tween 80 to LTC soil resulted in a TPH removal of 23%,

14%, 6% and 4%, respectively, while for HTC soil a significantly higher TPH

removal efficiency of 63%, 62%, 40% and 35%, respectively, was observed. These

results indicated that among four (bio) surfactants, rhamnolipid and surfactin showed

superior performance on TPH removal, compared to synthetic surfactants. The two

biosurfactants examined in this work have the potential to be used as biostimulation

agents for bioremediation of oil-polluted soils.

2.10.4. Microbial Enhanced Oil Recovery (MEOR)

Biosurfactants can also be involved in microbial enhanced oil recovery

(MEOR). MEOR methods are used to recover oil remaining in reservoirs after

primary (mechanical) and secondary (physical) recovery procedures (Banat et al.,

2000; Sen, 2008). It is an important tertiary process where microorganisms or their

metabolites, including biosurfactants, biopolymers, biomass, acids, solvents, gases

and also enzymes, are used to increase recovery of oil from depleted reservoirs.

Application of biosurfactants in enhanced oil recovery is one of the most

promising advanced methods to recover a significant proportion of the residual oil.

The remaining oil is often located in regions of the reservoir that are difficult to

access and the oil is trapped in the pores by capillary pressure (Sen, 2008).

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Biosurfactants reduce interfacial tension between oil/water and oil/rock. This reduces

the capillary forces preventing oil from moving through rock pores Figure (2.5).

Biosurfactants can also bind tightly to the oil-water interface and form emulsion. This

stabilizes the desorbed oil in water and allows removal of oil along with the injection

water (Suthar et al., 2008).

Figure (2.5) Mechanism of enhanced oil recovery by biosurfactants (Pacwa-

et al., 2011).

2.11 Industrial applications of biosurfactants

Beside applications in MEOR, MS compounds can also be exploited for other

applications in the oil industry. For example, the de-emulsifying properties shown by

some biosurfactant-producing microorganisms may be used to break emulsions which

form at various steps in oil extraction and processing, thus allowing a better recovery

of the product. The surface tension decrease produced by MS can also be used to

separate oil from tank bottom sludge (Singh et al., 2007; Joseph and Joseph 2009;

Perfumo et al., 2010).

Due to their physico-chemical properties, the use of MS compounds has also

been proposed for various industrial applications, as additives in foods, cosmetics and

detergent formulations (Banat et al., 2000). In the food industry, the most useful

property is the ability to form stable emulsions, which improves the texture and

creaminess of dairy products. Biosurfactants are also used to retard staling, solubilise

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flavour oils and improve organoleptic properties in bakery and ice cream formulations

and as fat stabilisers during cooking of fats.

Although the addition of rhamnolipids has been suggested to improve dough

characteristics of bakery products, the use as food ingredients of compounds derived

from an opportunistic pathogen such as P. aeruginosa is not practically feasible.

Instead, it has been suggested to use biosurfactants obtained from yeasts or

Lactobacilli, which are generally recognised as safe and are already involved in

several food-processing technologies (Nitschke and Costa, 2007).

Wetting, dispersing and surface-tension reduction properties, as well as low

toxicity and high biodegradability, suggested the application of biosurfactants,

especially glycolipids, as components of detergent formulations. Low-foaming SLs

from C. bombicola appear suitable due to their high detergency ability, low

cytotoxicity and high biodegradability and general environmentally acceptable

properties (Hirata et al., 2009). Also, cyclic lipopeptide biosurfactants from B. subtilis

improved wash performance by acting additively with other detergent components.

Since they have shown better results at low temperature, such formulations are

promising from an energy-saving point of view, allowing laundering at lower

temperatures (Mukherjee, 2007).

2.12 Biomedical applications

As described in Table (2.3) (Rodrigues et al., 2006a), a broad range of

chemical structures, such as glycolipids, lipopeptides, polysaccharide protein

complexes, phospholipids, fatty acids and neutral lipids, have been attributed to

biosurfactants (Morikawa et al., 1993; Lin, 1996; Angelova and Schmauder, 1999;

Ahimou et al., 2001). Some of these biosurfactants were described for their potential

to act as biologically active compounds and applicability in the medical field.

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Table (2.3) Examples of biosurfactant applications in the medical field(Rodrigues et al., 2006a)

Microorganism Biosurfactanttype

Activity/application Reference(s)

Pseudomonasaeruginosa

Rhamnolipid * antimicrobial activity againstMycobacterium tuberculosis * anti-adhesive activity againstseveral bacterial and yeaststrains isolated from voiceprostheses

(Gerard et al., 1997;Lang and Wullbrandt,1999; Maier andSoberón-Chávez,2000; Rodrigues etal., 2006b )

Bacillus subtilis Surfactin * antimicrobial and antifungalactivities* inhibition of fibrin clotformation* haemolysis and formation ofion channels in lipidmembranes* antitumour activity against

cells* antiviral activity againsthuman immunodeficiencyvirus 1 (HIV-1)

(Bernheimer andAvigad, 1970;Kameda et al., 1974;Vollenbroich et al., 1997a; Vollenbroichet al., 1997b;Sheppard et al., 1991;Itokawa et al., 1994)

Bacillus pumilus pumilacidin(surfactinanalogue)

* antiviral activity againstherpes simplex virus 1 (HSV-1)* inhibitory activity againstH+, K+-ATPase and protectionagainst gastric ulcers in vivo

(Naruse et al., 1990)

Bacillus subtilis Iturin * antimicrobial activity andantifungal activity againstprofound mycosis* effect on the morphology andmembrane structure of yeastcells* increase in the electricalconductance of biomolecularlipid membranes* non-toxic and non-pyrogenicimmunological adjuvant

(Besson et al., 1976;Thimon et al., 1995;Mittenbuhler et al.,1997; Tanaka et al.,1997; Ahimou et al.,2001)

Bacilluslicheniformis

Lichenysin * antibacterial activity * chelating properties thatmight explain the membrane-disrupting effect oflipopeptides

(Jenny et al., 1991;Lin et al., 1994;Yakimov et al., 1995;Grangemard et al.,2001)

Candida antartica mannosylerythritollipids

* antimicrobial, immunologicaland neurological properties* induction of celldifferentiation in the humanpromyelocytic leukemia cell

(Kitamoto et al.,1993; Isoda et al.,1997; Isoda et al.,1999; Zhao et al.,1999; Shibahara et al.,

Literature Review

line HL60* induction of neuronaldifferentiation in PC12 cells

2000; Zhao et al.,2000; Wakamatsu etal., 2001)

Rhodococcuserythropolis

treahalose lipid * antiviral activity against HSVand influenza virus

(Uchida et al., 1989a;Uchida et al., 1989b)

Streptococcusthermophilus

Glycolipid * anti-adhesive activity againstseveral bacterial and yeaststrains isolated from voiceprostheses

(Busscher et al., 1994;Busscher et al., 1997;Busscher et al., 1999;Rodrigues et al.,2004; Rodrigues etal., 2006c)

Streptococcus mitis not identified * anti-adhesive activity againstStreptococcus mutans

(Pratt-Terpstra et al.,1989; Van Hoogmoedet al., 2000)

Lactobacillus Surlactin * anti-adhesive activity againstseveral pathogens includingenteric bacteria

(Reid et al., 1984;Velraeds et al., 1996a; Velraeds et al.,1996b ; Velraeds etal., 1997; Reid et al.,1999)

Lactococcus lactis not identified * anti-adhesive activity againstseveral bacterial and yeaststrains isolated from voiceprostheses

(Rodrigues et al.,2004a; Rodrigues etal., 2004b)

2.12.1. Antimicrobial activity of biosurfactants

The high demand for new antimicrobial agents following increased resistance

shown by pathogenic microorganisms against existing antimicrobial drugs has drawn

have been reported to be suitable alternatives to synthetic medicines and antimicrobial

agents and may therefore be used as effective and safe therapeutic agents (Banat et

al., 2000; Cameotra and Makkar, 2004; Singh and Cameotra, 2004).

Lipopeptides form the most widely reported class of biosurfactants with

antimicrobial activity. Surfactin, produced by B. subtilis, is the best-known

lipopeptide (Arima et al., 1968). Other antimicrobial lipopeptides include fengycin,

iturin, bacillomycins and mycosubtilins produced by B. subtilis (Vater et al., 2002).

Lichenysin, pumilacidin and polymyxin B (Naruse et al., 1990; Yakimov et al., 1995;

Grangemard et al., 2001; Landman et al., 2008) are other antimicrobial lipopeptides

produced by B. licheniformis, Bacillus pumilus and Bacillus polymyxa, respectively.

The production of antimicrobial lipopeptides by Bacillus probiotic products is one of

the main mechanisms by which they inhibit the growth of pathogenic microorganisms

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in the gastrointestinal tract (Hong et al., 2005). Other reported biosurfactants having

antimicrobial activity are daptomycin, a cyclic lipopeptide from Streptomyces

roseosporus (Baltz et al., 2005), viscosin, a cyclic lipopeptide from Pseudomonas

(Neu et al., 1990; Saini et al., 2008), rhamnolipids produced by P. aeruginosa

(Abalos et al., 2001; Benincasa et al., 2004) and sophorolipids produced by C.

bombicola (Kim et al., 2002; Van Bogaert et al., 2007). Mannosylerythritol lipids

(MEL-A and MEL-B) produced by Candida antarctica strains have also been

reported to exhibit antimicrobial action against Gram-positive bacteria (Kitamoto et

al., 1993).

A rhamnolipid surfactant produced from soybean oil waste had antimicrobial

activity against several bacteria and fungi, namely Bacillus cereus, S. aureus,

Micrococcus luteus, Mucor miehei and Neurospora crassa (Nitschke et al., 2009).

Sophorolipids are also claimed to have activity against human

immunodeficiency virus (Shah et al., 2005). Similarly, a rhamonolipid and its

complex with alginate, both produced by a Pseudomonas sp. strain, showed

significant antiviral activity against herpes simplex virus types 1 and 2 (Remichkova

et al., 2008). The suppressive effect of the compounds on herpes simplex virus

replication was dose-dependent and occurred at concentrations lower than the CMC.

2.12.2. Biomedical and therapeutic applications of biosurfactants

Although there is an increasing potential for the application of biosurfactants

in the biomedical field, some of these molecules may pose a risk for humans. For

instance, P. aeruginosa is a bacterium responsible for severe nosocomial infections,

life threatening infections in immunocompromised persons and chronic infections in

cystic fibrosis patients; thus, rhamnolipids have to be well investigated prior to such

uses.

The virulence of a P. aeruginosa strain depends on a large number of cell-

associated and extracellular factors (Ishigami and Suzuki, 1997; Van Delden and

Iglewski, 1998; Ramisse et al., 2000). Cell-to-cell signalling systems control the

expression and allow a coordinated, cell-density-dependent production of many

extracellular virulence factors. The possible role of cell-to-cell signalling in the

Literature Review

pathogenesis of P. aeruginosa infections and a rationale for targeting cell-to cell

signalling systems in the development of new therapeutic approaches were discussed

by Van Delden and Iglewski, (1998). Synthesis of rhamnolipids is regulated by a very

complex genetic regulatory system that also controls different P. aeruginosa

virulence- associated traits (Maier and Soberón-Chávez, 2000).

The cosmetic and healthcare industries use large amounts of surfactants for a

wide variety of products, including insect repellents, antacids, acne pads, contact lens

solutions, hair colour and care products, deodorants, nail care products, lipstick, eye

shadow, mascara, toothpaste, denture cleaners, lubricated condoms, baby products,

foot care products, antiseptics and shaving and depilatory products (Kosaric, 1992).

Biosurfactants are known to have advantages over synthetic surfactants such

as low irritancy or anti-irritating effects and compatibility with skin. Rhamnolipids in

particular are being used as cosmetic additives and have been patented to make some

liposomes and emulsions, (Ishigami and Suzuki, 1997; Ramisse et al., 2000) both of

which are important in the cosmetic industry.

Materials and Methods

CHAPTER III

MATERIALS AND METHODS

3.1. Microorganisms and their isolation

The Microorganism used in all experiments was isolated by using Cetrimide

agar (Difco, U.S.A) from Compost pile waste collected from El-Sharkia region in

Delta, Egypt obtained in September 2010.

Cetrimide agar was melted and poured into separate sterile Petri dishes and

allowed to solidify. Thin layer of the compost pile was spread on the surface of

Cetrimide agar and incubated at 37oC for 24 hours and observed for signs of growth

and colony appearance.

The green colonies were picked with sterile inoculating wire and streaked on

the surface of MacConkey agar plates, labeled and incubated at 37oC for 24 hours and

observed for signs of growth and colony appearance.

White and golden to brown colonies on the MacConkey Agar plates were also

picked with sterile inoculating wire and streaked on the surface of blood agar plates,

labeled and incubated at 37oC for 24 hours.

The yellowish-green colonies observed were streaked onto nutrient agar slants

for various sub-cultures and incubated at 37oC for 24 hours and then kept (in the

refrigerator at -20oC) for further identification.

3.2. Identification of micro-organisms

3.2.1. Gram Stain

A clean grease free slide was obtained by cleaning with hot water and soap.

After which it was rinsed and the excess water blotted out with a lint-free cloth. A

drop of distilled water was placed on the slide and with the help of an inoculating

needle; the isolate picked from the surface of the agar plate was gently and thoroughly

emulsified in the drop of water and thinly spread on the glass slide. The smear was

dried in air and passed through a Bunsen flame to fix. It was then flooded with

Materials and Methods

ammonium oxalate crystal violet solution for about one minute and rinsed off with

washed off. The excess water was blotted off with bibulous paper and 95% ethyl

alcohol was applied and allowed to drain off until no more colour was seen leaving

the smear. It was again washed with water and Safranin was added and left for about

30 seconds. The contrast stain was then washed off and the smear dried in air (Harley

and Prescott, 1990). The slides were examined with the aid of a light microscope

(Olympus, Tokyo, Japan).

3.2.2. Biochemical Tests

Various biochemical tests were performed on the isolates to confirm their

identities. These included motility, catalase, oxidase, growth at 42oC and on Triple

Sugar Iron Agar.

3.2.2.1 Motility test

The suspected Pseudomonas isolates were inoculated into 10 ml nutrient

broths and incubated at 37 oC for 18-24 hours. Small drop of liquid bacterial culture

was placed in the centre of a cover slip and a small drop of water was placed at each

corner of the cover slip then a slide was inverted with a central depression over the

cover slip. The cover slip was stacked to the slide and when the slide was inverted the

drop of bacterial culture was suspended in the well (Health Protection Agency,

2010a). Motile organisms were examined using the light microscope (Olympus,

Tokyo, Japan).

3.2.2.2. Catalase test

A loopful of the isolate was streaked on nutrient agar and incubated at 37°C

for 24 hours. 0.2 ml of hydrogen peroxide solution was placed in a test tube. The

culture of the isolate was rubbed on the inside wall of the bottle above the surface of

the hydrogen peroxide solution and hydrogen peroxide solution was allowed to cover

the colony. A positive reaction was indicated by the appearance of vigorous bubbling

occurring within 10 seconds (Health Protection Agency, 2010b).

Materials and Methods

3.2.2.3. Oxidase production test

The oxidase reagent was prepared by dissolving 0.1 g of tetramethyl-p-

phenylenediamine in 10 ml of sterile distilled water. A loopful of the isolate was

streaked on nutrient agar and incubated at 37°C for 24 hours. Clean Whatman No.2

filter paper was placed in Petri dish and three drops of the freshly prepared oxidase

reagent added to it. The culture of the isolate was smeared across the impregnated

paper with a platinum loop (Harley and Prescott, 1990). A positive reaction was

indicated by the appearance of a dark purple colour on the paper within 10 seconds.

3.2.2.4. Growth at 42oC

The suspected Pseudomonas isolates were inoculated into 10 ml nutrient

broths and incubated at 42oC for 24 hours and observed for signs of growth (Hsueh et

al., 1998).

3.2.2.5. Reaction in Triple Sugar Iron Agar

The isolates grown on nutrient agar slants were picked with a sterile platinum

wire and then inoculated into triple sugar iron agar. The tubes were capped and

incubated at 37°C for 24 hours. A control of TSI Agar which was not inoculated was

also incubated alongside the tests (Mahon et al., 2007). After incubation, the tubes

were examined for the presence of sugar fermentation, gas and hydrogen sulfide

production (Harley and Prescott, 1990). A yellow butt and yellow slant due to the

fermentation of lactose, sucrose and glucose was recorded as a positive test.

3.2.3. API 20E Identification for selected isolate

The API20E strip (this strip is the same one used for the identification of

Enterobacteriaceae) contains 20 microtubes with substrates for the following 23 tests:

0-nitrophenyl- -D-galactosidase (ONPG); arginine dihydrolase; lysine and ornithine

decarboxylase; citrate utilization; hydrogen sulfide; urease; tryptophan deaminase;

indole; Voges-Proskauer (acetoin); gelatin liquefaction; fermentation of the

carbohydrates glucose, mannitol, inositol, sorbitol, rhamnose, sucrose, melibiose,

amygdalin, and arabinose; nitrate reduction and nitrogen gas production, tested in the

glucose microtube; and catalase production, in any other carbohydrate microtube. A

Materials and Methods

complete description of the strip is given in other reports (Washington et al., 1971;

Smith et al., 1972; Brooks et al., 1974; Nord et al., 1974).

Twenty-two of the initial tests were checked and recorded after 24 h of

incubation and again at 48 h. At this point, reagents were added for the five remaining

tests (tryptophan deaminase, indole, Voges-Proskauer, nitrate reduction, and nitrogen

gas production), and those results were recorded. The Analytical Profile Index and (if

necessary) the computer service were consulted for identification of the isolates.

When indicated, supplemental tests recommended by the manufacturer for

identification were done using conventional media. When sufficient reactions were

clear at 24 h, identification was determined on those readings.

3.2.4. 16S rRNA gene sequence analysis for selected isolate

For molecular identification, the selected bacterial isolate was referred for 16S

rRNA gene sequence analysis. DNA extraction was performed from 2 ml bacterial

cultures collected at the mid-exponential growth phase using Roche Kit (Germany)

according to the manufacturer's instructions and run in triplicate through polymerase

chain reaction (PCR). Three sets of primers are listed in Table (3.1).

V3 and V6 primers were used to amplify the V3 and V6; the most two

relevant hyper variable regions of the bacterial 16S rRNA gene, giving a product of

203 and 124 bp, respectively (Chakravorty et al. 2007). The reaction was carried out

Pfu DNA polymerase (Fermentas, St. Leon-Rot, Germany)

and 1 ng of template DNA. PCR amplification was performed as following: initial

denaturation at 95°C for 5 min, followed by 25 cycles each of 94°C for 1 min, 55°C

of annealing for 45 s, and a 45 s extension at 72°C. The 203 and 124 bp PCR products

amplified from the bacterial isolate which appeared as a single band were purified

using a High Pure PCR product purification kit (Roche Applied Science, Germany)

and sequenced on an ABI Prism 377 automatic sequencer (Applied Biosystems, CA,

USA). Sequence homologies were examined using BLAST version 2.2.12 of the

National Center for Biotechnology Information (Altschul et al. 1990). Multiple

sequence alignments were carried out using ClastalW and a consensus neighbor-

joining tree was constructed using Molecular Evolutionary Genetics Analysis

Materials and Methods

(MEGA) software (version 4.0) (Tamura et al. 2007). Finally potent biosurfactant

producing isolates were maintained on nutrient agar slants for further studies.

Table (3.1) List of primers used in this study

Name Sequence Size (bp) Reference

27 2

27f

-AGAGTTTGATCCTGGCTCAG- (Lane,

1991)

1492

1492r

-TACGGTTACCTTGTTACGACTT-

V3 VV3F

-CCAGACTCCTACGGGAGGCAG- 203 (Chakravorty

et al. 2007).

VV3R

-CGTATTACCGCGGCTGCTG-

V6 VV6F

-TCGATGCAACGCGAAGAA- 124 (Chakravorty

et al. 2007).

VV6R

-ACATTTCACaACACGAGCTGACGA-

3.3. Production of biosurfactant

3.3.1. Preparation of Culture medium

The strain from nutrient agar slants was streaked on Cetrimide agar plate,

which was then incubated at 37 C for 14 16 h. After that, a single colony was taken

from the plate and transferred into 50 ml liquid broth (LB) to prepare the seed culture.

The cultivation condition for the seed culture was 37 C, 200 rpm, and 14 16 h of

incubation time.

3.3.2. Fermentation medium and condition

For liquid fermentation, the seed culture (5% inoculum) was inoculated into a

500-ml flask containing 150 ml mineral salts (MS) medium consisting of (g/l):

NH4NO3, 4; KH2PO4, 4.08; Na2HPO4, 5.68; CaCl2, 7.77 · 10_4; MgSO4 · 7H2O, 0.2;

sodium EDTA, 1.49 · 10_3; FeSO4 · 7H2O, 5.56 · 10_4 (Wei and Chu, 1998). In

general, the MS medium was amended with 40 g/l glucose as a sole carbon substrate

(designated as GMS medium) (Wei and Chu, 1998; Wei et al., 2005). The culture

temperature and agitation rate was 37 C and 200 rpm, respectively in orbital incubator

Materials and Methods

SI50 (Stuart Scientific, UK) for 7 days. The pH of the medium was initially adjusted

to 6.8 by 1.0 M HCl.

3.4. Dried weight cell determination

After cultivation for 7 days, the collected fermentation broth was first

centrifuged at 9000g for 15 min to remove bacterial cells. 50 ml of the filtrate was

taken to detect surface tension, emulsification index, and oil displacement test which

will be described later. The removed bacterial cells were collected and placed in an

oven at 110 oC for 12 h to obtain a dried weight reported in terms of g/l, which is used

to express the microbial concentration.

3.5. Extraction of the produced biosurfactants

The pH of the remaining supernatant was adjusted to pH 2.0 with 1N HCl to

precipitate rhamnolipid. The precipitate was harvested by centrifugation (9000g, 20

min) and was then extracted three times with ethyl acetate at room temperature. The

organic phase was collected and the solvent was removed in a rotary evaporator,

allowing the yield of viscous honey-colored rhamnolipid product (Rotary vacuum

evaporator RE 10, Bibby Co., UK) (Chen et al., 2005; Wei et al., 2005).

3.6. Optimization of Biosurfactant Production

Type, quality and quantity of biosurfactant production is dependent on the

cultural conditions i.e. pH, temperature, agitation, aeration, dilution rate, the

concentration of metal ions, the nature of the carbon and nitrogen sources.

3.6.1. Optimum Carbon Substrate

For the experiments exploring the effect of carbon substrates on rhamnolipid

production, the carbon source in GMS medium (i.e., glucose) was replaced by

glycerol (40 g/l), sucrose (40 g/l), hexane (40 g/l), olive oil (80 g/l) and oleic acid (80

g/l) (Wu et al. 2008). Growth studies were done in 500-ml flasks containing 150 ml

medium at 37 C with pH 6.8 and shaking at 200 rpm for 7 days in orbital incubator

SI50. Experiment were done in duplicate and results reported as the average.

Biosurfactant production was monitored by measuring the weight and surface tension

of the partially purified biosurfactant.

Materials and Methods

3.6.2. Optimum Nitrogen Source

The nitrogen source in GMS medium (i.e. NH4NO3) was also replaced by

NH4Cl (50 mM), NaNO3 (50 mM), urea (50 mM), and yeast extract (10 g/l) to

investigate the effect of nitrogen source on rhamnolipid production. The concentration

of carbon and nitrogen source was identical to GMS medium, while that of oil and

yeast extract was determined according to the information from literature and

preliminary tests (Wu et al. 2008). The media was incubated at 37 C with pH 6.8 and

shaking at 200 rpm for 7 days in orbital incubator SI50.

3.6.3. Optimum pH

The optimum pH of MS medium for growth and biosurfactant production of

the tested bacterial isolates was investigated. The pHs tested were 4, 5, 6, 7, and 8.

The pH of the medium was adjusted using a solution of HCl or NaOH (1 M), with the

aid of a Sentron 2001 pH meter, and the inoculated medium was amended with

optimum carbon substrate and optimum nitrogen source.

3.6.4. Optimum Temperature

The growth and biosurfactant production of the isolated microorganisms were

monitored at a range of temperatures to determine the optimum one for each isolate.

The tested temperatures were 25, 30, 35, 37, 40 and 45oC. The inoculated medium

was adjusted to the optimum pH and supplied with the optimum carbon substrate and

optimum nitrogen source.

3.6.5. Optimum Agitation Rate

The growth and biosurfactant production of the isolated microorganisms were

monitored at a range of rpms to determine the optimum one for each isolate. The

tested rpms were 100, 150, 200, and 250 rpm. The inoculated medium was adjusted to

the optimum pH and supplied with the optimum carbon substrate and optimum

nitrogen source and adjusted at optimum temperature.

3.7. Purification of biosurfactant

The extracted viscous honey-colored rhamnolipid product was collected for

further purification using the chromatographic procedures. A 1.04 g sample of crude

Materials and Methods

biosurfactant was dissolved in chloroform, mixed with a small amount of silica gel 60

and dried with a rotary evaporator. The column (1.5cm×35 cm) was packed with

silica gel in hexane and not allowed to dry. The sample was applied on the surface of

the column and eluted with hexane. After that, it was eluted with solvents of gradually

increasing polarity: hexane > acetone > chloroform > chloroform:methanol (2:1, v/v)

>methanol (Darvishi et al., 2011).

Each fraction was evaporated with a rotary evaporator and its oil displacing

area was measured. The fractions that demonstrated the oil displacement test were

further separated by TLC using aluminum sheets of silica gel 60 F254 plates and a

chloroform:methanol:20% aqueous acetic acid (65:15:2) solvent system (Koch et al.,

1991). Rhamnolipids can be visualized using the orcinol test (Koch et al., 1991) or

Molisch test, with reagents specific for sugars or fatty acids or with reagents that are

used to reveal (Mechaly et

al., 1997).

3.8. Structural characterization of biosurfactant

The active fraction was analyzed by Fourier transform infrared spectroscopy

(FTIR), Nuclear magnetic resonance (1HNMR) and Electrospray ionisation mass

spectrometry (ESI MS).

3.8.1. Fourier transform infrared spectroscopy (FTIR)

FT-IR spectra of the dried biosurfactants were recorded on a Bruker 113V FT-

IR spectrometer equipped with a mercury cadmium telluride (MCT) detector cooled

with liquid N2. About 2mg of dried biomaterial was milled with 200mg of KBr to

form a very fine powder. This powder was then compressed into a thin pellet which

could be analyzed by FT-IR spectra measurement in wave number range of 4000 400

Cm 1 (Lotfabad et al., 2009). The analysis of IR spectra was carried out by using

OPUS 3.1 (Bruker Optics) software.

3.8.2. Nuclear magnetic resonance analysis

The 1H NMR spectra of fraction obtained from the fractionation step were

achieved from an FT NMR 500 mHz spectrometer (JEOL, JNM-A500) using

Materials and Methods

deuterated methanol as a solvent. The sample for analysis should be pure as possible

and free from insoluble and paramagnetic material (Pornsunthorntawee et al., 2008).

3.8.3. Electrospray ionisation mass spectrometry (ESI MS)

The analyses were performed with a triple Quadrupole Quattro LC

(Micromass, Manchester, UK) in negative-ion mode. 2 mg of rhamnolipid fraction

was dissolved in 1ml chloroform methanol (1:1, v/v), and aliquots of 0.1ml were

removed and diluted in 1.9 ml of acetonitrile water (7:3, v/v), which was introduced

by direct infusion with a syringe pump at a flow rate of 10µL min 1. ESI tandem mass

spectra were acquired by mass-selecting the target ion using a quadrupole mass

analyzer. The conditions of the analyses were: F1 capillary 2.03 kV, cone 15V and

collision energy 18V for collision induced dissociation (CID); F2 capillary 2.03 kV,

cone 27V and collision energy 45V for CID (Monteiro et al., 2007).

3.9. Physicochemical characterization

3.9.1. Oil displacement test

The oil displacement test is a method used to determine the surface activity by

measuring the diameter of the clear zone, which occurs after dropping a surfactant-

containing solution on a thin layer of oil on water. The binomial diameter allows an

evaluation of the surface tension reduction efficiency of a given biosurfactant

(Rodrigues et al., 2006d). The oil displacement test was done by adding 40 ml of

distilled water to a Petri dish with a diameter of 15 cm. After that, 15µl of crude oil

was dropped to form a thin oil layer on the surface of the water, and then 10µl of a

test solution was dropped onto the surface of oil. The test was conducted at room

temperature (25 27 C). The maximum diameter of the clear zone was observed under

light and measured. The larger the diameter of the clear zone, the higher the surface

activity of the test solution.

3.9.2. Surface tension measurement

The surface tension of the aqueous solution at different surfactant

concentrations was measured by Du-Nouy ring method with a Kruss Tensiometer

(Kruss, Hamburg, Germany). The surface tension measurement was carried out at 25

± 1 C after dipping the platinum ring in the solution for a while in order to attain

Materials and Methods

equilibrium conditions. The measurement was repeated three times and an average

value was obtained. CMC was then determined from the break point of the surface

tension versus its log of bulk concentration curve. For the calibration of the

instrument, the surface tension of the pure water was measured before each set of

experiments (Pornsunthorntawee et al., 2008).

3.9.3. Measurement of critical micelle concentration (CMC)

CMC is an important parameter during the evaluation of activity of

biosurfactant. The surface tension of surfactant reaches the lowest at its CMC. Above

this concentration, no further effect can be observed on the surface activity.

Measuring surface tension of serially diluted biosurfactant solution, the CMC was

determined by plotting the surface tension versus concentration of biosurfactant in the

solution. The determination of CMC was performed by several dilutions of crude

biosurfactant. These experiments were conducted in three independent replicates and

the results presented were the average data.

3.9.4. Determination of emulsification index

A mixture of 2 ml supernatant and 3 ml kerosene (or diesel) was vortically

stirred for 2 min and the height of emulsion layer was measured after 24 h to

determine the emulsification index (Cooper and Goldenberg, 1987; Wu et al., 2008).

The equation used to determine the emulsification index (E24 (%)) is as follows:

Experimental Results

CHAPTER IV

EXPERIMENTAL RESULTS

4.1. Isolation and identifiction of selected Biosurfactant-producing

bacterial strain

A total of 6 (non-lactose fermenter) isolates recovered on MacConkey agar

were suspected as P. aeruginosa and showed green fluorescence colour on Cetrimide

-hemolytic activity lysis and complete digestion of red

blood cell when grown on blood agar Figure (4.1).

The bacterial isolate was examined based on its morphological and

biochemical characteristics. As shown in Figure (4.1), the isolated strain produced a

diffusible green colored fluorescent pigment with excessive amounts of foam when it

grown on glucose-containing MS medium. The bacterial isolate was a non-spore

forming-gram negative, rod shape, motile, the catalase and oxidase tests were

positive, growth on triple sugar iron was neutral bottom and neutral slant, no

production of hydrogen sulfide (H2S) when grown on MacConkey agar, it could not

use lactose as a carbon source and can grow at 42oC.

Identification of selected bacterial isolate was done using morphological and

biochemical methods with API 20E kit (Biomérieux, Mercy, France). Results of

various biochemical tests and general characteristics of selected isolate are presented

in Table (4.1). Preliminary results indicated that strain was a member of the genus

Pseudomonas.

Further identification was performed based on the 16S rRNA gene sequence

analysis. Examination with three sets primers for almost fully and partial 16S rRNA

sequence alignment revealed that strain TMN was closely related to the species in

genus Pseudomonas. Strain TMN also exhibited the highest similarity (98%) to P.

aeruginosa so, strain TMN was tentatively classified as P. aeruginosa TMN Figure

(4.2).

Experimental Results

Table (4.1) Biochemical characteristics of the 6 isolates of gram negative bacteriagrown on the API 20E strip and identified as Pseudomonas aeruginosa

Test Result- galactosidase

arginine dihydrolaselysine decarboxylaseornithine decarboxylasecitrate utilizationhydrogen sulfide productionurease productiontryptophan deaminaseindole productionVoges-Proskauer (acetoin)gelatin liquefactionAcid from:glucose mannitolinositol sorbitol rhamnose sucrosemelibioseamygdalinarabinoseOxidase production

---++-----+

+--------+

Experimental Results

A: Pseudomonas aeruginosa on B: gram stain for Pseudomonas blood agar aeruginosa

C: Pseudomonas aeruginosa on D: Pseudomonas aeruginosa on MacConkey agar Cetrimide agar

E: Pseudomonas aeruginosa on glucose-containing MS medium

Figure (4.1) Growth of Pseudomonas aeruginosa on blood agar, MacConkey

agar, Cetrimide agar, glucose-containing MS medium and its gram stain.

Experimental Results

Figure (4.2) The neighbor-joining tree based on the 16S rRNA sequence,

demonstrating the phylogenetic position of strain TMN within Pseudomonas

species.

Experimental Results

4.2. Optimization of Biosurfactant Production

Factors affecting on the production of biosurfactant from Pseudomonas

aeruginosa were evaluated to detect the optimum media used to produce biosurfactant

based on the amount of biosurfactant and its surface tension.

4.2.1. Effects of carbon sources on rhamnolipid production

Pseudomonas aeruginosa was grown on different carbon sources to optimize

the media used for biosurfactant production. The carbon sources used in this

experiment include glucose, glycerol, sucrose, hexane, olive oil, and oleic acid (80

g/l).

The biosurfactant production was monitored by measuring the reduction in

surface tension, dry cell weight and the weight of crude extract. Figure (4.3) indicates

that the properties and amount of biosurfactant produced depend on the type of

substrate.

Glucose was the best carbon substrate for rhamnolipid production based on the

quantity of rhamnolipid produced (0.3+0.01 g/l) then glycerol produced 0.25+0.02 g/l

of rhamnolipid. Meanwhile, the dry cell weight for glucose and glycerol was 2.9+

0.02 and 1.35+ 0.01 g/l, respectively. Olive oil, oleic acid, hexane and sucrose

produced 0.19+0.03, 0.13+0.01, 0.07+0.02 and 0.02+0.01 g/l rhamnolipid

respectively. Dry cell weight for olive oil, oleic acid, hexane and sucrose were 5.07+

0.04, 5.0+ 0.06, 0.2+ 0.01 and 0.91+ 0.03 g/l respectively.

On the other hand the surface tension measurement for the produced

biosurfactants from different carbon sources indicate that glucose was the best carbon

source which had a surface tension of 34 mN/m and achieved an emulsification index

of over 46% for kerosene.

4.2.2. Effects of nitrogen sources on rhamnolipid production

Medium constituents other than carbon sources also affect the production of

rhamnolipids. Nitrogen source is one of the most important factors affecting

rhamnolipid production. Five different organic and inorganic nitrogen sources used in

this study is shown in Figure (4.4).

Experimental Results

It was found that NaNO3 was the most efficient nitrogen source for P.

aeruginosa TMN and ammonium nitrate follows it to produce rhamnolipid, giving a

high rhamnolipid yield of 0.34+0.01 and 0.3+0.01 g/l, respectively. Moreover, using

urea and yeast extract as the organic nitrogen sources led to a rhamnolipid yield of

0.3+0.01 and 0.25+0.01 g/l, respectively. Surface tension measurement for the

produced biosurfactants from different nitrogen sources indicate that NaNO3 was the

best nitrogen source which had a surface tension of 34 mN/m and achieved an

emulsification index of over 46% for kerosene. Hence, in the following experiments,

NaNO3 was chosen as the nitrogen source and glucose as carbon source to achieve a

higher rhamnolipid yield.

4.2.3. Effect of pH on rhamnolipid production

It was found that the yield of biosurfactant increased when pH varied from 4

to 8 and reached the highest at pH 7 (0.36+ 0.03 g/l); however, when pH reached 8,

the biosurfactant yield declined and came to its lowest point (0.12+ 0.02 g/l). Thus, it

can be inferred that the TMN strain excreted biosurfactant more effectively under

neutral conditions as shown in Figure (4.5).

4.2.4. Effect of temperature on rhamnolipid production

TMN strain was grown in GMS medium at 25 45 C to explore the influence

of culture temperature on rhamnolipid production. As indicated in Figure (4.6),

rhamnolipid production increased with temperature from 25 (0.15+ 0.01 g/l) to 37 C

(0.34+ 0.07 g/l), and decreased slightly when temperature was further increased to 45

C (0.19+ 0.05 g/l). P. aeruginosa was unable to grow at 47 C, leading to negligible

rhamnolipid production at that temperature. These results suggest that the optimal

temperature for rhamnolipid production with TMN was in the range of 37 40 C.

Experimental Results

Figure (4.3) Effect of different carbon sources on the yield and surface tension.

Figure (4.4) Effect of different nitrogen sources on the yield and surface tension.

0

10

20

30

40

50

60

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

Glucose Glycerol Sucrose Hexane Oleic acid Olive oil

Carbon Source

Surf

ace

Tens

ion

(dyn

e/cm

)

Rha

mno

lipid

Yiel

d (g

/L)

Rhamnolipid Yield (g/L) Surface Tension (dyne/cm)

0

5

10

15

20

25

30

35

40

45

50

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

0.4

NH4Cl (50mM)

NaNO3 (50mM)

Urea (50 mM) Yeast extract(1%)

NH4NO3 (50mM)

Nitrogen Source

Surf

ace

Tens

ion

(dyn

e/cm

)

Rha

mno

lipid

Yiel

d (g

/L)

Rhamnolipid Yield (g/L)Surface Tension (dyne/cm)

Experimental Results

Figure (4.5) Effect of different pH on rhamnolipid yield and dry cell weight.

Figure (4.6) Effect of different temperatures on rhamnolipid yield and dry cell

weight.

0

0.5

1

1.5

2

2.5

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

0.4

0.45

4 5 6 7 8

pH

Rha

mno

lipid

Yiel

d (g

/L)

Dry

Cel

lWei

ght (

g/L)

Rhamnolipid Yield (g/L) Dry cell weight (g/L)

0

0.5

1

1.5

2

2.5

3

3.5

4

4.5

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

0.4

0.45

25 30 35 "37" 40 45

Temperature (oC)

Rha

mno

lipid

Yiel

d (g

/L)

Dry

Cel

lWei

ght (

g/L)

Rhamnolipid Yield (g/L) Dry cell weight (g/L)

Experimental Results

4.2.5. Effect of agitation rate on rhamnolipid production

Agitation rate also affect rhamnolipid production. Results from batch

fermentation under different agitation rates (100 250 rpm) show that as the agitation

rate increased from 100 to 200 rpm, rhamnolipid production increased and dry cell

weight was also improved from 0.14+ 0.01 and 0.94+ 0.05 to 0.3+ 0.01 and 1.8+ 0.03

g/l respectively as shown in Figure (4.7). The rhamnolipid production and dry cell

weight are slightly decreased to 0.15+ 0.02 and 1.59+ 0.06 g/l respectively at 250

rpm. As a result, an agitation rate of 200 rpm was used in the rest of experiments.

Figure (4.7) Effect of different agitation rates on rhamnolipid yield and dry cell

weight.

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

2

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

100 150 200 250

Agitation Rate (rpm)

Rha

mno

lipid

Yiel

d (g

/L)

Dry

Cel

lWei

ght (

g/L)

Rhamnolipid Yield (g/L) Dry cell weight (g/L)

Experimental Results

4.3. Purification of biosurfactant

The purification steps of biosurfactant produced by Pseudomonas aeruginosa

were carried out as mentioned in Materials and Methods. The purification profile of

biosurfactant was presented in Table (4.2). The fractions were collected and classified

into 5 parts by solvents of gradually increasing polarity: hexane > acetone >

chloroform > chloroform: methanol (2:1, v/v) >methanol. Each fraction was dried and

weighed. The fractions were varied in colour, where the fraction 1 was light yellow in

colour, fraction 2 was light brown, fraction 3 was brown, fraction 4 was dark brown

and fraction 5 was light brown in colour. The fractions were weighted, where

fractions 1,2,3,4 and 5 were 0.03, 0.64, 0.02, 0.14 and 0.05 g respectively.

Oil displacement test was carried out for each fraction and represented in

Table (4.2). Fraction 4 show best oil displacement result where the diameter of clear

zone on the oil surface was 10.5 cm, while fractions 1, 2, 3 and 5 the diameter of clear

zone on the oil surface were ve, 3, 3.5 and 4 respectively. From the previous results,

fraction 4 which have dark brown colour and diameter of clear zone on the oil surface

10.5 cm at concentration of 0.14 g/l was the best biosurfactant. This purified

biosurfactant was characterized to elucidate the chemical structure for it.

4.4. Structural characterization

4.4.1. Thin Layer Chromatography

The formation of various rhamnolipids was checked by TLC as shown in Figure

(4.8). The crude rhamnolipids contained both monorhamnolipid and dirhamnolipids.

The most active fragment is dirhamnolipid with diameter 10.5 cm by oil

displacement test.

Experimental Results

Table (4.2) Purification of biosurfactant using different mobile phases and oil

displacement test for each fraction

Fraction Quantity(g)

Mobile phase Colour Oil displacementtest (cm)

1 0.03 Hexane Light yellow -ve2 0.64 Acetone Light brown 33 0.02 Chloroform Brown 3.54 0.14 Chloroform: Methanol

( 2 : 1 v/v)Dark brown 10.5

5 0.05 Methanol Light brown 4

Figure (4.8) Thin layer chromatography of crude extract of biosurfactant

produced by P. aeruginosa TMN.

Experimental Results

4.4.2. Fourier transform infrared spectroscopy (FTIR)

Infrared spectrum of biosurfactant which is shown in Figure (4.9) indicated a

broad stretching peak at 3429cm 1, characteristic of O H stretching vibrations.

Absorption around 2922cm 1 is assigned to the symmetric stretch (- C H) of CH2 and

CH3 groups of aliphatic chains. The corresponding symmetric stretch is seen at

2853cm 1. Also, a weak symmetric stretching peak at 1715cm 1 indicate the presence

of ester carbonyl groups (C=O in COOH) in the biosurfactant. The ester carbonyl

group was proved from the band at 1321cm 1 which corresponds to C- O deformation

vibrations, although other groups also absorb in this region.

The presence of the carboxylic acid functional group in the molecule was also

confirmed by the bending of the hydroxyl (O H) of medium intensity bands in the

region (1455 1386 cm 1 -pyranyl II sorption band at 834 cm 1 showed the

presence of dirhamnolipid in the mixture.

These characteristic adsorption bands together demonstrate that both hold

chemical structures identical with those of rhamnolipids, which have rhamnose rings

and long hydrocarbon chains.

4.4.3. Nuclear magnetic resonance NMR

The 1H NMR spectrum of purified biosurfactant was represented in Figure

(4.10). Chemical shifts are assembled in Table (4.3).

4.4.4. Electrospray ionisation mass spectrometry (ESI MS)

In this study, thirteen rhamnolipid homologues with the pseudo molecular ions

-MS and summarized in Table

(4.4). The di rhamnolipid homologues which correspond to Rha-Rha-C10 C12

Figure (4.11).

Experimental Results

Figure (4.9) FTIR spectra of rhamnolipids produced by P. aeruginosa TMN.

Experimental Results

Figure (4.10) 1H-NMR spectrum of purified biosurfactant produced by P.

aeruginosa TMN.

Table (4.3) 1H-NMR chemical shift data for rhamnolipid components1H chemical shift(ppm) Multiplicity Assignment

0.8821.185, 1.2041.2761.574

2.545

3.372

3.3354.130

4.9385.275, 5.295

SingletDoubletMultipletMultiplet

Multiplet

Multiplet

MultipletMultiplet

SingletDoublet

CH3 (on b-hydroxyfatty acids)CH3 (on rhamnose moiety)(CH2)5 (on b-hydroxyfatty acids)(CH2) CH(O) CH2COO (on b-

hydroxyfatty acids)CH(O) CH2COO (on b-

hydroxyfatty acids)(CH2) CH(O-Rha) CH2COO (on

b-hydroxyfatty acids)CH OH (on rhamnose moiety)(CH2) CH( O C=O) CH2COO

(on b-hydroxyfatty acids)CH OH (on rhamnose moiety)CH O C (on rhamnose moiety)

Experimental Results

Table (4.4) Homologues of rhamnolipid produced by Pseudomonas aeruginosa,

using glucose as carbon source.

Rhamnolipid congeners Molecular massesRha-C12

Rha-C8 C8

Rha-C8 C10 Rha-C10 C8

Rha-C10 C12:1 Rha-C12:1 C10

Rha-C10 C12 Rha-C12 C10

Rha-Rha-C8

Rha-Rha-C10

Rha-Rha-C12

Rha-Rha-C8 C12:1 Rha-Rha-C12:1 C8

Rha-Rha-C10 C10

Rha-Rha-C10 C12 Rha-Rha-C12 C10

Rha-Rha-C12 C12:1 Rha-Rha-C10 C14:1

Rha Rha C12 C14

365447471529532456479508646651680703750

Experimental Results

Figure (4.11) ESI mass spectrum and chemical structure of Rha-Rha-C10 C12

rhamnolipid and its fragments.

Experimental Results

4.5. Oil displacement test for partially purified biosurfactants

Figure (4.12) indicate the diameters of the clear zones on the oil surface

obtained from oil displacement testing with the crude biosurfactant produced from

different carbon and nitrogen sources by Pseudomonas aeruginosa TMN. The type of

carbon and nitrogen substrates markedly affected the activity of biosurfactant. Oleic

acid, olive oil, glucose and glycerol showed the largest diameter of the clear zone

(14+0.22, 11+0.26, 10+0.85 and 10+0.37 cm respectively) among different carbon

substrates using NH4NO3 as nitrogen source. NaNO3 showed the largest diameter of

the clear zone (14+0.1 cm) among different nitrogen sources using glucose as carbon

sources.

4.6. Critical micelle concentration of partially purified

biosurfactants

The CMC can stand for surface activity of surfactant. A lower CMC of

surfactant means that the concentration for this surfactant to form micelle is also

lower, therefore, surfactant can change the surface property, exhibit some functions

such as emulsification, solubilization and foaming even at a relatively low

concentration. The CMC of the partially purified biosurfactant produced from glucose

as carbon source and NaNO3 as nitrogen source is presented in Figure (4.13). It can be

seen that the surface tension of biosurfactant decreased as its concentration increased,

until reaching the lowest, 34 mN/m. At this point, the concentration of biosurfactant,

namely CMC, was 18.75 mg/l. CMC for biosurfactants produced from different

carbon Figure (4.14) and nitrogen sources Figure (4.15) are summarized in Table

(4.5).

Experimental Results

Figure (4.12) Diameters of the clear zones on the oil surface obtained from oil

displacement testing with the crude biosurfactant produced from different

carbon and nitrogen sources by Pseudomonas aeruginosa TMN.

0

2

4

6

8

10

12

14

16

Glu

cose

Gly

cero

l

Suc

rose

Hex

ane

Ole

ic a

cid

Oliv

e oi

l

NH

4Cl

(50

mM

)

NaN

O3

(50

mM

)

Ure

a (5

0 m

M)

Yea

st e

xtra

ct(1

%)

NH

4NO

3 (5

0 m

M)

Carbon Source Nitrogen Source

Oil

Dis

plac

emen

tTes

t (cm

)

Experimental Results

Table (4.5) CMC of biosurfactants produced from different carbon and nitrogen

sources.

Different Carbon Sources CMC (mg/l)Glucose 18.75Glycerol 3.125Sucrose 2.5Hexane 4.38

Oleic acid 6.25Olive oil 6.25

Different Nitrogen SourcesNH4Cl (50 mM) 12.5NaNO3 (50 mM) 18.75Urea (50 mM) 12.5

Yeast extract (1%) 6.25NH4NO3 (50 mM) 18.75

Figure (4.13) CMC of partially purified biosurfactant produced using glucose ascarbon source and NaNO3 as nitrogen source.

Experimental Results

A. Glycerol as carbon source. B. Sucrose as carbon source.

C. Hexane as carbon source. D. Oleic acid as carbon source.

E. Olive oil as carbon source.

Figure (4.14) CMC of biosurfactants produced from different carbon sources.

Experimental Results

A. NH4Cl as nitrogen source. B. Urea as nitrogen source.

C. Yeast extract as nitrogen source. D. NH4NO3 as nitrogen source.

Figure (4.15) CMC of biosurfactants produced from different nitrogen sources.

Experimental Results

4.7. Estimation of Emulsification Activity

A partial purified biosurfactants produced from Pseudomonas aeruginosa

using different carbon and nitrogen sources were evaluated for their emulsification

activity using kerosene. Six carbon sources, i.e., glucose, glycerol, sucrose, hexane,

oleic acid, and olive oil were conducted for their effectiveness on biosurfactant

production. The results of various carbon sources for emulsification activity under

above mentioned conditions are as shown in Figure (4.16).

Emulsification activity for glucose, glycerol, sucrose, hexane, oleic acid, and

olive oil were 46+0.57, 49+0.86, 34+0.21, 5.67+0.13, 50+0.96 and 50+1.01 %

respectively. Among the six carbon sources tested, oleic acid and olive oil were the

best carbon sources with emulsification activity of 50%.

Nitrogen sources such as sodium nitrate, ammonium chloride, urea, yeast

extract, and ammonium nitrate were added to the fermentation media. Sodium nitrate

(46+0.35 %) and ammonium nitrate (46+0.21 %) were found most effective amongst

these nitrogen sources as shown in Figure (4.17). Other nitrogen sources urea

(41.7+0.34 %), ammonium chloride (41.7+0.23 %) and yeast extract (35.7+0.45 %)

gave low emulsification activity. Among the nitrogen sources added sodium nitrate

and ammonium nitrate gave similar results, but considering the cost factor sodium

nitrate was the most effective.

Experimental Results

Figure (4.16) Emulsifying activity (E24%) of biosurfactants obtained from

different carbon sources against kerosene

Figure (4.17) Emulsifying activity (E24%) of biosurfactants obtained from

different nitrogen sources against kerosene.

0

10

20

30

40

50

60

Glucose Glycerol Sucrose Hexane Oleic acid Olive oil

Carbon Source

Emul

sific

atio

nAc

tivity

(%)

05

101520253035404550

NH4Cl (50 mM) NaNO3 (50mM)

Urea (50 mM) Yeast extract(1%)

NH4NO3 (50mM)

Nitrogen Source

Emul

sific

atio

nAc

tivity

(%)

Discussion

CHAPTER V

DISCUSSION

Properties of Pseudomonas aeruginosa TMN isolated in this study are quite

similar with the literature (Lotfabad et al., 2009; Abbasi et al., 2012).

Literature revealed that the type and concentration of carbon and nitrogen

substrates markedly affected the production yield of rhamnolipid (Lang and

Wullbrandt, 1999; Mata-Sandoval et al., 1999; Maier and Soberón-Chávez, 2000;

Jeong et al., 2004).

Glucose was considered as the best carbon substrate based on rhamnolipid

yield and surface tension. P. aeruginosa TMN was unable to utilize sucrose

efficiently, resulting in a poor rhamnolipid yield and dry cell weight of 0.02 and 0.91

g/l, respectively. It is likely that the strain may lack the enzyme (i.e., invertase) for

sucrose hydrolysis (Wu et al., 2008). Boulton and Ratledge, 1987 mentioned that the

use of vegetable oil as carbon sources to produce rhamnolipids seemed to be an

interesting and low cost alternative but TMN strain attained a lower rhamnolipid yield

from olive oil than that from glucose and glycerol. The data showed that there seems

to be no clear trend between cell yield and rhamnolipid yield, which were, however,

strongly dependent on the carbon source used.

Use of oleic acid as the carbon source did not improve rhamnolipid production,

suggesting that hydrolysis of the oils was not the bottle- neck step. Some reports

showed that vegetable oils were more efficient substrates in rhamnolipid production

from P. aeruginosa TMN when compared with glucose, glycerol, and hydrocarbons

(Maier and Soberón-Chávez, 2000; Mata-Sandoval et al., 2001).

Moreover, hexane was also inefficient in cell growth and rhamnolipid

production; resulting in a low rhamnolipid yield and cell dry weight probably due to

its poor biodegradability (Chayabutra et al., 2001).

P. aeruginosa TMN used in this work showed a different trend, as glucose was

superior to the olive oils tested in terms of both rhamnolipid yield and surface tension.

Discussion

This suggests that the carbon source preference for rhamnolipid production seems to

be strain dependent.

These results are in concurrence with several previous reports, emphasizing the

utilization of glucose as cost-effective alternative substrates for microbial growth and

biosurfactant production (Lotfabad et al., 2009; Anyanwu and Chukwudi, 2010).

NaNO3 was the most efficient nitrogen source for P. aeruginosa TMN and

ammonium nitrate follows it to produce rhamnolipid. The observation that nitrate was

the better inorganic nitrogen source than ammonium ion for rhamnolipid production is

consistent with the finding of comparable studies in the literature (Guerra-Santos et

al., 1984; Arino et al., 1996; Wei and Chu, 1998; Anna et al., 2002; Santos et al.,

2002; Jeong et al., 2004).

Despite using urea and yeast extract as the organic nitrogen sources giving rise

to better cell growth, using those organic nitrogen sources had poorer performance

than using NaNO3 in terms of rhamnolipid yield. In fact, it has been reported that the

organic nitrogen source could help cell growth but was unfavorable to production of

glycolipid biosurfactant (Kim et al., 2006).

Addition of NaNO3 as the sole source of nitrogen in the glucose-containing

MS medium resulted in the highest biosurfactant production (0.98 g/l). Organic

nitrogen sources including yeast extract, and urea had approximately comparable

influence on the MR01 biosurfactant production (Lotfabad et al., 2009).

The yield of biosurfactant increased when pH varied from 4 to 8 and reached

the highest at pH 7; however, when pH reached 8, the biosurfactant yield declined and

came to its lowest point. Thus, indicated that the TMN strain excreted biosurfactant

more effectively under neutral conditions. These values agree with those previously

reported for rhamnolipid by the authors Priya and Usharani, 2009. They showed

maximum production of rhamnolipid at pH 7 using Pseudomonas aeruginosa.

Temperature and pH were two environmental factors that majorly affect biological

activities of prokaryotes (Yakimov et al., 1995).

Maximum rhamnolipid production was found at 37 C. Rhamnolipid

production increased with temperature from 25 to 37 C, and decreased slightly when

Discussion

temperature was further increased to 45 C. P. aeruginosa was unable to grow at 47

C, leading to negligible rhamnolipid production at that temperature.

These results suggest that the optimal temperature for rhamnolipid production

with TMN was in the range of 37 40 C. This result is comparable with the results

obtained by the authors Priya and Usharani, 2009. They showed maximum

biosurfactant production at 37 C.

Agitation rate affects the mass transfer efficiency of both oxygen and medium

components and is considered crucial to the cell growth and biosurfactant formation

of the strictly aerobic bacterium P. aeruginosa, especially when it was grown in a

shake flask. Rhamnolipid production increased and dry cell weight was also improved

at 200 rpm. The rhamnolipid production and dry cell weight are slightly decreased at

250 rpm. This result is comparable with the results obtained by the authors Maqsood

and Jamal, 2011. This can be attributed to the severe foaming while the flask was

shaken at 250 rpm. The heavy foaming caused by emulsification of rhamnolipid

during vigorous shaking (e.g., 250 rpm) may decrease the transfer efficiency of

oxygen gas into the liquid medium, and is thus unsuitable for rhamnolipid production

in shake-flask cultures. It should be pointed out that the heavy foaming during

biosurfactant fermentation is not necessarily a negative factor. Some researchers

collected the biosurfactant-rich foams from the fermenter for continuous recovery of

biosurfactant to avoid product inhibition and to simplify downstream processing

(Davis et al., 2001).

The fractions were collected and classified into 5 parts, Fraction 4 which has

dark brown colour and diameter of clear zone on the oil surface 10.5 cm at

concentration of 0.14 g/l was the best biosurfactant fraction. The biosurfactant

produced by P. aeruginosa have been characterized as rhamnolipids (Guerra-Santos

et al., 1984; Syldatk et al., 1985a; Parra et al., 1989; Rendell et al., 1990). They have

been found to increase the dispersion of hydrophobic compounds in water (Zhang and

Miller, 1992) and to enhance the recovery from soil of polyaromatic hydrocarbons

(PAH) (Van Dyke et al., 1993).

According to the results of the IR spectra, the rhamnolipids produced by P.

aeruginosa TMN belong to the glycolipid group, which is made up of aliphatic acid

Discussion

and ester. The absorption peak around 1051cm 1 was reported as C O C stretching in

the rhamnose (Pornsunthorntawee et al., 2008) -pyranyl II sorption band at 834

cm 1 showed the presence of dirhamnolipid in the mixture. The adsorption bands

obtained are consistent with the report of Guo et al. 2009 showing the presence of

rhamnose rings and hydrocarbon chains. In the FTIR spectrum, we could observe only

a minor shoulder; it might be because of the dirhamnolipid-rich biosurfactant

produced by TMN (Rahman et al., 2002).

The results obtained are consistent with the structure reported by (Stanghellini

and Miller, 1997; Pornsunthorntawee et al., 2008) consisting of aliphatic acid and the

glycolipid moiety.

The chemicals shifts were comparable to previous reports (Ramana and

Karanth, 1989; Sim et al., 1997; Wei et al., 2005) and the results were in accordance

with the structures as shown by ESI-spectra.

Thirteen rhamnolipid homologues were identified using ESI-MS (Lang and

Wullbrandt, 1999; Maier and Soberón-Chávez, 2000). The dirhamnolipid homologue

which corresponds to Rha-Rha-C10 C12

found. This work is in agreement with previous reports dealing with production of

rhamnolipids surfactant mixtures in which dirhamnolipids were the predominant

species (Arutchelvi and Doble, 2010). On the contrary, few reports (Arino et al.,

1996; Sim et al., 1997) describe a rhamnolipid mixture in which monorhamnolipid is

the predominant component.

The Oil displacement test is an indirect measurement of the surface activity of

a surfactant sample tested against oil; a larger diameter represents a higher surface

activity of the testing solution (Pornsunthorntawee et al., 2008). The oil spreading

method is rapid and easy to carry out, requires no specialized equipment and just a

small volume of sample et al., 2006). It can be applied when the activity and

quantity of biosurfactant is low. Youssef et al. 2004 and Plaza et al. 2006

demonstrated that the oil spreading technique is a reliable method to detect

biosurfactant production by diverse microorganisms. The assay was also applied for

screening by Huy et al. 1999. NaNO3 showed the largest diameter of the clear zone

among different nitrogen sources using glucose as carbon sources.

Discussion

The surface tension of biosurfactant decreased as its concentration increased,

until reaching the lowest, 34 mN/m. At this point, the concentration of biosurfactant,

namely CMC, was 18.75 mg/l. This CMC was much lower compared with some

chemical surfactant, for instance, sodium dodecyl sulfate (SDS) had a CMC value of

2100 mg/l (Chen et al., 2006). Nitschke and Pastore reported that the CMC of

biosurfactant obtained from Bacillus subtilis was 33 mg/l (Nitschke and Pastore,

2006). Li et al. 1984 showed that the CMC of rhamnolipid fermentation liquor was

386 mg/l (Daoshan et al., 2004).

Emulsification of rhamnolipids depends on carbon sources used to produce

rhamnolipids (Pornsunthorntawee et al., 2008). Six carbon sources, i.e., glucose,

glycerol, sucrose, hexane, oleic acid, and olive oil were conducted for their

effectiveness on biosurfactant production.

As evident from the results that Pseudomonas aeruginosa TMN was able to

grow in a medium containing oleic acid and olive oil with maximum emulsification

activity. Other carbon sources (glycerol, glucose, sucrose and hexane) used, showed

less emulsification activity, hexane being the lowest, as compared to other carbon

sources. This suggested that the carbon source preference of the strain for

biosurfactant production, which seems to be strain dependent (Wu et al., 2008). Most

microbial surfactants were substrate specific, solubilizing or emulsifying different

hydrocarbons at different rates.

Poor emulsification of other hydrocarbons might be due to the inability of the

biosurfactant to stabilize the microscopic droplets. Also inefficient oxygen supply in

the flask cultures may be responsible for poor growth in other carbon sources, as

biodegradation of these oils are known as an oxygen intensive metabolic event

(Panesar et al., 2009).

On other hand, oleic acid and olive oil are the carbon sources which are taken

up very easily than compared to others. Among the six carbon sources tested, oleic

acid and olive oil were the best carbon sources with emulsification activity of 50%.

Similar results were found with P. aeroginusa 44T1 (Robert et al., 1989). A probable

reason for this tendency that P. aeruginosa is lipase positive which facilitate

assimilation of fatty acids contained in oleic acid and olive oil.

Discussion

Sodium nitrate and ammonium nitrate were found most effective amongst the

nitrogen sources used. Other nitrogen sources (urea, ammonium chloride and yeast

extract) gave low emulsification activity. Among the nitrogen sources added sodium

nitrate and ammonium nitrate gave similar results, but considering the cost factor

sodium nitrate was the most effective. Similar results were reported elsewhere

(Ochsner et al., 1995; Arino et al., 1996; Healy et al., 1996).

Summary

SUMMARY

Surfactants are widely used for various purposes in industry, but for many

years were mainly chemically synthesized. It has only been in the past few decades

that biological surface active compounds (biosurfactants) have been described.

Biosurfactants are gaining prominence and have already taken over for a

number of important industrial uses, due to their advantages of low toxicity, high

biodegradability, low irritancy, and compatibility with human skin.

Biosurfactant-producing bacterial strain (Pseudomonas aeruginosa) was

isolated by using Cetrimide agar from Compost pile waste collected from El-Sharkia

region in Delta, Egypt. Morphological and biochemical characteristics of the isolate

indicated that strain was a member of the genus Pseudomonas. 16S rRNA gene

sequence analysis indicated that Strain TMN exhibited the highest similarity (98%) to

P. aeruginosa.

I. Several cultural conditions were studied to detect the optimum media used in

the production of biosurfactant from P. aeruginosa.

1. Different carbon sources (glucose, glycerol, sucrose, hexane, olive oil, and

oleic acid) were used to optimize the proper media used for biosurfactant

production. The result revealed that, maximum yield of biosurfactant (0.3 g

crude extract/l culture) and the maximum reduction in surface tension (34

mN/m) were obtained by using glucose as a carbon source and achieved an

emulsification index of over 46% for kerosene.

2. Different nitrogen sources (NH4NO3, NH4Cl, NaNO3, urea and yeast extract)

were investigated. NaNO3 was the most efficient nitrogen source to produce

rhamnolipid, giving a high rhamnolipid yield of (0.34 g crude extract/l

culture) and reduced surface tension of 34 mN/m and achieved an

emulsification index of over 46% for kerosene.

Summary

3. It was found that the yield of biosurfactant increased when pH varied from 4

to 8 and reached the highest at pH 7 (0.36 g crude extract/l culture).

4. It was found that the optimal temperature of rhamnolipid production was at

37 C where the rhamnolipid yield was 0.34 g crude extract/l culture.

5. The biosurfactant yield was maximized by increasing of the agitation rate.

Actually the maximum yield (0.3 g crude extract/l culture) was obtained at

agitation rate 200 rpm.

II. Column Chromatography was applied in the purification of biosurfactant;

fractions were collected and classified into 5 parts. The fraction of the fourth

part which had a dark brown in color and diameter of clear zone on the oil

surface 10.5 cm was the best biosurfactant. This purified biosurfactant was

characterized to elucidate its chemical formula.

III. FTIR, 1HNMR and ESI MS were applied to estimate the chemical

composition of the purified biosurfactant. The chemical structure is supposed

to be Rha-Rha-C10 C12.

IV. Oil displacement test for partially purified biosurfactants produced from

different carbon and nitrogen sources indicated that oleic acid showed the

largest diameter of the clear zone (14+0.22 cm) among different carbon

substrates and NaNO3 (14+0.1 cm) among different nitrogen sources.

V. CMC for biosurfactants produced from different carbon and nitrogen sources

indicated that glucose had higher CMC (18.75 mg/l) among different carbon

substrates while NaNO3 and NH4NO3 had higher CMC (18.75 mg/l for both)

among different nitrogen sources.

VI. Emulsification activity of the six carbon sources tested showed that oleic

acid and olive oil were the best carbon sources with emulsification activity of

50%. Sodium nitrate was found most effective amongst other nitrogen

sources with emulsification activity of 46%.

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Appendix

APPENDIX

Table (A.1): Effect of different carbon sources on the yield and surface tension:

Carbon Source Rhamnolipid Yield (g/l) Surface Tension (dyne/cm) P-value

Glucose 0.3+0.01 34+1.2 0.000*

Glycerol 0.25+0.02 46+0.99 0.000*

Sucrose 0.02+0.01 51+0.87 0.000*

Hexane 0.07+0.02 51+0.54 0.000*

Oleic acid 0.13+0.01 35+0.23 0.000*

Olive oil 0.19+0.03 36+0.45 0.000*

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent high Significant difference (P 0.001).

Table (A.2): Effect of different carbon sources on Dry cell weight and yield:

Carbon Source Dry cell weight (g/l) Rhamnolipid Yield (g/l) P-value

Glucose 2.9+ 0.02 0.3+0.01 0.000**

Glycerol 1.35+ 0.01 0.25+0.02 0.000**

Sucrose 0.91+ 0.03 0.02+0.01 0.000**

Hexane 0.2+ 0.01 0.07+0.02 0.004*

Oleic acid 5.0+ 0.06 0.13+0.01 0.000**

Olive oil 5.07+ 0.04 0.19+0.03 0.000**

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent Significant difference (P 0.05).

(**); represent high Significant difference (P 0.001).

Appendix

Table (A.3): Effect of different nitrogen sources on the yield and surface tension:

Nitrogen Source Rhamnolipid Yield (g/l) Surface Tension (dyne/cm) P-value

NH4Cl (50 mM) 0.04+0.01 37+1.0 0.000*

NaNO3 (50 mM) 0.34+0.01 34+0.69 0.000*

Urea (50 mM) 0.3+0.01 47+0.47 0.000*

Yeast extract (1%) 0.25+0.01 40+0.34 0.000*

NH4NO3 (50 mM) 0.3+0.01 36+0.22 0.000*

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent high Significant difference (P 0.001).

Table (A.4): Effect of different nitrogen sources on the dry cell weight and yield:

Nitrogen Source Dry cell weight (g/l) Rhamnolipid Yield (g/l) P-value

NH4Cl (50 mM) 3.61+ 0.1 0.04+0.01 0.000*

NaNO3 (50 mM) 4.2+ 0.03 0.34+0.01 0.000*

Urea (50 mM) 3.89+ 0.02 0.3+0.01 0.000*

Yeast extract (1%) 2.43+ 0.11 0.25+0.01 0.000*

NH4NO3 (50 mM) 2.9+ 0.08 0.3+0.01 0.000*

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent high Significant difference (P 0.001).

Appendix

Table (A.5): Effect of different PH on Rhamnolipid Yield and Dry cell weight:

PH Dry cell weight (g/l) Rhamnolipid Yield (g/l) P- value

4 0.9+ 0.02 0.11+ 0.01 0.000**

5 1.42+ 0.1 0.19+ 0.02 0.000**

6 1.51+ 0.04 0.27+ 0.05 0.000**

7 1.84+ 0.3 0.36+ 0.03 0.008*

8 1.04+ 0.02 0.12+ 0.02 0.000**

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent Significant difference (P 0.05).

(**); represent high Significant difference (P 0.001).

Table (A.6): Effect of different temperatures on Rhamnolipid Yield and Dry cell weight:

Temperature (oC) Dry cell weight (g/l) Rhamnolipid Yield (g/l) P-value

25 0.68+ 0.07 0.15+ 0.01 0.001*

30 1.42+ 0.01 0.21+ 0.02 0.000**

35 1.63+ 0.08 0.28+ 0.04 0.000**

"37" 4.2+ 0.05 0.34+ 0.07 0.000**

40 2.9+ 0.12 0.30+ 0.02 0.000**

45 2.1+ 0.10 0.19+ 0.05 0.000**

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent Significant difference (P 0.05).

(**); represent high Significant difference (P 0.001).

Appendix

Table (A.7): Effect of different agitation rates on Rhamnolipid Yield and Dry cell

weight:

Agitation Rate (rpm) Dry cell weight (g/l) Rhamnolipid Yield (g/l) P- value

100 0.94+ 0.05 0.14+ 0.01 0.000**

150 1.06+ 0.09 0.27+ 0.03 0.001*

200 1.8+ 0.03 0.3+ 0.01 0.000**

250 1.59+ 0.06 0.15+ 0.02 0.000**

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent Significant difference (P 0.05).

(**); represent high Significant difference (P 0.001).

Table (A.8): Effect of different carbon sources on yield and Oil displacement

test:

Carbon Source Rhamnolipid Yield (g/l) Oil displacement test (cm) P-value

Glucose 0.3+0.01 10+0.85 0.000*

Glycerol 0.25+0.02 10+0.37 0.000*

Sucrose 0.02+0.01 9+0.27 0.000*

Hexane 0.07+0.02 - -

Oleic acid 0.13+0.01 14+0.22 0.000*

Olive oil 0.19+0.03 11+0.26 0.000*

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent high Significant difference (P 0.001).

Appendix

Table (A.9): Effect of different nitrogen sources on the yield and Oil

displacement test:

Nitrogen Source Rhamnolipid Yield (g/l) Oil displacement test (cm) P-value

NH4Cl (50 mM) 0.04+0.01 5+0.07 0.000*

NaNO3 (50 mM) 0.34+0.01 14+0.1 0.000*

Urea (50 mM) 0.3+0.01 5+0.09 0.000*

Yeast extract (1%) 0.25+0.01 8+ 0.12 0.000*

NH4NO3 (50 mM) 0.3+0.01 10+0.02 0.000*

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent high Significant difference (P 0.001).

Table (A.10): Emulsification index of the carbon sources:

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent Significant difference (P 0.05).

(**); represent high Significant difference (P 0.001).

Carbon Source Emulsification Index P-valueGlucose 46+0.57 Ref.

Glycerol 49+0.86 0.043*

Sucrose 34+0.21 0.000**

Hexane 5.67+0.13 0.000**

Oleic acid 50+0.96 0.023*

Olive oil 50+1.01 0.026*

Appendix

Table (A.11): Emulsification index of the nitrogen sources:

Data were statistically described in terms of mean and standard error (±S.E.),

(*); represent high Significant difference (P 0.001).

( 0.05); represent non Significant difference.

Nitrogen Source Emulsification Index P-value

NH4Cl (50 mM) 41.7+0.23 Ref.

NaNO3 (50 mM) 46+0.35 0.000*

Urea (50 mM) 41.7+0.34 0.05

Yeast extract (1%) 35.7+0.45 0.000*

NH4NO3 (50 mM) 46+0.21 0.000*

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