Biosurfactants: From Genes to Applications (Microbiology Monographs, Vol. 20)

Microbiology Monographs Volume 20

Series Editor: Alexander Steinbu¨chel Mu¨nster, Germany

Microbiology Monographs Volumes published in the series

Inclusions in Prokaryotes Volume Editor: Jessup M. Shively Vol. 1, 2006

Microbial Megaplasmids Volume Editor: Edward Schwartz Vol. 11, 2009

Complex Intracellular Structures in Prokaryotes Volume Editor: Jessup M. Shively Vol. 2, 2006

Endosymbionts in Paramecium Volume Editor: Masahiro Fujishima Vol. 12, 2009

Magnetoreception and Magnetosomes in Bacteria Volume Editor: Dirk Schu¨ler Vol. 3, 2007

Alginates: Biology and Applications Volume Editor: Bernd H. A. Rehm Vol. 13, 2009

Predatory Prokaryotes – Biology, Ecology and Evolution Volume Editor: Edouard Jurkevitch Vol. 4, 2007

Plastics from Bacteria: Natural Functions and Applications Volume Editor: Guo Qiang Chen Vol. 14, 2010

Amino Acid Biosynthesis – Pathways, Regulation and Metabolic Engineering Volume Editor: Volker F. Wendisch Vol. 5, 2007

Amino-Acid Homopolymers Occurring in Nature Volume Editor: Yoshimitsu Hamano Vol. 15, 2010

Molecular Microbiology of Heavy Metals Volume Editors: Dietrich H. Nies and Simon Silver Vol. 6, 2007

Biology of Rhodococcus Volume Editor: He´ctor Alvarez Vol. 16, 2010

Microbial Linear Plasmids Volume Editors: Friedhelm Meinhardt and Roland Klassen Vol. 7, 2007

Structures and Organelles in Pathogenic Protists Volume Editor: W. de Souza Vol. 17, 2010

Prokaryotic Symbionts in Plants Volume Editor: Katharina Pawlowski Vol. 8, 2009

Plant Growth and Health Promoting Bacteria Volume Editor: Dinesh K. Maheshwari Vol. 18, 2010

Hydrogenosomes and Mitosomes: Mitochondria of Anaerobic Eukaryotes Volume Editor: Jan Tachezy Vol. 9, 2008

(Endo)symbiotic Methanogenic Archaea Volume Editor: Johannes H.P. Hackstein Vol. 19, 2010

Uncultivated Microorganisms Volume Editor: Slava S. Epstein Vol. 10, 2009

Biosurfactants Volume Editor: Gloria Sobero´n Cha´vez Vol. 20, 2011

Gloria Sobero´n-Cha´vez Editor

Biosurfactants From Genes to Applications

Editor Gloria Sobero´n Cha´vez Departamento de Biologı´a Molecular y Bioingenierı´a Instituto de Investigaciones Biome´dicas Universidad Nacional Auto´noma de Me´xico Apdo. Postal 70228, Ciudad Universitaria 04510, Me´xico D. F. Me´xico [email protected]

Series Editor Professor Dr. Alexander Steinbu¨chel Institut fu¨r Molekulare Mikrobiologie und Biotechnology Westfa¨lische Wilhelms Universita¨t Corrensstr. 3 48149 Mu¨nster Germany steinbu@uni muenster.de

ISSN 1862 5576 e ISSN 1862 5584 ISBN 978 3 642 14489 9 e ISBN 978 3 642 14490 5 DOI 10.1007/978 3 642 14490 5 Springer Heidelberg Dordrecht London New York Library of Congress Control Number: 2010937320 # Springer Verlag Berlin Heidelberg 2011 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Cover design: SPi Publisher Services Printed on acid free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface

The term surfactant refers to those compounds that have tension active properties, or in other words, the molecules that reduce interfacial tension. Their chemical composition can vary widely, but they have in common their amphiphilic or amphipatic nature and can thus be soluble in aqueous as well as in organic solvents. These characteristics make surfactants useful in a wide variety of industries, based on their abilities to lower surface tensions, increase solubility, their detergency power, wetting ability, and foaming capacity. However, the surfactants used by industry include so far almost exclusively synthetic surfactants. The term biosurfactant refers to different tensioactive compounds produced by living cells, but a large body of research has been carried out with those produced by bacteria, yeast, or fungi. These compounds include molecules with different chemical structures, which play different roles in the life-cycle of each of these microorganisms. Despite the very small amount of the surfactant market that is represented by biosurfactants, there is an increasing interest in microbial biosurfactants for several reasons. First, biosurfactants are considered environmentally “friendly” since they are relatively nontoxic and biodegradable. Second, biosurfactants have unique structures that are just starting to be appreciated for their potential applications to many different facets of industry, ranging from biotechnology to environmental cleanup. This “Microbiology Monographs” volume covers the current knowledge and the most recent advances in the field of microbial biosurfactants. Each chapter is written by one or more expert scientists working on one class of these biosurfactants. These reviews include the physicochemical properties of biosurfactants, their role in the physiology of the microbe that produced them, the biosynthetic pathway for their production, including the genetic regulation, and their potential biotechnological applications. We wish to thank the contributing authors of this book for the high quality of their work, their positive attitude in accepting suggestions, and very importantly for the generosity with their time to write lengthy manuscripts, which I am sure represented an important effort, despite having such tight agendas. Specially, v

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GS-Ch wants to thank Raina M. Maier for sharing her knowledge on the field of biosurfactants and for her friendship. We also want to acknowledge Jutta Lindenborn for her support in the editing process and Springer for publishing this monograph. We hope very much that this project of writing a book that reviewed the fascinating field of surfactants produced by microbes will be appreciated by the readers. Mexico City, Mexico Mu¨nster, Germany

Gloria Sobero´n-Cha´vez Alexander Steinbu¨chel

Contents

Biosurfactants: A General Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Gloria Sobero´n-Cha´vez and Raina M. Maier Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation, and Bioengineering of Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Ahmad Mohammad Abdel-Mawgoud, Rudolf Hausmann, Francois Le´pine, Markus M. Mu¨ller, and Eric De´ziel Surfactin and Other Lipopeptides from Bacillus spp. . . . . . . . . . . . . . . . . . . . . . . 57 Philippe Jacques Serrawettins and Other Surfactants Produced by Serratia . . . . . . . . . . . . . . . . 93 Tohey Matsuyama, Taichiro Tanikawa, and Yoji Nakagawa Trehalolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121 Zongze Shao Mannosylerythritol Lipids: Microbial Production and Their Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 J. Arutchelvi and M. Doble Sophorolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179 Inge N.A. Van Bogaert and Wim Soetaert Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211

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Biosurfactants: A General Overview Gloria Sobero´n-Cha´vez and Raina M. Maier

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 2 Physicochemical Properties, Formation of Micelles, and Other Aggregates . . . . . . . . . . . . . . . . 2 3 Biosurfactant Production in the Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 4 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

Abstract This Microbiology Monographs volume covers the current and most recent advances in the field of microbial surfactants. There is increasing interest in microbial biosurfactants for several reasons. First, biosurfactants are considered environmentally “friendly” since they are relatively nontoxic and biodegradable. Second, biosurfactants have unique structures that are just starting to be appreciated for their potential application to many different facets of the industry, ranging from biotechnology to environmental cleanup. The aim of this introductory chapter is to give a general overview of biosurfactants, their properties, their relationship to the synthetic surfactant industry, and their distribution in the environment.

G. Sobero´n Cha´vez (*) Departamento de Biologı´a Molecular y Bioingenierı´a, Instituto de Investigaciones Biome´dicas, Universidad Nacional Auto´noma de Me´xico, Apdo. Postal 70228, Ciudad Universitaria, Me´xico D. F. 04510, Me´xico e mail: [email protected] R.M. Maier Department of Soil, Water and Environmental Science, College of Agriculture and Life Sciences, The University of Arizona, Tucson, AZ, USA e mail: [email protected]

G. Sobero´n‐Cha´vez (ed.), Biosurfactants, Microbiology Monographs 20, DOI 10.1007/978 3 642 14490 5 1, # Springer Verlag Berlin Heidelberg 2011

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1 Introduction The demand for new specialty surfactants in the agriculture, cosmetic, food, pharmaceutical, and environmental industries is steadily increasing. Since these surfactants must be both effective and environmentally compatible, it is natural to turn to the microbial world to try to meet this demand (Banat et al. 2000; Mukherjee et al. 2006). Each chapter in this volume focuses on one class of biosurfactant produced by different microorganism and is written by one or more experts who work with these fascinating molecules. These reviews include the physicochemical properties of biosurfactants, their role in the physiology of the microbe that produces it, the biosynthetic pathway for their production, including the genetic regulation, and potential biotechnological applications.

2 Physicochemical Properties, Formation of Micelles, and Other Aggregates The term surfactant encompasses a wide variety of compounds, both synthetic and biological, all of which have tensioactive properties. These molecules are amphiphilic in nature, having both hydrophilic and hydrophobic domains that allow them to exist preferentially at the interface between polar and nonpolar media (Table 1, Fig. 1). Thus, surfactants tend to accumulate at interfaces (air water and oil water) as well as at air solid and liquid solid surfaces. Accumulation of surfactants at interfaces or surfaces results in the reduction of repulsive forces between dissimilar phases and allows the two phases to mix and interact more easily. Specifically, surfactants can reduce surface (liquid air) and interfacial (liquid liquid) tension. In fact, 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. The physicochemical characteristics that define a surfactant are its abilities to enhance the apparent water solubility of hydrophobic compounds, to form water hydrocarbon emulsions, and to reduce surface tension (Desai and Banat 1997). Typically, effective surfactants lower the surface tension between water and air from 72 to 35 mN/m and the interfacial tension between water and n-hexadecane from 40 to 1 mN/m. As surfactant monomers are added into the solution, the surface or interfacial tension will decrease until the surfactant concentration reaches what is known as the critical micelle concentration (cmc). Above the cmc, no further reduction in surface or interfacial tension is observed. At the cmc, surfactant monomers begin to spontaneously associate into structured aggregates such as micelles, vesicles, and lamellae (continuous bilayers). These aggregates form as a result of numerous weak chemical interactions between the polar head groups and the nonpolar tail groups including hydrophobic, Van der Waals, and hydrogen bonding. The cmc for any surfactant is dependent on the surfactant structure as

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Table 1 Examples of the more common biosurfactants and their origin Head group Biosurfactant Microorganism Corynebacterium lepus (Cooper et al. 1981) Fatty acids, neutral Fatty acid Neutral lipid N. erythropolis (Kretschmer et al. 1982) lipids, and Thiobacillus thiooxidans (Knickerbocker et al. 2000) phospholipids Phospholipid Lipopeptides Surfactin Bacillus subtilis, Bacillus pumilus A (Seydlova and Svobodova 2008) Viscosin Pseudomonas fluorescens, P. libanensis (Laycock et al. 1991) Serrawettin Serratia marcescens (Matsuyama et al. 1992) Glycolipids Mannosylerythritol Genus Pseudozyma (yeast), Candida antartica, lipids Ustilago maydis (Kitamoto et al. 2002) Sophorolipids C. batistae, T. bombicola, C. lypolytica, C. bombicola, T.apicola, T.petrophilum, C. bogoriensis (Van Bogaert et al. 2007) Rhamnolipids Pseudomonas sp., P. aeruginosa (Reiling et al. 1986) Trehaloselipids Rhodococcus sp., Arthrobacter sp., R. erythropolis, N. erythropolis (Lang and Philp 1998) Cellobiolipids Ustilago zeae, Ustilago maydis (Hewald et al. 2005) Polymeric Emulsan Acinetobacter calcoaceticus (Rosenberg and Ron 1999) Biodispersan A. calcoaceticus (Rosenberg and Ron 1997) Mannan lipid protein C. tropicalis (Rosenberg and Ron 1999) Alasan A. radioresistens (Navonvenezia et al. 1995) Siderophore Flavolipids Flavobacterium (Bodour et al. 2004)

Fig. 1 Representative structures of biosurfactants

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well as the pH, ionic strength, and temperature of the solution. Further, the aggregate structure is dictated by the polarity of the solvent in which the surfactant is dissolved. For example, in an aqueous solution, the polar head groups of a micelle will be oriented outward toward the aqueous phase, and the hydrophobic tails will associate in the core of the micelle (oil in water micelle). In contrast, in oil, the polar head groups will associate in the center of the micelle while the hydrophobic tails will be oriented toward the outside (water in oil micelle). Biosurfactants are produced by plants and animals as well as microorganisms. A large body of research has been carried out on surfactants produced by bacteria, yeast, and fungi, and the aim of this book is to provide a review of the various microbial surfactants that have been identified. Microbial surfactants have an amazingly wide range of chemical structures (Fig. 1) and each appears to play different roles in the life cycle of the producing microorganism (Ron and Rosenberg 2001). Biosurfactants display important biological activities, including antibiotic, antifungal, insecticidal, antiviral, immunomodulator, and antitumoral activities. These activities are the basis for growing interest in a number of specialty applications such as biological control of pests in agriculture (Stanghellini and Miller 1997), cancer treatment (Saini et al. 2008), and wound healing (Piljac et al. 2008; Stipcevic et al. 2006). The biosurfactants that have been studied in most detail include the rhamnolipids produced by Pseudomonas aeruginosa (Soberon-Chavez et al. 2005) and different Burkholderia species (Dubeau et al. 2009), and surfactin, a lipopeptide that is synthesized by Bacillus subtilis (Mulligan 2005). These are examples of effective biosurfactants that can each reduce the surface tension between pure water and air, from 73 mN/m to less than 30 mN/m. The worldwide surfactant industry was valued at $20 billion in 2006 with the United States/Canada, Western Europe, and China accounting for approximately 70% of the market (Janshekar et al. 2007). Surfactants are used in a wide variety of industries that produce household and industrial cleaners, personal care products, and in various types of manufacturing including food processing and the production of plastics, paints and coatings, textiles, pulp and paper, and agricultural products. These compounds are also used in the specialty chemical market as components of cosmetic products, pharmaceuticals, emulsifiers, wetting agents, and in the synthesis of fine chemicals. Presently, the vast majority of surfactants used are synthetic; however, in light of their unique chemical characteristics, biosurfactants have been recognized for their potential utility in many applications in a number of recent reviews, e.g., (Bonmatin et al. 2003; Desai and Banat 1997; Kosaric 2001; Kralova and Sjoblom 2009; Lang and Wullbrandt 1999; Maier and Soberon-Chavez 2000; Nitschke and Costa 2007; Ritter 2004; Rodrigues et al. 2006a; Ron and Rosenberg 2001; Rosenberg and Ron 1999; Singh et al. 2007). The most important limitation for the commercial use of biosurfactants is the complexity and high cost of production, which has limited the development of their use on a large scale. Thus far, the only commercially available biosurfactants are rhamnolipids and surfactin. Rhamnolipids are produced by a small number of companies such as Jeneil Biosurfactant Company (JBR products) and Rhamnolipid, Inc. (http://www.rhamnolipidholdings.com). However, even these companies do

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not always have readily available product for sale, underscoring the tenuous nature of the commercial availability of biosurfactants to date. One of the reasons that there are commercial sources of rhamnolipids is that they are the only biosurfactant thus far that has been approved by US Environmental Protection Agency for use in food products, cosmetics, and pharmaceuticals (Nitschke and Costa 2007). Despite the current limitations to commercial production of biosurfactants, there is great interest in these materials since they are considered to be “green” alternatives to synthetic surfactants (Ritter 2004). Biosurfactants are considered relatively nontoxic and biodegradable, but perhaps more importantly, the chemical structure of biosurfactants is unique and exhibits great structural diversity including glycolipids, lipopeptides, fatty acids and neutral lipids, siderophore lipids, and polymeric surfactants (Table 1). Biosurfactants are intriguing, in that they are produced as complex mixtures of up to 40 congeners where the hydrophilic head groups are fairly conserved and the hydrophobic tail groups have considerable variation (Bodour et al. 2004; Monteiro et al. 2007). Component congeners within these complex mixtures can have remarkably different properties, and behavioral differences between biosurfactant classes can be equally divergent. In contrast to common synthetic surfactants that typically possess alkyl chains of ten or more carbon units, many biosurfactants possess surprisingly short alkyl chains. Such structures enhance the aqueous solubility of these biosurfactants but render Van der Waals attractive interactions relatively weak. Despite this structural attribute, biosurfactants do aggregate in solution, supplemented by intermolecular forces such as hydrogen bonding. Moreover, not only do these biosurfactants aggregate in solution, but they also exhibit powerful surfactant activity at both liquid and solid surfaces. Thus, it seems that nature has uniquely and intentionally positioned these biosurfactants to possess characteristics that lie at the boundary between conventional hydrophilic and hydrophobic organic molecules. The structures of these biosurfactants are quite elegant and, in many cases, defy conventional chemical intuition that would predict little surface activity. Despite their aqueous solubility, they can have remarkably low cmcs when compared to structurally similar synthetic surfactants. For example, nonionic (i.e., low pH), multicomponent monorhamnolipid mixtures, in which the heptyl chain congener is most prevalent, have cmc values that range from <1 mM to ~10 mM depending on solution ionic strength (Lebron-Paler 2008). This cmc value increases as the pH increases and the rhamnolipids become deprotonated, reaching values on the order of ~100 mM at pH 8. In contrast, the nonionic alkyl glucosides and glucamides of comparable alkyl chain length that are structurally similar to the rhamnolipids at pH 4 have cmc values on the order of 10 3 M or higher (Nickel et al. 1992; Soderman and Johansson 2000; Zhang and Marchant 1996), at least one order of magnitude and in many cases several orders of magnitude greater than those of the rhamnolipids. As a result of their remarkable properties, there is tremendous interest in these molecules for uses as diverse as bioremediation of organics (e.g., Chen et al. 2005; Garcia-Junco et al. 2003; Mulligan 2009; Olivera et al. 2000; Schippers et al. 2000; Shin et al. 2005; Urum and Pekdemir 2004; Uysal and Turkman 2005) and metals (e.g., Dahrazma and Mulligan 2007; Mulligan et al. 2001; Neilson et al. 2003; Ron and Rosenberg

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2002; Sandrin et al. 2000; Tan et al. 1994; Torrens et al. 1998; Wang and Mulligan 2009; Wen et al. 2009), cosmetic additives (Hayase and Ishihata 2006; Pianelli et al. 2002; Yoneda et al. 2001), pharmaceutical preparations (Piljac et al. 2008; Singh and Cameotra 2004; Stipcevic et al. 2005, 2006), and coatings resistant to bacteria (Meylheuc et al. 2006; Rodrigues et al. 2004; Rodrigues et al. 2006b; Vollenbroich et al. 1997).

3 Biosurfactant Production in the Environment Biosurfactant production by bacteria has been studied mainly from the perspective of biotechnological potential. Thus, biosurfactant-producing organisms have been isolated from a wide diversity of environments including soil, sea water, marine sediments, oil fields (Yakimov et al. 1998) and even extreme environments (Cameotra and Makkar 1998). Not only are biosurfactant producers widely distributed but also many different microbes produce biosurfactants. Bacterial genera described to produce surfactants include: Pseudomonas, Rhodococcus, Mycobacterium, Nocardia, Flavobacterium, Corynebacterium, Clostridium, Acinetobacter, Thiobacillus, Bacillus, Serratia, Arthrobacter, and Alcanivorax (Bodour et al. 2003). Some of these genera have multiple species that produce different kinds of surfactants. For example, P. aeruginosa produces rhamnolipids, and several Pseudomonas species, including P. fluorescens, produce cyclic lipopeptides (CLP) (Raaijmakers et al. 2006), which are similar to surfactin and other CLP produced by Bacillus. There are still relatively few studies that have addressed the frequency and distribution of biosurfactant producers in the environment. These studies suggest that only a small fraction of the community is capable of biosurfactant production unless a selective pressure exists. For example, one report determined the distribution of culturable bacteria that were able to produce biosurfactants from undisturbed and contaminated sites (Bodour et al. 2003). Twenty sites were sampled resulting in 1,305 isolates of which 45, or 3.4% of the total, were found to be biosurfactant producers. Of the 45 biosurfactant producers, there were 16 unique isolates, which included B. subtilis, P. aeruginosa, Pseudomonas sp., and a Flavobacterium sp. Biosurfactant producers were obtained from a majority, but not all, of the soils tested. Of the three soils tested that were co-contaminated with both organics and metals, 8.4% of the 203 isolates obtained produced surfactants, suggesting that some environments may have greater selective pressure for biosurfactants production than others. The biosurfactant produced by the Flavobacterium isolate was subsequently identified and represents an entirely new class of biosurfactants, the flavolipids (Bodour et al. 2004). In a more recent study, (Toribio et al. unpublished) screened for biosurfactant production in a group of 700 bacterial isolates, mainly Pseudomonas, taken over a 4 year period (2003 2005 and 2007) from the extremely oligotrophic water column at Cuatro Cie´negas Basin in the Mexican state of Coahuila (Souza et al. 2006). Only six of the isolates obtained produced a biosurfactant including one P. aeruginosa,

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one Pseudomonas moseelii, three Pseudomonas koreensis, and one Serratia marcescens (Toribio et al. unpublished). Although these results are not completely comparable with the results discussed previously (Bodour et al. 2003), because the screening for bacteria was done in oligotrophic water samples and was biased for Pseudomonas, the screening was not restricted to this genus. Based on the results of these two studies, the fraction of culturable isolates that produce biosurfactants is small. However, it is possible and even likely that other biosurfactant-producing populations were present in the soil and water samples tested but were not enriched by the screening conditions used. Despite the limitations of the screening methods used in these studies, a diverse group of biosurfactantproducing organisms was obtained, including one novel biosurfactant producer (Flavobacterium sp.), suggesting that a more exhaustive screening has the potential to yield other new biosurfactant producers. The question remains: what is the role of biosurfactant production in the ecology of environmental bacteria? Although many roles have been proposed based on pure culture studies, researchers are just beginning to explore how biosurfactants function in situ. Since biosurfactant production requires valuable resources and energy from the producing isolate, it is likely that their production provides an advantage in the competition for resources or in protection under harsh environmental conditions. For example, (Toribio et al. unpublished) propose that biosurfactant production by P. koreensis isolates from the Cuatro Cie´negas Basin might play an ecological role in survival in this extremely oligotrophic environment, enabling the bacteria that produce these tensioactive compounds to restrain the growth of competitors when growing as a part of biofilms or bacterial mats. As a second example, a study of 57 polyaromatic hydrocarbon-degrading isolates obtained from hydrocarbon-contaminated soil sites showed that 67% of the isolates produced surfactants, suggesting that biosurfactants production is an important characteristic in this subset of microorganisms (Willumsen and Karlson 1997). Interestingly, the production of biosurfactants by these isolates did not necessarily correlate with their ability to degrade hydrocarbons. As a final example, a recent study examined heterotrophic bacteria that were cultured from a mine tailings site. The mine site is characterized by low pH (2.5 4) and high metal content (up to 4 g/kg of arsenic and lead respectively). Total cultural counts on R2A agar (at neutral pH) were approximately 600 CFU/g tailings, a very low count, which indicates the high level of stress in this environment. Five unique isolates were obtained from this study and all of them produce biosurfactants, suggesting that this may be an important survival trait for microbes in this site (Solis-Dominguez and Maier unpublished).

4 Conclusions The chapters presented in this book provide in-depth reviews of the best-studied major groups of biosurfactants discovered thus far including the rhamnolipids, surfactin and related lipopeptides, the serrawettins, trehalose lipids, mannosyl

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Number of Publications

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Fig. 2 The number of publications obtained from a year by year search for the term “biosurfac tant” on the Thomson Reuters ISI Web of Knowledge search platform (2009)

erythriol lipids, and sophorolipids. There is a rapidly increasing body of research on these molecules as well as other newly discovered biosurfactants as evidenced by the rapidly growing number of publications on the topic of biosurfactants (Fig. 2). We expect in the next 5 10 years, as yields increase and production costs decrease, new biosurfactants continue to be discovered, and the chemistry and potential applications of these molecules are better understood, that biosurfactants will begin to compete favorably with synthetic surfactants in the surfactant industry, particularly in specialty surfactant markets. Acknowledgments RMM gratefully acknowledges financial support of this work by the National Science Foundation (CHE 0714245) and invaluable discussions with collaborator Jeanne E. Pemberton on biosurfactant structure and chemistry. GS Ch acknowledges financial support from CONACYT (50201) and DGPA UNAM PAPIIT (IN200707.)

References Banat IM, Makkar RS, Cameotra SS (2000) Potential commercial applications of microbial surfactants. Appl Microbiol Biotechnol 53:495 508 Bodour AA, Drees KP, Maier RM (2003) Distribution of biosurfactant producing bacteria in undisturbed and contaminated arid southwestern soils. Appl Environ Microbiol 69:3280 3287 Bodour AA, Guerrero Barajas C, Jiorle BV, Malcomson ME, Paull AK, Somogyi A, Trinh LN, Bates RB, Maier RM (2004) Structure and characterization of flavolipids, a novel class of biosurfactants produced by Flavobacterium sp strain MTN11. Appl Environ Microbiol 70:114 120

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Bonmatin JM, Laprevote O, Peypoux F (2003) Diversity among microbial cyclic lipopeptides: iturins and surfactins. Activity structure relationships to design new bioactive agents. Comb Chem High Throughput Screen 6:541 556 Cameotra SS, Makkar RS (1998) Synthesis of biosurfactants in extreme conditions. Appl Micro biol Biotechnol 50:520 529 Chen G, Qiao MQ, Zhang HY, Zhu HL (2005) Sorption and transport of naphthalene and phenanthrene in silica sand in the presence of rhamnolipid biosurfactant. Sep Sci Technol 40: 2411 2425 Cooper DG, Zajic JE, Denis C (1981) Surface active properties of a biosurfactant from Coryne bacterium lepus. J Am Oil Chem Soc 58:77 80 Dahrazma B, Mulligan CN (2007) Investigation of the removal of heavy metals from sediments using rhamnolipid in a continuous flow configuration. Chemosphere 69:705 711 Desai JD, Banat IM (1997) Microbial production of surfactants and their commercial potential. Microbiol Mol Biol Rev 61:47 64 Dubeau D, Deziel E, Woods DE, Lepine F (2009) Burkholderia thailandensis harbors two identical rhl gene clusters responsible for the biosynthesis of rhamnolipids. BMC Microbiol 9:263 Garcia Junco M, Gomez Lahoz C, Niqui Arroyo JL, Ortega Calvo JJ (2003) Biosurfactant and biodegradation enhanced partitioning of polycyclic aromatic hydrocarbons from nonaqueous phase liquids. Environ Sci Technol 37:2988 2996 Hayase M, Ishihata S (2006) Development of skin care products by using fermentation technology. Fragr J 34:83 89 Hewald S, Josephs K, B€ olker M (2005) Genetic analysis of biosurfactant production in Ustilago maydis. Appl Environ Microbiol 71:3033 3040 Janshekar H, Chang RJ, Yokose K, Ma X (2007) Surfactants. (4/23/2010; http://www.sriconsul ting.com/SCUP/Public/Reports/SURFA000/) Kitamoto D, Isoda H, Nakahara T (2002) Functions and potential applications of glycolipid biosur factants from energy saving materials to gene delivery carriers. J Biosci Bioeng 94:187 201 Knickerbocker C, Nordstrom DK, Southam G (2000) The role of “blebbing” in overcoming the hydrophobic barrier during biooxidation of elemental sulfur by Thiobacillus thiooxidans. Chem Geol 169:425 433 Kosaric N (2001) Biosurfactants and their application for soil bioremediation. Food Technol Biotechnol 39:295 304 Kralova I, Sjoblom J (2009) Surfactants used in food industry: a review. J Dispersion Sci Technol 30:1363 1383 Kretschmer A, Bock H, Wagner F (1982) Chemical and physical characterization of interfacial active lipids from Rhodococcus erythropolis grown on normal alkanes. Appl Environ Micro biol 44:864 870 Lang S, Philp JC (1998) Surface active lipids in rhodococci. Antonie Van Leeuwenhoek Int J Gen Mol Microbiol 74:59 70 Lang S, Wullbrandt D (1999) Rhamnose lipids biosynthesis, microbial production and applica tion potential. Appl Microbiol Biotechnol 51:22 32 Laycock MV, Hildebrand PD, Thibault P, Walter JA, Wright JLC (1991) Viscosin, a potent peptidolipid biosurfactant and phytopathogenic mediator produced by a pectolytic strain of Psedomonas fluorescens. J Agric Food Chem 39:483 489 Lebron Paler A (2008) Solution and interfacila characterization of rhamnolpid biosurfactant from Pseudomonas aeruginosa ATCC 9027. PhD Dissertation, University of Arizona Maier RM, Soberon Chavez G (2000) Pseudomonas aeruginosa rhamnolipids: biosynthesis and potential applications. Appl Microbiol Biotechnol 54:625 633 Matsuyama T, Kaneda K, Nakagawa Y, Isa K, Hara Hotta H, Yano I (1992) A novel extracellular cyclic lipopeptide which promotes flagellum dependent and independent spreading growth of Serratia marcescens. J Bacteriol 174:1769 1776 Meylheuc T, Methivier C, Renault M, Herry JM, Pradier CM, Bellon Fontaine MN (2006) Adsorption on stainless steel surfaces of biosurfactants produced by gram negative and gram positive bacteria: consequence on the bioadhesive behavior of Listeria monocytogenes. Colloids Surf B Biointerfaces 52:128 137

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Monteiro SA, Sassaki GL, de Souza LM, Meira JA, de Araujo JM, Mitchell DA, Ramos LP, Krieger N (2007) Molecular and structural characterization of the biosurfactant produced by Pseudomonas aeruginosa DAUPE 614. Chem Phys Lipids 147:1 13 Mukherjee S, Das P, Sen R (2006) Towards commercial production of microbial surfactants. Trends Biotechnol 24:509 515 Mulligan CN (2005) Environmental applications for biosurfactants. Environ Pollut 133:183 198 Mulligan CN (2009) Recent advances in the environmental applications of biosurfactants. Curr Opin Colloid Interface Sci 14:372 378 Mulligan CN, Yong RN, Gibbs BF (2001) Heavy metal removal from sediments by biosurfactants. J Hazard Mater 85:111 125 Navonvenezia S, Zosim Z, Gottlieb A, Legmann R, Carmeli S, Ron EZ, Rosenberg E (1995) Alasan, a new bioemulsifier from Acinetobacter radioresistens. Appl Environ Microbiol 61:3240 3244 Neilson JW, Artiola JF, Maier RM (2003) Characterization of lead removal from contaminated soils by nontoxic soil washing agents. J Environ Qual 32:899 908 Nickel D, Nitsch C, Kurzendorfer P, von Rybinski W (1992) Trends in colloid and interface science VI. Springer, Berlin Nitschke M, Costa S (2007) Biosurfactants in food industry. Trends Food Sci Technol 18:252 259 Olivera NL, Commendatore MG, Moran AC, Esteves JL (2000) Biosurfactant enhanced degradation of residual hydrocarbons from ship bilge wastes. J Ind Microbiol Biotechnol 25:70 73 Pianelli G, Kado T, Yosioka T (2002) Characteristics and cosmetic applications of sopholiance (sophorolipid). Fragr J 30:86 92 Piljac A, Stipcevic T, Piljac Zegarac J, Piljac G (2008) Successful treatment of chronic decubitus ulcer with 0.1% dirhamnolipid ointment. J Cutan Med Surg 12:142 146 Raaijmakers JM, de Bruijn I, de Kock MJD (2006) Cyclic lipopeptide production by plant associated Pseudomonas spp.: diversity, activity, biosynthesis, and regulation. Mol Plant Microbe Interact 19:699 710 Reiling HE, Thaneiwyss U, Guerrasantos LH, Hirt R, Kappeli O, Fiechter A (1986) Pilot plant production of rhamnolipid surfactant by Pseudomonas aeruginosa. Appl Environ Microbiol 51:985 989 Ritter SK (2004) Green innovations. Chem Eng News 82:25 30 Rodrigues L, van der Mei HC, Teixeira J, Oliveira R (2004) Influence of biosurfactants from probiotic bacteria on formation of biofilms on voice prostheses. Appl Environ Microbiol 70:4408 4410 Rodrigues L, Banat IM, Teixeira J, Oliveira R (2006a) Biosurfactants: potential applications in medicine. J Antimicrob Chemother 57:609 618 Rodrigues L, van der Mei H, Banat IM, Teixeira J, Oliveira R (2006b) Inhibition of microbial adhesion to silicone rubber treated with biosurfactant from Streptococcus thermophilus A. FEMS Immunol Med Microbiol 46:107 112 Ron EZ, Rosenberg E (2001) Natural roles of biosurfactants. Environ Microbiol 3:229 236 Ron EZ, Rosenberg E (2002) Biosurfactants and oil biorremediation. Curr Opin Biotechnol 13:249 252 Rosenberg E, Ron EZ (1997) Bioemulsans: microbial polymeric emulsifiers. Curr Opin Biotech nol 8:313 316 Rosenberg E, Ron EZ (1999) High and low molecular mass microbial surfactants. Appl Micro biol Biotechnol 52:154 162 Saini HS, Barragan Huerta BE, Lebron Paler A, Pemberton JE, Vazquez RR, Burns AM, Marron MT, Seliga CJ, Gunatilaka AAL, Maier RM (2008) Efficient purification of the biosurfactant viscosin from Pseudomonas libanensis strain M9 3 and its physicochemical and biological properties. J Nat Prod 71:1011 1015 Sandrin TR, Chech AM, Maier RM (2000) A rhamnolipid biosurfactant reduces cadmium toxicity during naphthalene biodegradation. Appl Environ Microbiol 66:4585 4588

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Schippers C, Gessner K, Muller T, Scheper T (2000) Microbial degradation of phenanthrene by addition of a sophorolipid mixture. J Biotechnol 83:189 198 Seydlova G, Svobodova J (2008) Review of surfactin chemical properties and the potential biomedical applications. Cent Eur J Med 3:123 133 Shin KH, Ahn Y, Kim KW (2005) Toxic effect of biosurfactant addition on the biodegradation of phenanthrene. Environ Toxicol Chem 24:2768 2774 Singh A, Van Hamme JD, Ward OP (2007) Surfactants in microbiology and biotechnology: Part 2. Application aspects. Biotechnol Adv 25:99 121 Singh P, Cameotra SS (2004) Potential applications of surfactants in biomedical sciences. Trends Biotechnol 22:142 146 Soberon Chavez G, Lepine F, Deziel E (2005) Production of rhamnolipids by Pseudomonas aeruginosa. Appl Microbiol Biotechnol 68:718 725 Soderman O, Johansson I (2000) Polyhydroxyl based surfactants and their physico chemical properties and applications. Curr Opin Colloid Interface Sci 4:391 401 Souza V et al (2006) An endangered oasis of aquatic microbial biodiversity in the Chihuahuan desert. Proc Natl Acad Sci USA 103:6565 6570 Stanghellini ME, Miller RM (1997) Their identity and potential efficacy in the biological control of zoosporic plant pathogens. Plant Dis 81:4 12 Stipcevic T, Piljac T, Isseroff RR (2005) Di rhamnolipid from Pseudomonas aeruginosa displays differential effects on human keratinocyte and fibroblast cultures. J Dermatol Sci 40:141 143 Stipcevic T, Pijac A, Pijac G (2006) Enhanced healing of full thickness burn wounds using di rhamnolipid. Burns 32:24 34 Tan H, Champion JT, Artiola JF, Brusseau ML, Miller RM (1994) Complexation of cadmium by a rhamnolipid biosurfactant. Environ Sci Technol 28:2402 2406 Toribio J, Escalante AE, Caballero Mellado J, Gonza´lez Gonza´lez A, Zavala S, Souza V, Sobero´n Cha´vez G (2010) Characterization of a novel biosurfactant producing Pseudomonas koreensis lineage that is endemic to Cuatro Cie´negas Basin. Syst Appl Microbiol (Submitted) Torrens JL, Herman DC, Miller Maier RM (1998) Biosurfactant (rhamnolipid) sorption and the impact on rhamnolipid facilitated removal of cadmium from various soils under saturated flow conditions. Environ Sci Technol 32:776 781 Urum K, Pekdemir T (2004) Evaluation of biosurfactants for crude oil contaminated soil washing. Chemosphere 57:1139 1150 Uysal A, Turkman A (2005) Effect of biosurfactant on 2, 4 dichlorophenol biodegradation in an activated sludge bioreactor. Process Biochem 40:2745 2749 Van Bogaert INA, Saerens K, De Muynck C, Develter D, Soetaert W, Vandamme EJ (2007) Microbial production and application of sophorolipids. Appl Microbiol Biotechnol 76:23 34 Vollenbroich D, Pauli G, Ozel M, Vater J (1997) Antimycoplasma properties and application in cell culture of surfactin, a lipopeptide antibiotic from Bacillus subtilis. Appl Environ Microbiol 63:44 49 Wang SL, Mulligan CN (2009) Arsenic mobilization from mine tailings in the presence of a biosurfactant. Appl Geochem 24:928 935 Wen J, Stacey SP, McLaughlin MJ, Kirby JK (2009) Biodegradation of rhamnolipid, EDTA and citric acid in cadmium and zinc contaminated soils. Soil Biol Biochem 41:2214 2221 Willumsen PA, Karlson U (1997) Screening of bacteria, isolated from PAH contaminated soils, for production of biosurfactants and bioemulsifiers. Biodegradation 7:415 423 Yakimov MM, Golyshin PN, Lang S, Moore ERB, Abraham WR, Lunsdorf H, Timmis KN (1998) Alcanivorax borkumensis gen. nov., sp. nov., a new, hydrocarbon degrading and surfactant producing marine bacterium. Int J Syst Bacteriol 48:339 348 Yoneda T, Tsuzuki T, Ogata E, Fusyo Y (2001) Surfactin sodium salt: an excellent bio surfactant for cosmetics. J Cosmet Sci 52:153 154 Zhang TH, Marchant RE (1996) Novel polysaccharide surfactants: the effect of hydrophobic and hydrophilic chain length on surface active properties. J Colloid Interface Sci 177:419 426

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation, and Bioengineering of Production Ahmad Mohammad Abdel-Mawgoud, Rudolf Hausmann, Francois Le´pine, Markus M. M€ uller, and Eric De´ziel

Contents 1 2

Introduction and Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rhamnolipid Structure, Detection, and Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Methods of Detection and Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Biosynthesis and Genetic Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Biosynthesis of Rhamnolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Regulation of Rhamnolipid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Bioengineering of Rhamnolipid Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Production by P. aeruginosa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Alternatives to P. aeruginosa for Rhamnolipid Production . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusion: Prospectives for the Industrial Production of Rhamnolipids . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

14 16 16 17 21 22 29 34 34 41 43 43

Abstract As promising biotechnological products, rhamnolipids (RLs) are the most investigated biosurfactants. Over the years, important efforts have been spent and an array of techniques has been developed for the isolation of producing bacterial strains and the characterization of a large variety of RL homologs and congeners. Investigations on RL production by the best known producer, the opportunistic pathogen Pseudomonas aeruginosa, have shown that production of RLs proceeds through de novo biosynthesis of precursors. Over the last 15 years, the genetic details underlying RL production in P. aeruginosa have been mostly unraveled, revealing a complex regulatory mechanism controlled by quorum sensing pathways of intercellular communication. A number of nutritional and A. Mohammad Abdel Mawgoud, F. Le´pine, and E. De´ziel (*) INRS Institut Armand Frappier, Laval, QC, Canada H7V 1B7 e‐mail: [email protected] R. Hausmann, and M.M. M€ uller Institute of Engineering in Life Sciences, Section of Technical Biology, Research University Karlsruhe, Engler Bunte Ring 1, 76131 Karlsruhe, Germany

G. Sobero´n‐Cha´vez (ed.), Biosurfactants, Microbiology Monographs 20, DOI 10.1007/978 3 642 14490 5 2, # Springer Verlag Berlin Heidelberg 2011

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cultivation factors affecting RL productivity have also been identified, while the use of many affordable and renewable raw substrates has been described to optimize the production. Multidisciplinary approaches are increasingly adopted to develop methods for the safe, cost-effective, and highly efficient production of RLs at the industrial scale.

1 Introduction and Overview Rhamnolipids (RLs), the glycolipid biosurfactants produced mainly by Pseudomonas aeruginosa, are the most intensively studied biosurfactants. This arises from two contrasting facts. First, they display relatively high surface activities and are produced in relatively high yields after relatively short incubation periods by a well-understood, easy to cultivate microorganism. Second, they are one of the virulence factors contributing to the pathogenesis of P. aeruginosa infections, and consequently, many aspects of RL biosynthesis have been investigated, in part, to control their production and effects. The discovery of RLs dates back to 1946 when Bergstr€om et al. reported an oily glycolipid produced by Pseudomonas pyocyanea (now P. aeruginosa) grown on glucose. This substance was named pyolipic acid and its structural units were identified as L-rhamnose and b-hydroxydecanoic acid (Bergstr€om et al. 1946a, b; Hauser and Karnovsky 1954; Jarvis and Johnson 1949). The exact chemical nature of these biomolecules was unraveled by Jarvis and Johnson (1949) followed by Edwards and Hayashi (1965). Since then, extensive investigations have been conducted covering various aspects of RL research. Numerous research teams have contributed to decipher the biosynthetic pathway of RLs (Burger et al. 1963; De´ziel et al. 2003; Hauser and Karnovsky 1957, 1958; Rehm et al. 2001; Sobero´n-Cha´vez 2004; Zhu and Rock 2008). This was done in conjunction with efforts to identify the genes responsible for RL production, both at the enzymatic (Ochsner et al. 1994a; Rahim et al. 2000, 2001; Rehm et al. 2001; Zhu and Rock 2008) and regulatory (Ochsner et al. 1994b; Pearson et al. 1997; Pesci et al. 1997) levels. These advancements were made possible largely because of the major efforts conducted on the development of versatile and accurate methods for RL detection and analysis (De´ziel et al. 2000; Gartshore et al. 2000; Heyd et al. 2008; MataSandoval et al. 1999; Price et al. 2009; Rendell et al. 1990; Schenk et al. 1995; Siegmund and Wagner 1991). These investigations revealed a large diversity of RL congeners and homologs produced by various P. aeruginosa strains under many different culture conditions and also from other bacterial species (Abdel-Mawgoud et al. 2010; Dubeau et al. 2009; Ochsner et al. 1994a, b; Van Gennip et al. 2009). Another line of research is devoted to understanding the role of these biomolecules for the producing microorganisms as well as their interactions with other biological systems, especially the human body (Abdel-Mawgoud et al. 2010; Van Hamme et al. 2006). One of these roles is to promote the uptake of poorly soluble

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

15

hydrocarbons (Koch et al. 1991). Other physiological functions include the control of the bacterial cell surface hydrophobicity for attachment and detachment on different substrates (Al-Tahhan et al. 2000; Arino et al. 1998a; Sotirova et al. 2009; Yuan et al. 2007; Zhong et al. 2007, 2008), and the enhancement and modulation of surface motility (Caiazza et al. 2005; De´ziel et al. 2003; K€ohler et al. 2000; Tremblay et al. 2007). Studies about the interactions of RL with other biological systems are numerous. The antibacterial (Abalos et al. 2001; Bergstr€ om et al. 1946b; Haba et al. 2003b; Lang et al. 1989; Onbasli and Aslim 2008; Shen et al. 2009; Sotirova et al. 2008; Yilmaz and Sidal 2005), antifungal (Kim et al. 2000; Yoo et al. 2005), antiviral (Cosson et al. 2002; Remichkova et al. 2008), antiphytopathogenic (De Jonghe et al. 2005; Haferburg et al. 1987; Kim et al. 2000; Nielsen et al. 2005, 2006), and algicidal (Wang et al. 2005) properties of RLs have been extensively investigated. RLs released by P. aeruginosa have long been known as the heat-stable extracellular hemolysin (Fujita et al. 1988; Johnson and Boese-Marrazzo 1980; Kurioka and Liu 1967; Sierra 1960) and more recently, a RL congener produced by Burkholderia pseudomallei was shown to display hemolytic and cytotoxic activities (H€aussler et al. 1998, 2003). Because of their excellent surface activity, the physicochemical properties of RLs have received considerable interest (Abalos et al. 2001; Abdel-Mawgoud et al. 2009; Chen 2004; Cohen and Exerowa 2007; Cohen et al. 2004; Haba et al. 2003b; Hansen et al. 2008; Ochoa-Loza et al. 2001; Ozdemir and Malayoglu 2004; Ozdemir et al. 2004; Pornsunthorntawee et al. 2009). Due to their hydrocarbonsolubilizing properties, they also have been used in the fields of bioremediation and biodegradation (Arino et al. 1998a; Asci et al. 2007, 2008; Avramova et al. 2008; Beal and Betts 2000; Benincasa 2007; Cameotra and Singh 2009; Cho et al. 2004; Churchill et al. 1995). The potential industrial and biotechnological applications of RLs are thus quite diverse (Singh et al. 2007). RLs have been used for the synthesis and stabilization of nanoparticles (Palanisamy and Raichur 2009; Xie et al. 2006), the preparation of microemulsion (Nguyen and Sabatini 2009; Xie et al. 2007), as an antiagglomeration agent (York and Firoozabadi 2008), as dispersing agent (Raichur 2007; Tripathy and Raichur 2008), in cleaning soap mixtures (Ecover™ products) and as a source of rhamnose (Linhardt et al. 1989). Clinical testing of RLs as pharmacoactive compounds has been performed. Some successful trials proved the potential applications of RLs for the treatment of ulcers (Piljac et al. 2008) and of full-thickness wounds (Stipcevic et al. 2006). These promising properties and potential application of RLs have encouraged researchers to improve the production of RLs, using industrially safe and more affordable processes in order to reduce production costs, which currently restrict the competitiveness of RLs vis-a`-vis petroleum-derived surfactants. This goal has been sought through different approaches. First, many attempts have been made to isolate RL producers other than the opportunistic pathogen P. aeruginosa (Abouseoud et al. 2008; Celik et al. 2008; Chang et al. 2005; Christova et al. 2004; Gunther et al. 2005; Rooney et al. 2009) or to transfer the genes responsible

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for RL production into more industrially safe heterologous hosts, such as Escherichia coli (Cabrera-Valladares et al. 2006; Ochsner et al. 1994a). Second, important efforts have been dedicated to the identification of low-cost and renewable raw material as production substrates, such as agroindustrial wastes (Nitschke et al. 2005; Rahman et al. 2002). Finally, an even production of RLs through pure chemical synthesis was also reported (Bauer et al. 2006; Duynstee et al. 1998; Howe et al. 2006). As highlighted in this brief overview, it would be difficult to present all domains and aspects of RL research in one chapter. Therefore, our aim here is to provide a description of the chemical nature of RLs and to mention the different methods of RL detection and analysis. This chapter aims also to discuss the biosynthetic pathways of different RL precursors and describe the network of genetic regulation controlling their biosynthesis. Finally, different modalities of fermentative production of RLs on large scale are described, with an account of the associated problems and approaches to overcome them.

2 Rhamnolipid Structure, Detection, and Analysis RLs are among the best studied biosurfactants. As several methods for their detection and analysis have been developed, their structure and characteristics are largely known.

2.1

Structure

RLs are glycosides that are composed, of a glycon part and an aglycon part linked to each other via O-glycosidic linkage. The glycon part is composed of one (for mono-RLs) or two (for di-RLs) rhamnose moieties linked to each other through a-1,2-glycosidic linkage (Edwards and Hayashi 1965). The 2-hydroxyl group of the distal (relative to the glycosidic bond) rhamnose group remains generally free, although in some rare homologs it can be acylated with a long chain alkenoic acid (Yamaguchi et al. 1976). The aglycon part, however, is composed of mainly one or two [in few cases, three (Andr€a et al. 2006)] b-hydroxyfatty acid chains. These fatty acid chains are most commonly saturated or, less abundantly, mono- or polyunsaturated. Their chain lengths vary from C8 to C16 (Abalos et al. 2001; De´ziel et al. 1999a, 2000). These fatty acid chains are linked to each other through an ester bond formed between the b-hydroxyl group of the distal (relative the sugar part) chain with the carboxyl group of the proximal chain (Fig. 1). In most cases, the carboxyl group of the distal b-hydroxyfatty acid chain remains free. However, few homologs have this group esterified with a short alkyl group (Hirayama and Kato 1982). Figure 1 displays the structure of the best known RL congener, a-L-rhamnopyranosyl-a-L-

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

17

OH

O

O H3C

O

O

O CH3

HO HO O H3C

CH3

O

HO HO OH

Fig. 1 Chemical structure of the first identified rhamnolipid; known as a L rhamnopyranosyl a L rhamnopyranosyl b hydroxydecanoyl b hydroxydecanoate (Rha Rha C10 C10). Its full IUPAC name is (R) 3 {(R) 3 [2 O (a L rhamnopyranosyl) a L rhamnopyranosyl]oxydecanoyl} oxydecanoate; Or the synonym name: (R) 3 ((R) 3 ((2R,3R,4R,5R,6S) 4,5 dihydroxy 6 methyl 3 ((2S,3R,4R,5R,6S) 3,4,5 trihydroxy 6 methyltetrahydro 2H pyran 2 yloxy)tetrahydro 2H pyran 2 yloxy)decanoyloxy)decanoic acid

rhamnopyranosyl-b-hydroxydecanoyl-b-hydroxydecanoate, which is typically symbolized as Rha-Rha-C10-C10. The stereochemical configuration of the b-hydroxy groups of the fatty acid chains is in the R-configuration (Bauer et al. 2006; Schenk et al. 1997). To date, about 60 different RL congeners and homologs have been reported, as recently reviewed by Abdel-Mawgoud et al. (2010). While P. aeruginosa synthesizes a mixture of mono- and di-RLs with hydroxyacyl moieties mostly from C8 up to C12, species from the Burkholderia genus produce principally di-RLs with two rhamnose units and mainly C14 hydroxy acyl chains.

2.2

Methods of Detection and Analysis

Several methods with variable precision and purposes are available for the detection and analysis of RLs.

2.2.1

Qualitative Methods

The most widely used method for qualitative, high throughput screening of RLproducing bacterial strains is the cetyltrimethylammonium bromide (CTAB) agar test (Pinzon and Ju 2009a; Siegmund and Wagner 1991). In this method, the anionic RLs form an insoluble complex with this cationic bromide salt, and the complex is revealed using methylene blue present in the agar. The RL-producing strains are

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revealed by a dark blue halo around the colony, allowing for facile identification of the presence of RLs. Another indirect way to detect RLs is based on their hemolytic properties. This approach can be performed in solution, using an erythrocyte suspension to which the RL solution is added. After a predetermined time, the residual erythrocytes are removed by centrifugation and the hemoglobin released is measured at 540 nm (Johnson and Boese-Marrazzo 1980). Alternatively, RL production can be tested using blood agar plates on which the bacteria are directly inoculated. Formation of a halo around the colony can be indicative for the presence of RLs (Carrillo et al. 1996). However, because bacteria can secrete other hemolytic factors such as proteases, this test often leads to false positive results (Siegmund and Wagner 1991) and is less reliable than those based on the tensioactive effects of RLs, such as those described below. The drop collapsing test (Jain et al. 1991) is a sensitive method for the rapid screening of RL production by various isolates. This assay consists of applying a drop of a bacterial culture supernatant to be tested over a polystyrene plate containing shallow wells covered with oil. The droplet will spread over the oil only if the culture supernatant sample contains RLs. A similar approach is used in the oil spreading test, in which 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 duNouy-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 which the surface tension starts to increase and provides the Critical Micelle Concentration (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.

2.2.2

Quantitative Methods

The quantification of RLs can be performed through different strategies.

Spectrophotometric Methods One of the most widely used methods for RL quantification is the orcinol test. It consists of heating the solvent-obtained extracts of culture supernatants in the

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

19

presence of sulfuric acid and orcinol (1,3-dihydroxy-5-methylbenzene). The rhamnose groups of RL are hydrolyzed and transformed into methyl furfural, which then reacts with the orcinol to produce a blue-green color that can be measured spectrophotometrically at 421 nm (Chandrasekaran and BeMiller 1980; Koch et al. 1991). A standard curve is prepared with rhamnose, or preferably with a standard RL mixture, for quantification. When rhamnose is used for building up the calibration curve, a correction factor must be applied to compensate for the extra mass of the lipidic portion of RLs. De´ziel et al. (2000) calculated a correction factor of 2.25. One problem with this approach is that the results will vary with the proportion of mono- to di-RLs in the culture to be analyzed. A variation of the orcinol test uses anthrone (9,10-dihydro-9-oxoanthracene) instead of orcinol to create a dye that can be quantified at 625 nm (Helbert and Brown 1957; Hodge and Hofreiter 1962). A quantitative method based on the interaction of methylene blue, CTAB, and RLs, as illustrated in the CTAB agar test, was described recently. It involves extracting the RLs in chloroform to which is added the two other chemicals, and the complex formed is detected at 638 nm (Pinzon and Ju 2009b). Chromatographic Methods The different approaches for RL measurement based on chromatographic procedures are presented. Thin Layer Chromatography One of the problems of RL quantification is that these compounds are produced as complex mixtures of congeners (see below), 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 nonpolar 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. One such method is thin-layer chromatography (TLC). In normal phase, the polar stationary phase (silica gel) is eluted with a relatively polar mobile phase, for instance, chloroform:methanol:20% aqueous acetic acid (65:15:2) (Koch et al. 1991). This allows for straightforward separation of mono- from the more polar and later eluting di-RLs. Alternatively, a reverse phase TLC method has been developed in which the stationary phase is a hydrophobic C8 matrix eluted with methanol:water:trifluoroacetic acid (90:10:0.25) (deKoster et al. 1994). With this approach, RLs congeners are separated according to the length of their alkyl chains. Once the separation is completed, RLs can be visualized using the orcinol test (Koch et al. 1991), with reagents specific for sugars or fatty acids or with reagents that are used to reveal most organic compounds on TLC such as the “ceric dip” (Mechaly et al. 1997). Densitometric analysis of the revealed spots can be performed for more quantitative data (Matsufuji

20

A. Mohammad Abdel Mawgoud et al.

et al. 1997), but this approach 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. Gas Chromatography Because of their relatively high molecular weights, RLs cannot be directly analyzed by gas chromatography (GC). Typically, prior to analysis, RLs are thus hydrolyzed with acid or with a strong base, their acid groups are modified into methyl esters (Van Dyke et al. 1993), and optionally, the hydroxyl groups are further transformed into a trimethylsilyl (TMS) ether (Arino et al. 1996). Rhamnose can be analyzed by GC as a TMS derivative (Arino et al. 1996). The various 3-hydroxyfatty acids are then identified and quantified using flame ionization detection (FID) or mass spectrometry (MS), using standards to determine their retention times and response factors. The main problem with GC analysis is that the relationship between the 3-hydroxyfatty acids in the dilipid portion of the RLs is lost, along with the relationship between the dilipids and their substitution with one (or two) rhamnose moieties. 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. However, because they only absorb UV at very short wavelengths, RL detection is problematic. One approach is to derivatize them with para-bromoacetophenone in order to produce the corresponding para-bromophenacyl esters, which can be detected at 265 nm (Schenk et al. 1995). Alternatively, an Evaporative Light Scattering (ELS) detector, which rapidly evaporates the solvent and monitors the diffraction of a beam of light by the analyte, has been used on occasions (Arino et al. 1996; Noordman et al. 2000). With either type of detection, 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. 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 (Benincasa et al. 2004; De´ziel et al. 1999b, 2000; Haba et al. 2003a; Monteiro et al. 2007). In negative ESI, the molecular weight of the

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

21

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. 1999b, 2000). Fragmentation of the pseudomolecular ion using MS/MS analysis of the parent ion can provide further structural information, if required. For example, this approach allows the discrimination of Rha-C10-C8 from the isomeric Rha-C8-C10, even though they are not chromatographically resolved (De´ziel et al. 1999b). The ability to predict the compound eluting at a given retention time permits quantification, even if the corresponding compound is not available as a pure standard. The response factor of mono-RLs differs from that of the di-RL congeners (De´ziel et al. 1999b). But within the same family, the molar response factor is very similar, thus allowing for quantification of all the members of such a family of congeners if one member of the family can be purified. Quantification of a given congener can be performed by integrating the intensity of the peak occurring at the correct retention time in the ion chromatograph of the corresponding pseudomolecular ion. Another alternative is to perform a MS/MS experiment in which a given pseudomolecular ion is fragmented and only one of its fragments is monitored. This approach, called Multiple Reaction Monitoring (MRM), increases the signal-tonoise ratio of the analysis, thus providing a lower limit of detection. To perform quantification of RLs, an internal standard, such as 16-hydroxyhexadecanoic acid (De´ziel et al. 2000), is added to compensate for differences in the ionization efficiencies from sample to sample. 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. 2003a; Monteiro et al. 2007) rather than for quantification of complex RL congener mixtures.

3 Biosynthesis and Genetic Regulation Details of the pathways involved in RL biosynthesis, including synthesis of the fatty acid and sugar moieties, have been in large part elucidated. Furthermore, a good deal of information is available on the regulation of genes important for RL production.

22

3.1

A. Mohammad Abdel Mawgoud et al.

Biosynthesis of Rhamnolipids

Following early studies to understand the metabolic pathway of RL biosynthesis (Hauser and Karnovsky 1957, 1958), Burger et al. (1963) reported a putative mechanism of rhamnosylation of fatty acid chains to form RLs according, as an example, to the following reactions for Rha-Rha-C10-C10, (Burger et al. 1963): 2b
Hydroxydecanoyl
CoA ! b
hydroxydecanoyl
b
hydroxydecanoate þ2 CoA
SH TDP
l
rhamnoseþb
hydroxydecanoyl
b
hydroxydecanoate ! TDPþl
rhamnosyl
b
hydroxydecanoyl
b
hydroxydecanoate TDP
l
rhamnoseþl
rhamnosyl
b
hydroxydecanoyl
b
hydroxydecanoate ! TDPþl
rhamnosyl
l
rhamnosyl

(1)

(2)

(3)


b
hydroxydecanoyl
b
hydroxydecanoate The first reaction involves dimerization of two b-hydroxydecanoic acid chains. The dimer then undergoes two sequential rhamnosylation reactions with two different rhamnosyltransferases: rhamnosyltransferase 1 (Rt-1) in reaction (2) and rhamnosyltransferase 2 (Rt-2) in reaction (3) (Burger et al. 1963). It was initially hypothesized that the biosynthesis of biosurfactants, in general, and especially of glycolipids, proceeds through one of the three possible pathways: Both moieties are synthesized independently of the growth substrate (de novo). With a hydrophobic carbon source such as fatty acids and triglycerides, the lipid moieties are directly derived from the carbon source, but the sugar is synthesized de novo. The sugar moiety is directly derived from the carbon source, but the lipid component is synthesized de novo. The biosynthesis of RLs has been largely elucidated. It is present here under three sections, namely: biosynthesis of the lipid moiety, biosynthesis of the sugar moiety, and finally, the enzymatic dimerization and the rhamnosyl transfers, which yield the final products.

3.1.1

Biosynthesis of the Lipid Moiety of Rhamnolipids

A number of reports demonstrate that the biosynthesis of the lipid components of RLs proceeds through the classical pathway of fatty acid synthesis from 2-carbon units. First, Hauser and Karnovsky (1957) reported in vivo experiments where the lipidic component of RLs incorporated radioactivity from various labeled

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

23

precursors supplied to the cultures, such as 14C-acetate and 14C-glycerol (Hauser and Karnovsky 1957). Second, the lipidic portion of RL is not substantially altered if different carbon sources are used (Hommel and Ratledge 1993). The observation that the stereochemistry of the b-hydroxyacids in the lipidic portion of RLs matches that of the intermediates in fatty acid biosynthesis, as opposed to that of the intermediates in fatty acid b-oxidation, suggests that fatty acid synthesis is the source for this moiety of RLs (Zhu and Rock 2008). The lipidic moiety of RLs is thus most probably synthesized de novo (Fig. 2). The classical fatty acid synthetases of type-II (FAS II), which are found in most bacteria and plants, differ from those common to mammals (FAS I), in that these former fatty acid synthetases are not a single multifunctional polypeptide produced from a single gene (White et al. 2005). They are instead part of a dissociated fatty acid synthetase system, in which the individual reactions are catalyzed by separate proteins that are encoded by separate genes (Hoang and Schweizer 1997; White et al. 2005). The reference system for FAS II biochemistry is E. coli (White et al. 2005). In general, however, these enzymes are homologous among bacteria. For example, many isozymes of FabI, an enoyl-acyl carrier protein reductase, have been found in other bacteria such as FabL, FabV, and FabK (Zhu et al. 2009). Yet, the similarities and differences between homologous enzymes in different bacteria remain to be identified (White et al. 2005). In E. coli, fatty acid biosynthesis can be separated into two stages, initiation and cyclic elongation. Each round of elongation requires four chemical reactions (Hoang and Schweizer 1997). Three b-ketoacyl-acyl carrier protein (ACP) synthetases, KAS I (FabB), KAS II (FabF), and KAS III (FabH), the products of fabB, fabF, and fabH, play pivotal roles in fatty acid synthesis. Initiation requires malonyl coenzyme A (CoA) and malonyl-ACP. Malonyl-CoA is synthesized by acetyl-CoA carboxylase, and malonyl-ACP is derived from malonyl-CoA and ACP by the action of malonyl-CoA:ACP transacylase, the product of fabD. The first cycle of elongation is initiated by KAS III (FabH), which condenses malonyl-ACP to acetylCoA. Subsequent cycles are initiated by condensation of malonyl-ACP with acylACP, catalyzed by KAS I (FabB) and KAS II (FabF). In the second step, the resulting b-ketoester is reduced to a b-hydroxyacyl-ACP by a single, NADPHdependent b-ketoacyl-ACP reductase (FabG). The third step in the cycle is catalyzed by either the fabA- or the fabZ-encoded b-hydroxyacyl-ACP dehydratase. The fourth and final step in each cycle involves the conversion of trans-2-enoylACP to acyl-ACP, a reaction catalyzed by a single NADH-dependent enoyl-ACP reductase (FabI) (Hoang and Schweizer 1997) (Fig. 2). For P. aeruginosa, a model of FAS II was proposed that is composed of the same enzymatic machinery (Hoang and Schweizer 1999). However, the fabH homolog in P. aeruginosa has been only tentatively identified. It is therefore only hypothesized that initiation of FAS II is mediated by KAS III (FabH), and subsequently, initiation of elongation cycles are catalyzed by KAS I (FabB) for saturated fatty acid substrates and KAS II (FabF) for unsaturated fatty acid substrates (Hoang and Schweizer 1999). The same team has also characterized FabA, FabB, and FabI proteins in P. aeruginosa (Hoang and Schweizer 1997, 1999).

CoA-SH

TDP

Monorhamnolipids

TDP

HHQs

PqsABCD ?

Anthranilic acid

RhlG

RhlB

PHAMCL

Acetyl-CoA Carboxylase

RhlC

dTDP-L-rhamnose

RmlD

dTDP-4-oxo-6-deoxy-L-mannose

RmlC

dTDP-4-oxo-6-deoxy-D-glucose

RmlB

dTDP-D-glucose

RmlA

D-glucose-1-P

LasI

2 ACP-SH

RhlA

PhaC

5-MTA

SAM

CO2+ACP

CoA-SH

Malonyl-ACP

FabH

FabG

RhlI

SAM

H2O

FabI

NADH

+

NADH + H

trans-2-enoylACP

FabA/Z

5-MTA

Acyl-ACP

FAS II

β-hydroxyacylACP

N-butanoyl-L-HSL

FabB, F

β-ketoacylACP

+

CoA-SH

NADP NADPH + + H

ACP-SH

PhaG

AcetoacetylACP

FabD

ACP-SH

Malonyl-CoA

Acetyl-CoA

CoA-SH + CO2

β-hydroxyacylCoA

N-(3-oxo-dodecanoyl)-L-HSL

HAA

HAA

CoA-SH

β-oxidation

+

Fig. 2 Metabolic pathway of rhamnolipid biosynthesis and its relation to the biosynthesis of polyhydroxyalkanoates (PHA) and quorum sensing signaling molecules. AlgC: phosphomannomutase, RmlA: glucose-1-phosphate thymidylyltransferase, RmlB: dTDP-D-glucose 4,6-dehydratase, RmlC: dTDP-4-dehydrorhamnose 3,5-epimerase, RmlD: dTDP-4-dehydrorhamnose reductase, FabD: malonyl-CoA:ACP transacylase, FabH, FabB and FabF: b-ketoacyl-ACP synthetases, FabG: NADPH-dependent b-ketoacyl-ACP reductase, FabA, FabZ: b-hydroxyacyl-ACP dehydratases, FabI: NADH-dependent enoyl-ACP reductase, HAA: 3-(3-hydroxyalkanoyloxy)alkanoic acid, PqsABCD: 4-hydroxy-2-heptylquinoline synthesis cluster, RhlA: 3-(3-hydroxyalkanoyloxy)alkanoate synthetase, RhlB: rhamnosyltransferase 1, RhlC: rhamnosyltransferase 2, RhlI: N-butanoyl-L-HSL synthase, RhlG: b-ketoacyl reductase, PhaG: (R)-3hydroxydecanoyl-ACP:CoA transacylase, PhaC: PHA synthase, LasI: N-(3-oxo-dodecanoyl)-L-HSL synthase, HHQ: 4-hydroxy-2-heptylquinoline, PHAMCL: Poly(3-hydroxyalkanoates) of medium chain length C6–C14, SAM: S-adenosylmethionine, 5-MTA: 50 -methylthioadenosine

Kreb's Cycle

Acetyl-CoA

Pyruvate

AlgC

Dirhamnolipids

Glyceraldehyde-3-P

Fructose-1,6-P2

Fructose-6-P

D-Glucose-6-P

Glucose

24 A. Mohammad Abdel Mawgoud et al.

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

25

Interestingly, ACP intermediates in the FAS II pathway are suggested to be contributing to the enormous diversity of bacterial products because they are diffusible entities that can be diverted into other biosynthetic pathways (White et al. 2005) like RL biosynthesis (De´ziel et al. 2003; Rehm et al. 2001; Zhu and Rock 2008), polyhydroxyalkanoates (Pham et al. 2004; Rehm et al. 2001), as well as the biosynthesis of quorum sensing signal molecules (Bredenbruch et al. 2005). The exact link between FAS II intermediates and RL biosynthesis, however, is still a matter of debate. Recently, Zhu and Rock (2008) proved that RhlA directly utilizes b-hydroxydecanoyl-ACP intermediates to generate 3-(30 -hydroxydecanoyloxy) decanoic acid, the ten carbon member of the 3-(30 -hydroxyalkanoyloxy)alkanoic acids (HAAs) portion of RLs. This had been previously suggested (De´ziel et al. 2003), based on the amino acid homology of the rhlA gene product with PhaG, a 3-hydroxyacyl ACP:CoA transacylase identified in various species of Pseudomonas (De´ziel et al. 2003; Rehm et al. 1998). However, these findings contradict previous data suggesting that an enzyme called RhlG is responsible for diverting fatty acid synthesis intermediates into the RL biosynthetic pathway in P. aeruginosa, based on its similarity to FabG (Campos-Garcia et al. 1998). However, the work of Zhu and Rock (2008) indicates that there is no enzyme upstream of RhlA for diverting b-hydroxy fatty acid intermediates from FAS II cycle, and that RhlA acts more like as a molecular ruler that preferentially diverts appropriate intermediates from FASII for the synthesis of the HAA moiety of RLs. This finding was based on the biochemical properties of the purified RhlA protein and its products when heterologously expressed in an E. coli host (Zhu and Rock 2008). That RhlG has no role in picking up b-hydroxydecanoyl-ACP for HAA synthesis was further supported by the fact that, although the overall structures of the RhlG-NADP+ and FabG-NADP+ complexes are indeed similar, there are key differences related to their function, making RhlG 2,000-fold less active than FabG in carrying out the same reaction. These findings entail that RhlG is indeed a NADPH-dependent b-ketoacyl reductase, but its substrate is not carried by the ACP of fatty acid synthesis (Miller et al. 2006). Another important issue is whether the substrate of the enzyme dimerizing b-hydroxydecanoyl moieties into HAA is carried by an ACP or by CoA. An earlier experiment performed in vitro with crude cellular extract showed that b-hydroxydecanoyl-CoA is a precursor of HAA (Burger et al. 1963). However, this experiment did not exclude the possibility that the reaction could involve one bhydroxydecanoyl-CoA and free b-hydroxydecanoyl acid (Burger et al. 1963). A recent experiment performed in vitro with purified RhlA showed that HAAs are produced when b-hydroxydecanoyl-ACP is used as substrate, while no HAAs are obtained when b-hydroxydecanoyl-CoA is used at concentrations up to ten times higher than those of b-hydroxydecanoyl-ACP (Zhu and Rock 2008). Although these results seem in contrast with those of Burger et al. (1963), it remains to be seen whether b-hydroxydecanoyl-CoA could be a precursor of HAAs in vivo. Remains also to be elucidated is the biosynthesis of some RLs having only one 3-hydroxy-fatty acid instead of the dimmer, attached to one or two rhamnose(s).

26

A. Mohammad Abdel Mawgoud et al.

Are they degradation products of other RLs or are they formed by direct rhamnosylation of a 3-hydroxy-fatty acid by RhlB and RhlC? The biosynthetic pathway of RLs having one more unsaturation in their HAA chains will also require further investigation. Biosynthetic Link Between Rhamnolipid-Lipid Moiety and PHA As many other Pseudomonas sp, P. aeruginosa is capable of accumulating poly (3-hydroxyalkanoates) (PHAs) granules containing medium chain length (MCL) (C6 to C14) 3-hydroxyfatty acids (PHAMCL) (Madison and Huisman 1999; Rehm et al. 2001). The biosynthetic pathway of PHAMCL proceeds mainly through FASII, when grown on carbon sources metabolized into acetyl-CoA like carbohydrates. On the other hand, when grown on hydrocarbons, it proceeds mainly through b-oxidation (Madison and Huisman 1999; Rehm et al. 1998, 2001; Timm and Steinbuchel 1990). The substrate of the PHAMCL synthases is (R)-3hydroxydecanoyl-CoA, which is formed from ACP-thioester precursors by the action of the transacylase PhaG (Rehm et al. 2001). Thus, PhaG directly links fatty acid de novo biosynthesis to PHA biosynthesis (Rehm et al. 2001). Based on that, it is understood that RLs and PHAMCL biosynthesis compete with each other for the b- hydroxydecanoyl-ACP precursor, which is an intermediate in FAS-II (Rehm et al. 2001) (Fig. 2). Although that PHA synthase could be responsible for supplying the HAA moieties for RL synthesis has been postulated (CamposGarcia et al. 1998), another study proved that PHA-synthase negative mutants are still capable of RL production (Pham et al. 2004).

Biosynthetic Link Between the Rhamnolipids Lipidic Moiety and Quorum Sensing Signal Molecules P. aeruginosa produces two classes of signal molecules, the acyl homoserine lactones (AHLs) and the 4-hydroxy-2-alkylquinolines (HAQs). The most abundant AHLs are N-(3-oxododecanoyl)-L-HSL and N-butanoyl-L-HSL, while the HAQs include 3,4-dihydroxy-2-heptylquinoline [Pseudomonas Quinolone Signal (PQS)] and its precursor 4-hydroxy-2-heptylquinoline (HHQ) (De´ziel et al. 2004). Some of these molecules are involved in the regulation of RL synthesis genes expression. The biosynthesis of these signal molecules also requires substrates derived from FAS II by diverting FAS intermediates of a specific fatty acid chain length for their own synthesis (Schaefer et al. 1996). Using in vitro and in vivo experiments, Hoang and Schweizer (1999) showed that butanoyl-ACP serves as substrate for N-butanoyl-L-HSL biosynthesis. They observed that FabI (the enzyme that supplies acyl-ACP like butanoyl-ACP in FAS II) plays a central role in AHL biosynthesis in vivo because a fabI mutant of P. aeruginosa produced only 50% of the AHL levels found in wild-type cells (Hoang and Schweizer 1999). Moreover, when coupled to FabI, purified P. aeruginosa N-butanoyl-L-HSL synthase (RhlI) produced N-butanoyl-L-HSL from crotonyl-ACP

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

27

(the enoyl-ACP precursor of butanoyl-ACP in FAS II) and S-adenosylmethionine (SAM) (Hoang and Schweizer 1999) (Fig. 2). Similarly, N-(3-oxododecanoyl)-L-HSL is synthesized from the b-ketododecanoyl-ACP intermediate of FAS II by the action of N-(3-oxododecanoyl)-L-HSL synthase (LasI) in the presence of SAM (Fuqua and Greenberg 2002; Hoang and Schweizer 1999; Schaefer et al. 1996) (Fig. 2). That HAQs also obtain some of their biosynthetic precursors from the FAS II cycle was hypothesized (Sobero´n-Cha´vez et al. 2005). Ritter and Luckner (1971) and Calfee et al. (2001) proposed a synthetic scheme for HAQs where anthranilate and b-ketodecanoic acid are condensed in a multistep reaction that would produce HHQ, followed by PQS after the release of a one carbon unit as CO2. This pathway has been verified using labeled substrates (Bredenbruch et al. 2005; De´ziel et al. 2004). These reactions are catalyzed by the pqsABCD and pqsH gene products (Dubern and Diggle 2008). Bredenbruch et al. (2005) suggested that the b-ketoacyl reductase RhlG plays a role, although an intact rhlG gene is not required for the production of HAQs. A more detailed description of the biosynthetic pathway of HAQs was recently described (Gross and Loper 2009).

3.1.2

Biosynthesis of Rhamnolipids-Rhamnose Moiety

Rhamnose is a component of the cell wall lipopolysaccharide (LPS) core and of several O-antigen polysaccharides in a variety of gram-negative bacteria, including several strains of Pseudomonas (Burger et al. 1963; Rahim et al. 2000). Early studies on the catabolic pathway of rhamnose were performed using radioactive carbon sources. These showed that the carbons of rhamnose are derived from glycerol and not from acetate, apparently through the condensation of two three-carbon units formed from glycerol without cleavage or rearrangement of its carbon carbon bonds (Hauser and Karnovsky 1957). In a later study (Hauser and Karnovsky 1958), both glycerol and propane-1,2diol were found to provide carbon to the rhamnose of RL and to be equally converted into the precursors of the two halves of the sugar (Hauser and Karnovsky 1958). For fructose as the carbon source, it was suggested that this sugar is cleaved into two triose units that are subsequently recombined to form rhamnose (Hauser and Karnovsky 1958). However, this latter study did not clarify the detailed steps of RL biosynthesis with glucose as the sole carbon source. The biosynthetic conversion of glucose to rhamnose in P. aeruginosa was clarified with the in vivo and in vitro studies of Southard et al. (1959) and of Glaser and Kornfeld (1961). These reports showed that glucose is converted into rhamnose without randomization of the carbon chain and that the carbon at the position 1 of glucose is found at the same position in rhamnose (Glaser and Kornfeld 1961). Glaser and Kornfeld (1961) also explained the previous results of Hauser and Karnovsky (1958) with labeled glycerol, suggesting that two three-carbon compounds initially condense to form glucose. They also showed that in order for D-glucose to be converted into L-rhamnose, the configuration of the carbon 3, 4, and 5 should be inverted and that a reduction at the carbon 6 is required. They also

28

A. Mohammad Abdel Mawgoud et al.

showed that D-glucose is converted into L-rhamnose through a 4-keto-6-deoxyglucose intermediate and they further postulated that the inversion of the configuration at the carbon 3 and 5 is performed by an isomerization reaction facilitated by enolization of the ketone (Glaser and Kornfeld 1961). Recently, the biosynthetic link between glucose and rhamnose was found to proceed through the phosphoglucomutase AlgC, as an algC mutant does not produce detectable amounts of RLs (Olvera et al. 1999). This phosphoglucomutase converts D-glucose-6-phosphate into D-glucose-1-phosphate, and this compound is then used by RmlA, RmlB, RmlC, and RmlD to produce dTDP-L-rhamnose (Fig. 2) (Olvera et al. 1999; Robertson et al. 1994). dTDP-L-rhamnose is also the precursor for the L-rhamnose present in the outer core oligosaccharide of the lipopolysaccharide (LPS) (Rahim et al. 2000). Furthermore, dTDP-L-rhamnose was recently shown to provide the L-rhamnose as a component of both the flagellin glycan of b-type flagellin (Lindhout et al. 2009) and the psl-encoded polysaccharide, which consists of a repeating pentasaccharide containing D-mannose, D-glucose, and L-rhamnose (Byrd et al. 2009). This pathway only explains how L-rhamnose is probably synthesized when the bacteria are grown with glucose as the carbon source. However, many other carbon sources are more efficient for RL production, such as mannitol (De´ziel et al. 1999b), vegetable oils (Trummler et al. 2003), glycerol, or ethanol (Chen et al. 2007a). Under these conditions, the exact biosynthetic pathways remain to be elucidated.

3.1.3

Three Last Enzymatic Reactions in Rhamnolipids Biosynthesis

As mentioned earlier, three enzymatic reactions are required in the final steps of RL biosynthesis in P. aeruginosa (Sobero´n-Cha´vez et al. 2005): (1) RhlA is involved in the synthesis of the HAAs, the fatty acid dimers, from two 3-hydroxyfatty acid precursors (De´ziel et al. 2003; Le´pine et al. 2002; Zhu and Rock 2008); (2) the membrane-bound RhlB rhamnosyltransferase uses dTDP-L-rhamnose and an HAA molecule as precursors, yielding mono-RL; (3) these mono-RLs are in turn the substrates, together with dTDP-L-rhamnose, of the RhlC rhamnosyltransferase to produce di-RLs. Unfortunately, few works have characterized these three enzymes (Fig. 2). Burger et al. (1966) were unable to characterize RhlA because it was relatively labile under their conditions, thus preventing purification. Nonetheless, they showed that b-hydroxydecanoyl-CoA is a precursor of HAAs. Ochsner et al. (1994a) were also unable to purify RhlA and suggested that it is involved in the synthesis or supply of the precursors of subsequent rhamnosyl transferase or that it is necessary for the stabilization or anchoring of RhlB in the cytoplasmic membrane. Based on the analysis of the amino acid sequences derived from the nucleotide sequence of rhlA, they deduced that RhlA is 32.5 kDa protein that harbors a putative signal sequence, suggesting that it is located in the periplasm (Ochsner et al. 1994a). De´ziel et al. (2003) later proposed that RhlA is responsible for the formation of HAAs, based on the observation that an rhlB mutant produce HAAs

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

29

but no RLs while a rhlA mutant does not produce neither HAAs nor RLs. Recently however, Zhu and Rock (2008) purified and characterized RhlA. They found that RhlA formed one molecule of b-hydroxydecanoyl-b-hydroxydecanoate (HAA) from two molecules of b-hydroxydecanoyl-ACP. They also showed that RhlA has a greater affinity for ten-carbon substrates (Zhu and Rock 2008). From these findings, they conclude that RhlA uses b-hydroxyacyl-ACP selectively picked from FAS-II cycle. In contrast to the findings of Burger et al. (1966), they did not detect any HAA production using b-hydroxyacyl-CoA, even at a concentration ten times higher than the one used with b-hydroxyacyl-ACP (Zhu and Rock 2008). It remains to be seen whether RhlA can use both substrates in vivo (Rehm et al. 2001), but maybe with different levels of affinity. Burger et al. (1966) purified RhlB, the enzyme catalyzing the second reaction, and partially purified RhlC, the enzyme catalyzing the third reaction. They showed that they both accepted L-rhamnosyl-b-hydroxydecanoyl-b-hydroxydecanoate, b-hydroxydecanoyl-b-hydroxydecanoate, and b-Hydroxydecanoyl-CoA as glycosyl acceptors, while free b-hydroxydecanoate was not a substrate (Burger et al. 1966). Ochsner et al. (1994a) later described RhlB, based on the analysis of the amino acid sequences deduced from rhlB as a protein with at least two putative membrane-spanning domains, which would allow anchoring in the inner membrane. They also partially purified RhlB from the membrane fraction and determined its size to be around 47 kDa (Ochsner et al. 1994a). Rahim et al. (2001) were the first to identify rhlC, which encodes for the second rhamnosyl transferase. They found that RhlC contains a transmembrane hydrophobic region, suggesting that it is also an inner membrane bound protein. RhlC specifically converts mono-RL into di-RL (Rahim et al. 2001). They suggested that both mono- and di-RLs are synthesized at the cytoplasmic side of the inner membrane before being transported to the extracellular milieu (Rahim et al. 2001). Finally, it should be mentioned that the rhlA, rhlB, and rhlC genes were recently identified in the RL-producing species Burkholderia thailandensis and B. pseudomallei (Dubeau et al. 2009). Interestingly, and in contrast with P. aeruginosa, where rhlC is separate from rhlAB, in these species, they are grouped together in a gene cluster. Furthermore, this RL synthesis gene cluster is duplicated on the chromosome of these bacteria, and both copies are functional and contribute to RL production (Dubeau et al. 2009).

3.2

Regulation of Rhamnolipid Biosynthesis

The control of RL production is complex, since it is influenced by numerous factors at both genetic control and environmental/nutritional levels. As typical secondary metabolites, biosynthesis primarily occurs from the end of the logarithmic or the onset of the stationary growth phases. Two factors concur to explain this: cell density-dependent regulation and limitation of specific nutrients.

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3.2.1

A. Mohammad Abdel Mawgoud et al.

Genetic Regulation of Rhamnosyltransferases

At the genetic level, the foundations of our current understanding were established by Urs Ochsner and colleagues in the mid-1990s (Ochsner et al. 1994a, b; Ochsner and Reiser 1995). Using a strategy of random transposon mutagenesis and genetic complementation, they identified the primary biosynthetic and regulatory genes, grouped in an rhl gene cluster, responsible for the production of RLs (Fig. 3). RhlA and RhlB are encoded by genes organized in an operon, which is flanked by the regulatory genes rhlR and rhlI. The main finding was that the expression of rhlAB is positively controlled in a cell-density manner by a cell-to-cell communication system called quorum sensing (Ochsner and Reiser 1995; Pearson et al. 1997). The production of secondary metabolites and virulence factors such as antibiotics and proteases is often controlled by quorum sensing (Miller and Bassler 2001). Gene regulation by quorum sensing implicates that bacteria produce and release chemical signal molecules for which increases in their external concentrations mirrors the cell-population density. Bacteria detect their accumulation and, once a minimal threshold stimulatory concentration is reached, they respond and alter gene expression, and therefore the behavior of the whole population (Fuqua et al. 1994). Gram-negative bacteria typically carry at least one quorum sensing mechanism mediated by a regulator of the LuxR-type and by an AHL synthase of the LuxI-type (Lazdunski et al. 2004). P. aeruginosa regulates the transcription of an array of genes by quorum sensing. A large proportion of these are directing the production of virulence factors, including proteases, lectins, HCN, phenazines, and RLs (Bjarnsholt and Givskov 2007; Williams and Ca´mara 2009). In the case of RL biosynthesis, the product of RhlI is the signal butanoyl-homoserine lactone, C4-HSL, which acts as the activating ligand of the transcriptional regulator RhlR (Fig. 3). The RhlR/C4-HSL complex then binds to a specific sequence in the rhlAB regulatory region to activate the transcription. Interestingly, RhlR was suggested to act as a transcriptional repressor when not bound to its signaling ligand (Medina et al. 2003c). The level of expression of rhlAB is thus dependent on the local environmental concentration of this signal. The expression of the second rhamnosyltransferase, encoded by rhlC, is coordinately regulated with rhlAB by the same quorum sensing regulatory pathway (Rahim et al. 2001). Besides RhlR/C4-HSL quorum sensing, two additional cell-to-cell communication systems participate in the quorum sensing circuitry of P. aeruginosa and influence rhlAB transcription (Fig. 3). First, the rhl system is positively upregulated by another LuxR-type regulator called LasR, which is activated by its cognate AHL N-3-oxo-dodecanoyl-HSL (3-oxo-C12-HSL) (Latifi et al. 1996; Pearson et al. 1997). Second, besides the LuxR/AHL-type circuits, P. aeruginosa carries a distinct quorum sensing system composed of the transcriptional regulator MvfR (PqsR), which directs the biosynthesis of HAQs (De´ziel et al. 2004; Gallagher et al. 2002; Pesci et al. 1999) and the activation of many quorum sensing-controlled genes via PqsE (De´ziel et al. 2005; Diggle et al. 2003; Farrow et al. 2008). Among

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31

the HAQs, HHQ and PQS act as inducing ligands of MvfR regulator (Xiao et al. 2006). This third level of quorum sensing regulation controls production of RLs by stimulating the RhlR/C4-HSL QS system through MvfR and PqsE: rhlAB expression and production of RLs are reduced in PQS-deficient and pqsE mutants (De´ziel et al. 2005; Diggle et al. 2003; Jensen et al. 2007). This is largely explained by the fact that PqsE upregulates rhlAB transcription by increasing the activity of

PtxR VqsR

N-3-oxo-dodecanoyl HSL

QscR

RsaL

N-butanoyl HSL 4-hydroxy-2-heptylquinoline 3,4-dihydroxy-2-heptylquinoline lasR

rsaL

LasR

AlgR

lasI

LasI

LasR

rhlC

PA1131

rhlA

rhlB

rhlR

rhlI

RhlR

RhlI

RhlR

PqsE

pqsH pqsA

PqsH

pqsB

pqsC

PqsABCD

pqsD

pqsE

phnA

phnB

mvfR

MvfR

Activation MvfR

MvfR

Repression

Fig. 3 Genetic regulation of rhamnolipid (RL) biosynthesis in P. aeruginosa. Multiple systems of quorum sensing (QS) participate in the control of RL synthesis genes (rhlA, rhlB, rhlC). Two QS systems, LasR/I and RhlR/I, depend on acyl homoserine lactones (AHL) ligands, N 3 oxo dodecanoyl HSL and N butanoyl HSL, respectively, which bind to their cognate transcriptional regulators, LasR and RhlR, respectively, for regulation of expression of several genes, among which are RL biosynthesis genes. LasR/I and RhlR/I activate the expression of their own auto inducer synthase genes, lasI and rhlI, respectively, as a positive feedback. The transcription of lasI and rhlI is also controlled by other regulators. RhlR/C4 HSL complex is positively regulating expression of the operon rhlAB as well as the operon encoding the rhlC gene. These last three genes encodes the three enzymes responsible for biosynthesis of RLs. LasR/oxo C12 HSL acti vates the other QS system in which the transcriptional regulator MvfR (PqsR) binds to its co inducers 4 hydroxy 2 heptylquinoline (HHQ) and 3,4 dihydroxy 2 heptylquinoline (Pseudo monas Quinolone Signal; PQS). LasR/oxo C12 HSL activates the expression of mvfR

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RhlR (Farrow et al. 2008). Thus both the LasR/3-oxo-C12-HSL and the MvfR/PQS/ PqsE quorum sensing signaling pathways end up upregulating the activity of RhlR. Accordingly, while a rhlR mutant does not produce any RLs (Ochsner and Reiser 1995), PQS system-deficient (e.g., mvfR or pqsE ) or lasR mutants still express rhlAB and produce RLs, but at reduced or delayed rates (Dekimpe and De´ziel 2009; De´ziel et al. 2005; Diggle et al. 2003; Ochsner and Reiser 1995). Additional regulatory factors modulate the expression of the rhlAB operon, all of which acting essentially on some levels of the quorum sensing global circuitry (Fig. 3). Here are some of these factors. The RsaL protein represses the transcription of lasI, inducing a large-scale downregulation of quorum sensing-regulated genes, including rhlAB (Rampioni et al. 2009). Besides the primary quorum sensing regulators LasR and RhlR, two additional LuxR-type regulators have been reported: QscR represses rhlAB transcription (Lequette et al. 2006), while VqsR activates it (Juhas et al. 2004). Additionally, VqsM, largely through modulation of vqsR expression, plays a role in regulation of QS signaling in P. aeruginosa, incidentally and indirectly upregulating rhlAB (Dong et al. 2005). Other regulators known to indirectly affect rhlAB transcription include PtxR, which negatively regulates the expression of RhlR target genes through rhlI downregulation (Carty et al. 2006) and DksA, which also seems to reduce rhlI expression, but nevertheless increases rhlAB translation (Jude et al. 2003). On a different level of quorum sensing control, the AlgR regulator was shown to repress RhlR-controlled genes, but in biofilmgrowing cells only (Morici et al. 2007); moreover, AlgR directly binds to the rhlAB promoter and prevents transcription and RLs production. At the posttranscriptional level, the production of RLs is positively controlled by the small RNA-binding protein RsmA (Heurlier et al. 2004); however, this control is indirect and the precise mechanism of control has not been identified (Brencic and Lory 2009). GidA, another factor recently reported, primarily activates RhlRcontrolled quorum sensing genes also at the posttranscriptional level, and thus controls rhlAB transcription and RL production (Gupta et al. 2009).

3.2.2

Genetic Regulation of Biosynthesis of Sugar Moiety

As presented above, the rhamnosyltransferase 1, encoded by rhlB, is responsible for catalyzing the coupling of the activated sugar dTDP-L-rhamnose to a b-hydroxyalkanoic acid dimer to yield mono-RLs (Burger et al. 1963). Together with another dTDP-L-rhamnose molecule, mono-RLs are in turn substrates of rhamnosyltransferase 2, encoded by rhlC, to produce di-RLs (Rahim et al. 2001). In P. aeruginosa, the rmlBDAC operon encodes the enzymes catalyzing the conversion of glucose-1phosphate to dTDP-L-rhamnose (Rahim et al. 2000). The regulation of rmlBDAC transcription is not characterized. There is a potential s70-like promoter sequence upstream of rmlB (Rahim et al. 2000). Interestingly, transcriptomic results suggest that this operon is upregulated by the RhlR quorum sensing pathway (Schuster et al. 2003; Wagner et al. 2003), but this has yet to be confirmed.

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

3.2.3

33

Regulation of Rhamnolipid Production by Environmental Factors – Links Between Quorum Sensing and the Environment

Restriction in the availability of a number of nutrients, except the carbon source, is known to promote the production of RLs (Guerra-Santos et al. 1986) (see below). For instance, the transcription of rhlAB and the production of RLs are inversely proportional with the concentration of iron (Fe) available to the bacterial cells (De´ziel et al. 2003; Glick et al. 2010). An explanation is provided by the wellestablished link between iron availability and quorum sensing in P. aeruginosa. Indeed, the expression of lasIR (Bollinger et al. 2001; Duan and Surette 2007; Kim et al. 2005) and rhlIR (Bredenbruch et al. 2006; Duan and Surette 2007; Jensen et al. 2006) is enhanced by Fe limitation and/or repressed by Fe supplementation. Production of RLs is inhibited by the presence of NH4+, glutamine, asparagine, and arginine as nitrogen source and promoted by NO3 , glutamate, and aspartate (K€ ohler et al. 2000; Mulligan and Gibbs 1989; Van Alst et al. 2007; Venkata Ramana and Karanth 1989). Several reports demonstrate that NO3 is the best nitrogen source for RL production (Arino et al. 1996; Manresa et al. 1991; Venkata Ramana and Karanth 1989), and it indeed elicits higher rhlAB expression than NH4+ (De´ziel et al. 2003). The basis for the preference for nitrate in RL production is unknown. On the other hand, high levels of NH4+ or glutamine reduce the production of RLs, and this is correlated with a lower glutamine synthase activity (Mulligan and Gibbs 1989). Synthesis of this enzyme, which is upregulated by environmental signals such as nitrogen-limiting conditions, is controlled by the RpoN s factor (s54) (Totten et al. 1990). In addition to the major housekeeping s factor, s70, many bacteria have alternative s factors that direct the expression of particular subsets of genes (Potvin et al. 2008). Hence, it is noteworthy that s54 is also required for production of RLs (Ochsner et al. 1994a). While it has been suggested that the rhlAB transcription start site contains a s54 promoter (Ochsner et al. 1994a; Pearson et al. 1997), this was later refuted (Medina et al. 2003c). The alternative explanation is provided by the finding that rhlR transcription is partially s54-dependent (Medina et al. 2003a). All this corroborates the frequent observation that production of RLs is increased under nitrogen-limited conditions (Mulligan and Gibbs 1989). Van Alst et al. (2007) recently contributed intriguing new observations to the connection between utilization of nitrate and production of RLs. Investigating the nitrate sensor-response regulator NarX/NarL, they found that a narL mutant strain produces significantly (approximately sixfold) more RLs than the wild type. They proposed that in the absence of its cognate response regulator NarL, NarX may activate an alternative response regulator that, either directly or indirectly, activates rhlAB (Van Alst et al. 2007). They also presented some evidence that nitric oxide, the product of nitrite reductase activity in the nitrate dissimilation pathway, activates RL production (Van Alst et al. 2007). In conclusion, the expression of the rhlAB operon and the production of RLs are regulated by both quorum sensing signals and environmental/nutritional factors. However, De´ziel et al. (2003) observed that nutritional conditions can supersede

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A. Mohammad Abdel Mawgoud et al.

cell-to-cell communication in RL production. Accordingly, exogenously added signals do not modify the onset of induction for genes controlled by both the RpoS s factor (sS) and quorum sensing (Diggle et al. 2002; Medina et al. 2003b; Schuster et al. 2003); now, the rhlAB promoter appears partially dependent on sS for its expression (Medina et al. 2003b). Indeed, high cell density and/or presence of both RhlR and its ligand signal C4-HSL do permit upregulation or advancement of rhlAB expression before late logarithmic-early stationary phase, when rpoS is induced (Medina et al. 2003b; Schuster et al. 2003). Further studies will be required to elucidate the complex interplay between nutrition-based and cell-density-based gene regulation in P. aeruginosa.

4 Bioengineering of Rhamnolipid Production 4.1

Production by P. aeruginosa

The bioproduction of RLs has been investigated almost exclusively with P. aeruginosa strains. In this section, we will present different aspects of such bioprocesses. 4.1.1

Fermentation Strategies

RLs are secondary metabolites and their production coincides with the onset of the stationary phase. Therefore, all cultivation strategies for the microbial production of RLs aim at inducing RL biosynthesis by limiting at least one medium component, for example, the nitrogen or the phosphorous source (Guerra-Santos et al. 1984; Sobero´n-Cha´vez et al. 2005). Corresponding to the growth-limited character of batch cultivations, the growth curve can satisfactorily be fitted to a logistic equation for biomass growth (4) or alternatively to a modified Gompertz equation (5) (Zwietering et al. 1990). The specific growth rate, biomass yield coefficient, and maintenance coefficients can be obtained by this approach (Ramana et al. 1991). Figure 4a illustrates the theoretical time courses of biomass and RL concentration in batch cultivation under growthlimiting conditions. The deviation of the concentrations of biodrymass and RLs based on the current biodrymass results, respectively, in the specific growth rate and specific RL production rate per cell that are shown in Fig. 4b. Actual time courses for growth and RL production in a batch cultivation were recently reported for the sequenced P. aeruginosa strain PA01 (M€ uller et al. 2010). x ¼ ½1 þ expfmmax  ðtc
tÞg xmax

1

(4)

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

5

10 15 20 25 Process time [h]

30

µ [1/h]

Biodrymass [g/l]

0

qRL, spec [g/g*h]

b Rhamnolipid [g/l]

a

35

0

5

10 15 20 25 Process time [h]

30

Fig. 4 Schematic representation of (a) the time courses of biodrymass and rhamnolipid concen tration in a batch cultivation under growth limiting conditions and (b) the specific growth rate and specific rhamnolipid production rate per cell in a batch cultivation under growth limiting conditions

99 8 == < x mmax e
  l
tÞ þ 1 ¼ exp
exp : ;; :ln x1 x1 x0 8 <

(5)

Cultivation strategies applied to RL production involve batch, fed-batch, continuous, and integrated microbial/enzymatic processes. Dextrose, glycerol, nalkanes, and triglycerides have been mostly used as carbon sources. Reported nitrogen sources include nitrate, ammonium, urea, corn steep liquor, and complex amino acids containing supplements (Lee et al. 2004; Syldatk and Wagner 1987; Zhang and Miller 1992). The reported biotechnological cultivation strategies applied to the production of RLs are: l l l

l l

(Fed-)batch cultivations under growth-limiting conditions Batch cultivations under resting cells conditions Semicontinuous productions with immobilized cells (excluding any nitrogen source) Continuous cultivations and production with free cells Solid state fermentations

Batch and Fed-Batch Strategies In general, fed-batch cultivation is the most effective process strategy for achieving high bioproductivities. This is because optimal low concentrations of all substrates can be set and the specific growth rate can be controlled by the feeding. In contrast to continuous cultures there is a limited contamination risk. However, for RL production, this strategy has not been effectively adopted yet. Even though considerable, final RL concentrations in the range of 6 95 g/L have been reported (Chen et al. 2007b; Giani et al. 1997; Hembach 1994; Lee et al. 2004; Trummler et al. 2003), these concentrations are in the same order of magnitude than those reported

36

A. Mohammad Abdel Mawgoud et al.

for batch cultivations, for which about 5 112 g/L RLs have been achieved (Chen et al. 2007b; Giani et al. 1997; Hembach 1994; Lee et al. 2004; Syldatk et al. 1985). The main difficulties for exploiting the fundamental benefits of fed-batch cultivations are the complex genetic regulation of RL production and the excessive foam formation during aerobic cultivation. Additionally, most of the reported fed-batch strategies just rely on heuristic approaches. Batch and fed-batch processes with P. aeruginosa DSM 7107 and DSM 7108 achieved the best-reported RL production in terms of maximum yield and specific productivity (qP). In their patent Giani et al. (1997) claimed a production of more than 100 g/L RLs. Unfortunately, insufficient technical details are provided; especially, no information on the applied analytical method for the RL determination is presented (Giani et al. 1997). Fed-batch cultivation of P. aeruginosa BYK-2 with fish oil as carbon source and urea as nitrogen source resulted in the highest specific yield yet reported, (YP/S) of 0.75 g/g (Lee et al. 2004). A final concentration of 17 g/L of RLs was achieved after 216 h of cultivation. In contrast, a batch strategy resulted in a specific yield YP/S of 0.68 g/g (Lee et al. 2004). A very interesting strategy was proposed by Chen et al. (2007b) who used a pH-stat fed-batch strategy to improve RLs production with 6% glucose in their feed medium. They reported that by excessive feeding of glucose, the accumulation of acidic metabolites occurred, while insufficient supply of glucose lowered RL productivity. Therefore, a pH-stat feeding strategy was applied to control the pH by adjusting the glucose feeding; in that study, a final RL concentration of about 6 g/L was achieved (Chen et al. 2007b). Resting Cells Cultivations Trummler et al. (2003) reported an integrated microbial/enzymatic process with resting cells of Pseudomonas sp. DSM 2874. By a two-step process, the biomass was first produced and harvested. The resting cells were then suspended in a buffer solution and RL production was induced by addition of the carbon source (rapeseed oil). A volumetric productivity (PV) of about 0.14 g/L h was achieved by this method. With the same strain, Syldatk et al. (1985) had reported an improvement of RLs yield coefficient YP/S from 0.16 to 0.23 g/g and YP/X from 0.61 to 3.30 g/g when cultivated under resting cell conditions compared to growth-limiting conditions with nitrogen limitation.

Continuous and Semicontinuous Cultivations Because of foaming problems, semicontinuous strategies have been developed with integrated continuous product removal by flotation. Screenings with Pseudomonas sp. DSM 2874 revealed that the combination of calcium alginate-immobilized cells with glycerol as the carbon source was the best condition for semicontinuous production of RLs (Syldatk et al. 1984). It is possible to reuse the immobilized

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37

biocatalyst several times after appropriate regeneration of the cells (Siemann and Wagner 1993). Continuous processes for the production of RLs are very promising in terms of productivity; relatively high specific and volumetric productivities have been reported for such processes. However, few attempts have been made to promote this process strategy, probably because they are more complex in terms of preparation, realization, and control. Cultures under a continuous process also have a higher risk of contamination. Most of the reported continuous cultivations for RL production have been performed with P. aeruginosa DSM 2659 and dextrose as carbon source. The main characteristics of the performed experiments involved carbon and phosphate excess in addition to nitrogen and iron limitation. The peak of specific productivity occurred at relatively low growth rates (Guerra-Santos 1985; Guerra-Santos et al. 1984, 1986) when strain DSM 2659 was cultivated under continuous conditions (33 C, pH 6.25, 20 g/L dextrose). Ochsner et al. (1996) reported volumetric productivities (PV) of 2 g/L h and a product yield (YP/S) of 0.48 g/g when using corn oil as carbon source for continuous cultivation of P. aeruginosa DSM 2659.

Solid State Fermentation Since they are potent surfactants, foaming is a serious obstacle when producing RLs in an aerated stirred tank bioreactor with a liquid medium. A neat circumvention of this problem is the application of solid state fermentation. Camilios Neto et al. (2008) optimized RL production by P. aeruginosa UFPEDA 614 grown on a solid medium impregnated with a solution containing glycerol. On the basis of the volume of impregnating solution added to the solid support, the yield was in the order of 46 g/L of RLs.

4.1.2

Foaming Problems Encountered During Fermentative Production

A serious challenge encountered during RL production under aerobic conditions is the excessive foaming due to the aeration and agitation of the culture broth in the bioreactor (Chayabutra et al. 2001; Reiling et al. 1986; Walter et al. 2010). Conventionally, chemical antifoaming agents are applied, e.g., based on silicone oil, polyethylene glycol, or polypropylene. This applies to the production of RLs as well (Giani et al. 1997). However, the utilization of chemical antifoaming agents may negatively affect the product quality. Mechanical foam control is an alternative to be considered. Without foam control, the generated foam may drain into the exhaust air duct and block the exhaust air filter. This increases the risk of infections, decreases the productivity, and endangers the whole process. Consequently, the working volume of the bioreactor is usually not completely exploited; rather, it has to be reduced substantially to handle foam formation. Typically, about 40% of the nominal reactor capacity is used (Trummler et al. 2003; Walter et al. 2010).

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In the future, in situ product removal (ISPR) could represent an interesting option for minimizing foam problems during RL production. While ISPR offers some significant advantages, the most important is mitigation of the pronounced foaming. Until now, all trials on ISPR of RLs by filtration have been ineffective due to rapid fouling of the filtration membranes (Gruber 1991). Most bioreactors employed in the production of RLs are stirred-tank reactors for general microbial fermentations equipped with conventional radial impellers (Rushton turbine). These reactors may additionally be equipped with a mechanical foam separator (M€ uller et al. 2010). On a large scale, this may lead to phase separation when utilizing vegetable oils as the carbon source. In such reactors, mixing the highly foamy broth becomes ineffective because of the mode of action of the Rushton turbine. A proposed solution is the combination of an axial propeller and a radial impeller housed in a draft tube, which enhances the dispersion of the hydrophobic substrate by forced vertical circulation (Walas 1997).

4.1.3

Nutritional Factors Affecting Rhamnolipid Production

The effect of different medium culture components, such as carbon and nitrogen sources, and the availability of minerals, on RL production by P. aeruginosa are presented.

Carbon Sources Both water-soluble or water-insoluble carbon sources have been utilized for production of RLs. However, hydrophobic carbon sources such as vegetable oils, are especially effective at promoting the production of RLs. Production processes utilizing a wide range of both natural and petrochemical carbon sources have been published, e.g., l l l l

Vegetable oils; e.g., (Giani et al. 1997; Trummler et al. 2003) Sugars; e.g., (Guerra-Santos et al. 1984; Lee et al. 2004; Reiling et al. 1986) Glycerol; e.g.,(Chen et al. 2007a; Syldatk et al. 1985) Hydrocarbons; e.g., (De´ziel et al. 1996; Syldatk et al. 1985)

Nitrogen, Minerals, and Iron Sources Nitrate is the best nitrogen source for the induction of RLs production (e.g., Arino et al. 1996; Manresa et al. 1991; Mulligan and Gibbs 1989). For the induction of RL formation in a biotechnological set-up, an appropriate limitation must be achieved. For this purpose, the limitation of nitrogen, phosphorus, or multivalent ions in combination with an excess carbon are employed. Interestingly, nitrate as nitrogen source promotes RL production, while ammonium does not (Arino et al. 1996;

Rhamnolipids: Detection, Analysis, Biosynthesis, Genetic Regulation

39

Guerra-Santos et al. 1986). As presented above, this is likely explained by regulatory factors (De´ziel et al. 2003). Under anaerobic, denitrifying conditions with P. aeruginosa ATCC 10145 (28 C, pH 6.8, 2% (v/v) hexadecane), Chayabutra et al. (2001) showed that phosphorus limitation resulted in a four- to fivefold higher productivity as compared to a nitrogen-limited conditions. For RL production under batch and fed-batch conditions at 25 C and pH 7 with P. aeruginosa BYK-2, urea turned out to be the best nitrogen source in combination with fish oil as carbon source (Lee et al. 2004). Not only the type of carbon and nitrogen source but also the respective C/N ratios strongly influence total RL productivity (Guerra-Santos et al. 1984; Santa Anna et al. 2002; Wu et al. 2008). Guerra-Santos et al. showed that for P. aeruginosa DSM 2569 (37 C, pH 6.5, glucose, nitrate) C/N ratios between 16/1 and 18/1 lead to the highest RL productivity while no RLs could be observed at C/N ratios lower than 11/1 (Guerra-Santos et al. 1984, 1986). Apart from phosphorus and nitrogen limitations, restricted availability of multivalent ions like Mg, Ca, K, Na and trace element salts also often result in increased RL yields. For instance, highest final RL concentrations (30 C, pH 6.3, sunflower oil) were observed in calcium-free media (Giani et al. 1997). Abalos et al. (2002) identified the carbon source, the nitrogen source, the phosphate content, and the iron content as critical factors for the medium when producing RLs with P. aeruginosa AT10. The maximum biodrymass of 12.06 g/L was obtained, when the medium contained 50 g/L carbon source, 9 g/L NaNO3, 7 g/L phosphate, and 13.7 mg/L FeSO47H2O. However, the maximum concentration of RLs, 18.7 g/L, was attained in medium that contained 50 g/L carbon source, 4.6 g/L NaNO3, 1 g/L phosphate, and 7.4 mg/L FeSO47H2O.

Low Cost Substrates (Nitrogen and Carbon Sources) RLs can be considered fine chemicals, for example, for pharmaceuticals or cosmetics, or as bulk surfactants, for example, for cleaning products. For highly pure products, the product costs are mainly determined by the downstream processing. However, if high purity is not required, e.g., for bulk applications, the raw material costs are all-dominant and can amount to 50% of the overall production costs (Mulligan and Gibbs 1993). For example, in batch RL production processes, YP/S of 0.13 up to 0.69 g/g have been reported, meaning that between 1.5 and 7.7 times more substrate is consumed than product is synthesized. Therefore, low-cost raw materials should be used. In general, less pure materials are less expensive and they are usually tolerated by the microorganisms. Crude materials or waste materials like soap stock, corn steep liquor, molasses, or nonrefined plant oils are promising carbon sources. On the other hand, this is not the case for the nitrogen source; Inorganic sources like ammonia, nitrate, or urea are generally less expensive than complex nitrogen sources like yeast extract, soybean meal, or casein, if comparing the price in terms of elemental nitrogen content. However, in this respect, corn steep liquor is an exception. This by-product of corn wet-milling is an important

40

A. Mohammad Abdel Mawgoud et al.

constituent of many industrial growth media and is an excellent source of organic nitrogen. Use of different waste substrates has already been reported, e.g., olive oil mill effluents (Mercade et al. 1993), waste frying oils (Haba et al. 2000), soap stock (Benincasa et al. 2002), or waste free fatty acids (Abalos et al. 2001). Additionally, the production of P. aeruginosa biosurfactants, most probably RLs, on whey and distillery waste was reported (Dubey et al. 2005).

4.1.4

Recovery of Rhamnolipids

Downstream processing can represent a significant proportion of the final cost of production of RLs. Most methods of recovery of RLs have been very well reviewed by Heyd et al. (2008). Methods range from those yielding mixtures of different RL congeners to those yielding specific congeners in pure forms. The criteria that govern the selection of a specific recovery method include: (1) the cost associated with the extraction method, which adds to the price of the final product, (2) the proposed purpose of the final product, which influences the level of purity required, and (3) the adaptability of the method to a particular industrial fermentation process. One of the simplest methods of recovery is by acid (De´ziel et al. 1999b; Van Dyke et al. 1993; Zhang and Miller 1992) or aluminum sulfate precipitation (Schenk et al. 1995). Acid precipitation depends on acidification of RL to low pH (e.g., around 2), which neutralizes the negative charges on RLs, making them less soluble in the aqueous phase. Aluminum sulfate precipitates RLs by salting out. The precipitated RLs can then be recovered by centrifugation. Another more commonly used method is recovery by solvent extraction (Le´pine et al. 2002; Mata-Sandoval et al. 1999; Schenk et al. 1995). In this method, molecules are precipitated by acidification and then extracted with organic solvents such as ether or ethyl acetate. Acidification is not a critical step in this method, but it enhances the net yield (Heyd et al. 2008). Other methods adapted to downstream processing in continuous fermentative production processes include: adsorption (Dubey et al. 2005), ion exchange chromatography (Abadi et al. 2009; Reiling et al. 1986; Schenk et al. 1995), ultrafiltration (H€aussler et al. 1998; Mulligan and Gibbs 1990), and foam fractionation (Gruber 1991; Sarachat et al. 2010). Adsorption methods are based on the use of hydrophobic adsorbent such as amberlite XAD 2 or 16 polystyrene resin that retain hydrophobic (or amphiphilic) substances through hydrophobic interactions. Adsorbed RLs are then released by elution, e.g., with methanol. Ion exchange chromatography exploits the fact that RLs behave as anions at high pHs, which allows their retention on columns of weak anion exchange resins such as (diethylamino)ethyl-sepharose. RLs are released from these resins by adding at least 0.6 M NaCl to the equilibration buffer. Yet, this method has been improved by Abadi et al. (2009), who applied phospholipid-coated colloidal magnetic nanoparticles ion exchange media for the recovery and purification of RLs from culture mixtures. Ultrafiltration with a membrane cutoff of 10 kDa leads to an almost complete

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41

retention of RLs even at neutral pH. Foam fractionation depends on the foaming capabilities of RLs; the foam is directed out of the fermentation vessel to a fractionation column where it collapses in a separate receptacle by the action of acids or shear forces, The water in the film surface, known as the lamella, is then allowed to drain by gravitational force, causing a higher concentration of the surfactant in the collapsed foam (Heyd et al. 2008; Sarachat et al. 2010). Most of the aforementioned methods result in the recovery of mixtures of different RLs congeners. Alternatively, chromatographic methods are usually the best solutions for separation of specific RL congeners in a pure form. These methods, however, work better after application of one of the extraction methods mentioned above. On the small scale, preparative TLC is a good choice (Monteiro et al. 2007; Sim et al. 1997); however, for large scale downstream processing, preparative column chromatography using silica gel is a better option (Burger et al. 1966; Monteiro et al. 2007). Recrystallization or repurification using TLC can be applied, if necessary (Heyd et al. 2008).

4.2

Alternatives to P. aeruginosa for Rhamnolipid Production

As P. aeruginosa is an opportunistic human pathogen, there have been several attempts to address the safety issues when producing RLs on a commercial scale. The two primary strategies are the heterologous production of RLs in nonpathogenic bacteria and the utilization of wild-type RL non-pathogenic producers other than P. aeruginosa.

4.2.1

Heterologous Production of Rhamnolipids

The heterologous production of RLs brings along two major advantages as compared to the production with P. aeruginosa. The first is the increased safety when handling large amounts of culture broths. The second is the possibility of constitutive RL production, in contrast to the very tightly regulated production in P. aeruginosa. Several attempts to produce Pseudomonas RLs in heterologous hosts have been reported. Yet, none produces RLs in comparable levels as the best P. aeruginosa strains. In view of a commercial production of RLs, there is still a huge potential for genetic optimization. Ochsner et al. (1995) cloned the rhlAB rhamnosyltransferase gene into various hosts, Pseudomonas fluorescens, Pseudomonas oleovorans, Pseudomonas putida, and E. coli. The best RL production was 60 mg/L and was achieved with P. putida, whereas no production was obtained with E. coli. Cabrera-Valladares et al. (2006) succeeded in producing mono-RLs in E. coli. They found that the availability in E. coli of dTDP-L-rhamnose restricts the production of mono-RLs in this species. By coexpression of the rhlAB and the rmlBDAC operons, the latter encoding the dTDP-L-rhamnose biosynthesis enzymes, they

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generated an RL-producing E. coli strain (productivity was 52.2 mg/L). In contrast, Wang et al. (2007) claimed RL production in E. coli expressing only rhlAB. Cha et al. (2008) reported the heterologous production of RLs in P. putida, using recombinant rhlABRI genes. A maximum yield of 7.2 g/L of RLs was achieved.

4.2.2

Non-P. aeruginosa Rhamnolipid Producers

Conventionally, P. aeruginosa is utilized as production strain for the production of RLs. However, RL-producing bacteria have been found in other species and genera as well. This topic was recently reviewed in details by Abdel-Mawgoud et al. (2010). Most RL-producing species belong to the closely related genera Pseudomonas and Burkholderia in the phylum proteobacteria (Walter et al. 2010). The genus Burkholderia arose from the genus Pseudomonas and was classified as a new genus in 1992 based on 16S rRNA sequence analysis (Yabuuchi et al. 1992). Consequently, bacteria of this genus have characteristics similar to Pseudomonas, and some species indeed produce RLs. B. glumae (formerly Pseudomonas glumae) (Pajarron et al. 1993), B. plantarii, (Andr€a et al. 2006), B. pseudomallei (Dubeau et al. 2009; H€aussler et al. 1998), and B. thailandensis (Dubeau et al. 2009) primarily produce one RL species, Rha-RhaC14-C14. However, a number of other congeners were recently detected in cultures of the two latter (Dubeau et al. 2009), including mono-RLs, mostly Rha-C14-C14. These authors indeed confirmed the very high ratio of di-RLs vs. mono-RLs produced by these species, compared to what is observed in P. aeruginosa cultures (Dubeau et al. 2009). This is probably due to the fact that, as stated above, the Burkholderia rhlC genes encoding the second rhamnosyltransferase are part of the same operon than the rhlA and RhlB homologs, in contrast to the situation in P. aeruginosa (Dubeau et al. 2009). Furthermore, many RL producers belong to Pseudomonas species other than P. aeruginosa (Gunther et al. 2005, 2006; Oliveira et al. 2009; Onbasli and Aslim 2009). In contrast to the di-RLs of Burkholderia species, Pseudomonas chlororaphis synthesizes only RLs with one rhamnose unit and two hydroxy acyl moieties (Gunther et al. 2005). The absence of an rhlC gene homolog is proposed to explain this finding. RLs have been also detected in cultures of many other genera and species of widely different taxonomical origins. Isolates identified as Acinetobacter calcoaceticus, Enterobacter sp (Rooney et al. 2009), Pseudoxanthomonas sp. (Nayak et al. 2009), Pantoea sp. (Rooney et al. 2009; Vasileva-Tonkova and Gesheva 2007), Renibacterium salmoninarum (Christova et al. 2004), Cellulomonas cellulans (Arino et al. 1998b), Nocardioides sp. (Vasileva-Tonkova and Gesheva 2005), and Tetragenococcus koreensis (Lee et al. 2005) have been reported to produce RLs. However, for most of these strains, a structure determination of the putative RLs has not been accomplished, and sometimes, the actual identification of the producing strain is not firmly confirmed.

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As P. aeruginosa is an opportunistic pathogen, another nonpathogenic species (Biosafety level 1) would represent a very interesting alternative if sufficient RL yields can be obtained. The most prominent nonpathogenic RL producers from the genus Pseudomonas are P. chlororaphis (Gunther et al. 2005), P. alcaligenes (Oliveira et al. 2009), and P. putida (Martinez-Toledo et al. 2006; Tuleva et al. 2002), and from the genus Burkholderia, they are B. glumae (Pajarron et al. 1993), B. plantarii (Andr€a et al. 2006), and B. thailandensis (Dubeau et al. 2009). Despite of the apparent safety advantage of these RL producers, very little is yet known about the biotechnological potential of these species.

5 Conclusion: Prospectives for the Industrial Production of Rhamnolipids We have gained a wealth of knowledge on rhamnolipidic surfactants of microbial origin. Still, even though over 60 years have passed since their first description (Jarvis and Johnson 1949), RLs have not yet been significantly employed in the industry. Indeed, there is still a long way before achieving widespread bulk bioproduction of RLs, for both technical and economical reasons. Currently, the economic competitiveness of RLs against synthetic surfactants is mainly determined by the low productivity of the bioprocesses employed. However, this is beginning to change, as environmental compatibility becomes an increasingly important factor for the selection of industrial chemicals. Major improvements can be expected if more productive strains can be found and if a better understanding of the underlying regulation can be attained. In view of the complex quorum sensing-regulated induction of RL production in P. aeruginosa, further optimization will almost certainly be dependent on a more precise understanding of the mechanisms of regulation. It is expected that significant insights on the regulation and biosynthesis of RLs will be gained from the current systems biology approaches. There are good chances of success in the near future if a more integrated biotechnological approach is effectively adopted for strain and process development. Additionally, the use of new heterologous RL-producing hosts will help to broaden the product spectrum and make it possible to produce single RL congeners.

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Walter V, Syldatk C, Hausmann R (2010) Microbial production of rhamnolipid biosurfactants. In: Flickinger MC (ed) Encyclopedia of industrial biotechnology. Wiley VCH, Weinheim, Germany Wang XL, Gong LY, Liang SK, Han XR, Zhu CJ, Li YB (2005) Algicidal activity of rhamnolipid biosurfactants produced by Pseudomonas aeruginosa. Harmful Algae 4:433 443 Wang QH, Fang XD, Bai BJ, Liang XL, Shuler PJ, Goddard WA, Tang YC (2007) Engineering bacteria for production of rhamnolipid as an agent for enhanced oil recovery. Biotechnol Bioeng 98:842 853 White SW, Zheng J, Zhang Y M, Rock CO (2005) The structural biology of type II fatty acid biosynthesis. Annu Rev Biochem 74:791 831 Williams P, Ca´mara M (2009) Quorum sensing and environmental adaptation in Pseudomonas aeruginosa: a tale of regulatory networks and multifunctional signal molecules. Curr Opin Microbiol 12:182 191 Wu JY, Yeh KL, Lu WB, Lin CL, Chang JS (2008) Rhamnolipid production with indigenous Pseudomonas aeruginosa EM1 isolated from oil contaminated site. Bioresour Technol 99:1157 1164 Xiao G, De´ziel E, He J, Le´pine F, Lesic B, Castonguay MH, Milot S, Tampakaki AP, Stachel SE, Rahme LG (2006) MvfR, a key Pseudomonas aeruginosa pathogenicity LTTR class regulatory protein, has dual ligands. Mol Microbiol 62:1689 1699 Xie YW, Ye RQ, Liu HL (2006) Synthesis of silver nanoparticles in reverse micelles stabilized by natural biosurfactant. Colloids Surf A Physicochem Eng Asp 279:175 178 Xie YW, Ye RQ, Liu HL (2007) Microstructure studies on biosurfactant rhamnolipid/n butanol/ water/n heptane microemulsion system. Colloids Surf A Physicochem Eng Asp 292:189 195 Yabuuchi E, Kosako Y, Oyaizu H, Yano I, Hotta H, Hashimoto Y, Ezaki T, Arakawa M (1992) Proposal of Burkholderia Gen Nov and transfer of 7 species of the genus Pseudomonas homology group II to the new genus, with the type species Burkholderia cepacia (Palleroni and Holmes 1981) Comb Nov. Microbiol Immunol 36:1251 1275 Yamaguchi M, Sato M, Yamada K (1976) Microbial production of sugar lipids. Chem Ind 17:741 742 Yilmaz ES, Sidal U (2005) Investigation of antimicrobial effects of a Pseudomonas originated biosurfactant. Biologia 60:723 725 Yoo DS, Lee BS, Kim EK (2005) Characteristics of microbial biosurfactant as an antifungal agent against plant pathogenic fungus. J Microbiol Biotechnol 15:1164 1169 York JD, Firoozabadi A (2008) Comparing effectiveness of rhamnolipid biosurfactant with a quater nary ammonium salt surfactant for hydrate anti agglomeration. J Phys Chem B 112:845 851 Yuan XZ, Ren FY, Zeng GM, Zhong H, Fu HY, Liu J, Xu XM (2007) Adsorption of surfactants on a Pseudomonas aeruginosa strain and the effect on cell surface lypohydrophilic property. Appl Microbiol Biotechnol 76:1189 1198 Zhang YM, Miller RM (1992) Enhanced octadecane dispersion and biodegradation by a Pseudo monas rhamnolipid surfactant (biosurfactant). Appl Environ Microbiol 58:3276 3282 Zhong H, Zeng GM, Yuan XZ, Fu HY, Huang GH, Ren FY (2007) Adsorption of dirhamnolipid on four microorganisms and the effect on cell surface hydrophobicity. Appl Microbiol Biotechnol 77:447 455 Zhong H, Zeng GM, Liu JX, Xu XM, Yuan XZ, Fu HY, Huang GH, Liu ZF, Ding Y (2008) Adsorption of monorhamnolipid and dirhamnolipid on two Pseudomonas aeruginosa strains and the effect on cell surface hydrophobicity. Appl Microbiol Biotechnol 79:671 677 Zhu K, Rock CO (2008) RhlA converts b hydroxyacyl acyl carrier protein intermediates in fatty acid synthesis to the b hydroxydecanoyl b hydroxydecanoate component of rhamnolipids in Pseudomonas aeruginosa. J Bacteriol 190:3147 3154 Zhu L, Lin J, Ma J, Cronan JE, Wang H (2009) The triclosan resistance of Pseudomonas aeruginosa PA01 is due to FabV, a triclosan resistant enoyl acyl carrier protein reductase. Antimicrob Agents Chemother doi: AAC.01152 01109 Zwietering MH, Jongenburger I, Rombouts FM, Van’t Riet K (1990) Modeling of the bacterial growth curve. Appl Environ Microbiol 56:1875 1881

Surfactin and Other Lipopeptides from Bacillus spp. Philippe Jacques

Contents 1

2

3

4

5

Introduction: History of Lipopeptide Discovery in Bacillus spp. . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1 Surfactins from Asia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2 Iturins from Africa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3 Concomitant Discovery of Fengycin and Plipastatin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4 Kurstakins, a New Family of Lipopeptides from Bacillus spp. . . . . . . . . . . . . . . . . . . . . . . 1.5 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A High Diversity of Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Surfactin and Related Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 The Family of Iturins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Fengycin or Plipastatin, Who’s Who? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Other Lipopeptide Compounds, the Kurstakins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Catalytic Assembly Lines for the Biosynthesis of Lipopeptides: From the Genes to the Biomolecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Modular Enzymes: A Complex Catalytic Machinery Dedicated to the Biosynthesis of Secondary Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Non ribosomal Peptide Synthesis of Surfactin and Lichenysin . . . . . . . . . . . . . . . . . . . . . . 3.3 The Hybrid PKS/NRPS Complex Involved in Iturin Biosynthesis . . . . . . . . . . . . . . . . . . . 3.4 Non ribosomal Peptide Synthesis of Fengycin and Plipastatin . . . . . . . . . . . . . . . . . . . . . . 3.5 The Recent Discovery of the Biosynthesis Mechanism of Kurstakin . . . . . . . . . . . . . . . . 3.6 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A Complex Regulation of the Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Quorum Sensing and Surfactin Efflux . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Influence of Environmental Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Physico chemical Properties and Biological Activities: A Strong Relationship . . . . . . . . . . . 5.1 Surfactin: A Potent Biosurfactant, Which Combines High Effect on Surface Tension and Low Critical Micellar Concentration . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Iturin: A Strong Antifungal Compound . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Fengycin and Plipastatin: Immunomodulators in Plants and Animals . . . . . . . . . . . . . . .

58 58 59 60 60 60 60 61 63 64 64 65 65 65 68 70 71 71 72 72 72 74 75 75 75 77 77

P. Jacques ProBioGEM, Polytech Lille, Univ Lille Nord de France, USTL 59655 Villeneuve d’Ascq, France e mail: Philippe.Jacques@polytech lille.fr

G. Sobero´n‐Cha´vez (ed.), Biosurfactants, Microbiology Monographs 20, DOI 10.1007/978 3 642 14490 5 3, # Springer Verlag Berlin Heidelberg 2011

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5.4 Lipopeptides: Versatile Weapons for Biocontrol of Plant Diseases . . . . . . . . . . . . . . . . . . 78 5.5 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78 6 New Strategies for an Optimal Production of Novel or Existing Lipopeptidic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78 6.1 Surfactin Synthetases Re engineering . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 6.2 Combinatorial Synthesis of Lipopeptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 6.3 Directed Biosynthesis and Molecular Optimisation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80 6.4 New Bioprocesses for Continuous Production and Extraction of Lipopeptides . . . . . . 80 6.5 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 7 Industrial Applications: Dream and Reality . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 7.1 Main Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 7.2 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83

Abstract Isolated during 1950s and 1960s, the group of lipopeptides from Bacillus spp. gather more than 30 different peptides linked to various fatty acid chains. More than a 100 different compounds can so be described. In this chapter, they are classified into four main families: the surfactins, the iturins, the fengycins or plipastatins and the kurstakins. The biochemical mechanism responsible for their biosynthesis, which involved nonribosomal peptide synthetases, is described in detail. The complex cascade of regulation of surfactin synthetase operon and the environmental factors, which influence the lipopeptide production, are discussed. The main physico-chemical properties of these remarkable biosurfactants and their possible relationships with the biological activities are also presented. A brief overview of the molecular strategies developed to get modified lipopeptide compounds and the last bioprocesses set up for their production are given. In the last chapter, the main applications of surfactin are proposed.

1 Introduction: History of Lipopeptide Discovery in Bacillus spp. In this section, we discuss a short history of the discovery of the various lipopeptides from different Bacillus species and the genes involved in their biosynthesis. Up to now, these lipopeptides were classified into three different families: surfactins, iturins and fengycins (or plipastatins). Following the discovery of kurstakins in 2000, we suggest that they should be added to the family of lipopeptides. A first overview of the different members of each lipopeptide family and the producing species is given.

1.1

Surfactins from Asia

In 1968, Arima et al. isolated an exocellular compound with an exceptional biosurfactant activity from the supernatant of a culture of Bacillus subtilis. This

Surfactin and Other Lipopeptides from Bacillus spp.

59

compound was named surfactin and its structure was elucidated as that of a lipopeptide (Kakinuma et al. 1968). It was characterised as: a valuable inhibitor of fibrin clot formation, an antibacterial, antitumour and hypocholesterolemic agent. Other strains or species producing surfactin derivatives (Bacillus coagulans, Huszcza and Burczyk 2006, Bacillus mycoides, Athukorala et al. 2009) or related compounds, such as esperin (Thomas and Ito 1969), halobacillin isolated from a marine Bacillus strain (Trischman et al. 1994), lichenysin from Bacillus licheniformis (Horowitz et al. 1990), pumilacidin from Bacillus pumilus (Morikawa et al. 1992) or bamylocin A from Bacillus amyloliquefaciens (Lee et al. 2007), were also identified. The srfA operon and the enzymes responsible for the biosynthesis of the surfactin were the first described for a lipopeptide from Bacillus subtilis (Nakano et al. 1991; Menkhaus et al. 1993). This discovery allowed for the confirmation that this lipopeptide is synthesised by the non-ribosomal pathway (see Sect. 3). The operon involved in lichenysin biosynthesis was then described by Konz et al. (1999).

1.2

Iturins from Africa

Mycosubtilin was the first antifungal lipopeptide from Bacillus subtilis mentioned in literature in 1949 (Walton and Woodruff). A second similar compound named iturin was described by Delcambe (1950). Its name is related to the Ituri, a region from Congo where the compound was isolated from a soil sample. Iturin was first characterised as a strong antifungal agent with a restricted antibacterial activity against Micrococcus and Sarcina strains. The precise structure of these two compounds and similar compounds from the same species was described during the 1970s and 1980s: mycosubtilin (Peypoux et al. 1976, 1986), bacillomycin L (Besson et al. 1977) identical to bacillomycin Lc or bacillopeptin (Eshita et al. 1995; Volpon et al. 2007), iturin A (Peypoux 1978), iturin C (Peypoux et al. 1978), bacillomycins D (Peypoux et al. 1981) and F (Peypoux et al. 1985). Production of iturinic compounds was also identified in other species such as: Paenibacillus koreensis (Chung et al. 2000), Bacillus amyloliquefaciens (Yu et al. 2002) and Bacillus pumilus (Cho et al. 2009). Mycocerein, an antifungal peptide produced by Bacillus cereus was partially described and could belong to the iturin family (Wakayama et al. 1984). The genes involved in the biosynthesis of iturinic compounds have been first characterised for mycosubtilin in Bacillus subtilis ATCC 6633 (Duitman et al. 1999) and then for iturin A (Tsuge et al. 2001a) and bacillomycin D (Moyne et al. 2004; Koumoutsi et al. 2004).

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P. Jacques

Concomitant Discovery of Fengycin and Plipastatin

In 1986, German (Vanittanakom et al. 1986) and Japanese teams (Nishikiori et al. 1986) discovered simultaneously a third family of lipopeptides: fengycin produced by Bacillus subtilis and plipastatin from Bacillus cereus. The first lipopeptide was determined as an antifungal agent and the other as a phospholipase A2 inhibitor. Only small structural differences exist between these two compounds and a doubt still exists today about them as well as the differences in their biological activities. Production of fengycin was also demonstrated by Bacillus thuringiensis (Kim et al. 2004). The operon encoding fengycin or plipastatin synthetases was first described in Bacillus subtilis 168 in 1997 (Tosato et al. 1997) and then in Bacillus subtilis b213 and A1/3 in 1999 (Steller et al. 1999) and in Bacillus amyloliquefaciens FZB42 in 2004 (Koumoutsi et al. 2004).

1.4

Kurstakins, a New Family of Lipopeptides from Bacillus spp.

The use of Matrix-assisted laser desorption/ionisation (MALDI) to characterise and differentiate between different species or strains of Bacillus spores allowed Hathout et al. (2000) to discover a new family of lipopeptides produced by Bacillus thuringiensis subsp. kurstaki and consequently called kurstakin. These compounds display antifungal activity against Stachybotrys charatum and are mainly adsorbed to the spore surfaces. Additionally, the potential operon encoding kurstakin synthetases was recently discovered (Bumpus et al. 2009; Abderrahmani et al. 2010).

1.5

Conclusion

Since the discovery of iturins, more than 30 different lipopeptides produced by different strains of Bacillus spp. have been characterised revealing the high potential of lipopeptide biosynthesis of this genus. The exponentially growing number of genome sequences should probably allow to discover new families or new variants in the near future.

2 A High Diversity of Structures This section is dedicated to the description of the structure of the different lipopeptides. The different variants of each family are described in detail. Fig. 1 shows a detailed structure of two compounds of each family. Table 1 gives an overview of the different peptide sequences and lipidic moiety of the different variants described in each family.

Surfactin and Other Lipopeptides from Bacillus spp.

a

61

b

c

O

H N 2

O

C

O

C

OH O

H C 2 CH

3

CH

CH

HC NH

CH

CH CH

3

C

CH

NH

C

2

O

O

2

CH 2O

H C 3 CH

H C 3

HC

2

CH CH

CH

2

CH

CH

CH

CH

CH

2

2

H C 2

C

CH 3

f

2

CH

HC

CH

C

NH

CH 2

CH

N

CH

HC CH

CH

2 HO

2

NH

NH

CH

2

CH

2

CH

CH

2

H C 2 CH

CH

CH

3

HO

OH 2

H C 2

OH

H N

C CH

H C 3

CH 2

H C 2 CH

CH

CH 2

C

CH

3

NH

CH

2

HC CH

3

HC

O

O

C

CH

CH

2

2

N

CH 2

C

H C 2

CH 2

CH

HC

CH

CH

O C

CH

C

H C 2

N CH

2

HC

NH

CH

HC C

CH

2

H N

C

O

NH 2

CH

HC

NH

O C

C HC

HO

OH 2

CH 2

O

O

CH

NH

CH 3

2

H C 2

3

C O

NH

CH

C

H C 2

CH

C

O C

NH

CH

H C 2

O

CH

CH 2

C

2

C NH

HC

NH

O

H

H

CH

O C

CH 2

O HC

C O

2

2

O

O

O

2

2

CH

CH 2

H C 2

CH

C

NH

OH

CH CH

CH 2

2

2

CH

2

HO

O

O

CH

CH

CH

2

2

2

2

2

HC CH

2

CH

NH

C

N H

3

2

2

2

C CH

CH

2

C

O

C

2

CH

CH

OH

C CH

CH

2

2

CH

2

H C 2

H N 2

C HC

CH CH

O

C HN

2

CH

3 CH

CH

2

C

CH

2

H C 2

O

CH

2

H C 2

2

2

CH CH

H N

2

2

H C 2

e

O

CH

2

HO

2

H C 2

O

CH

H C 3

NH

2

H C 2

CH CH

C

2

CH

O

O

H C 2

O

NH

O

H N 2

2

O HO

HC

CH 2

CH

2

O

CH

C

CH CH

2

d

C

2

NH

NH

C

H C 2 C

H C 3

O

C

CH H C 2

H C 2

CH

O

O

2

CH

O CH

C HO

CH 3

H C 3

H C 2 O

H C 2

H N 2

O

2

H C 2

C

NH

C

HC

C

CH

HC

2

O

C

CH

H C 2 CH

CH

CH

H N 2

H C 2

C

2

NH

NH

C

CH 2

C

CH

H C 2

O

C

CH

2

CH HC

NH

O

3

H C 2 C

CH

H N 2

HC

HC

CH

H C 3 O

HC

C

H C 2

C

O

NH

C

NH

C

CH

CH

C CH

HC

2

NH

O

CH 3

CH

HN

NH

O CH

HO

CH 2

HC

H C 3

O

CH

2O

O

H N 2

C

HC

C

H C 2

2

CH

C

O

C

CH

CH

2

HC

CH

NH

O

H C 3

H C 3

CH

CH

CH

N

C

2

NH

HN

H C 3

CH C

C

CH

NH

HC

CH

2

O 3

O

C

O 2

CH

CH

H C 2

HC

C

CH

CH 2

H C 3

C

NH

C

O

CH

3

O

CH

CH

CH CH

NH

O H N

O

H C 3

H C 2

OH

H C 2

3

C

C

CH 2

C

CH

CH

H N

O H C 3

H C 3

CH CH

2

NH

CH 2

2

CH

CH

C

O

C

O

C

NH

NH

H C 2

O

CH

O

2

HC CH 2

H C 2 CH

CH

O

NH

2 H C 3

HC

H C 3

O

HC

H

2

2

O

CH O

CH H C 3

C

NH C

HC

NH 2

O H C 3

NH

2

2

C

O

HC

O

CH O

CH

2

NH 2

NH C

HC

C

CH 2

CH

C CH

CH

CH

CH

CH

2

C C

C H C 3

HC C

CH 2

OH

CH

H C 3

OH

CH CH

CH

Fig. 1 Detailed structure of different lipopeptides (a) Lichenysin nC14 (b) Surfactin nC14 (c) Iturin nC14 (d) Bacillomycin F aC17 (e) Fengycin/plipastatin A nC15 (f) Fengycin/plipastatin B nC15

2.1

Surfactin and Related Compounds

The family of surfactin is composed of about 20 different lipopeptides (Bonmatin et al. 2003). With the exception of esperin (Thomas and Ito 1969), they have the common following structural traits: a heptapeptide with a chiral sequence LLDLLDL interlinked with a b-hydroxy fatty acid and with a D-Leu in position 3 and 6 and a L-Asp in position 4 (Figure 1). Amino acid residues in position 2, 4 and 7 belong to the aliphatic group including Val, Leu and Ile (Peypoux et al. 1991; Itokawa et al. 1994; Bonmatin et al. 1995). A surfactin with Ala in position 4 was also observed (Peypoux et al. 1994). The presence of these variants can be related to alterations in the culture conditions, in particular, the feeding with some specific amino acid residues. This is a result from the mechanism of biosynthesis of such

Heptapeptide with a lactone ring between carboxy-terminal group of Gln7 and OH group of Ser4 D-Thr-Gly-D-Ala-Ser-His-D-Gln-Gln Heptapeptide closed by a lactone ring with the b-OH group of the fatty acid chain Glu-Leu-Met-Leu-Pro-Leu-Leu-Leu L-Glu-L-Leu-D-Leu-L-Val-L-Asp-D-Leu-L-XE7-COOH L-XL1-L-XL2-D-Leu-L-XL4-L-Asp-D-Leu-L-XL7 L-Glu-L-Leu-D-Leu-L-Leu-L-Asp-D-Leu-L-XP7 L-Glu-L-XS2-D-Leu-L-XS4-L-Asp-D-Leu-L-XS7

L-Asn-D-Tyr-D-Asn-L-Gln-L-Pro-D-Ser-L-Asn

L-Asp-D-Tyr-D-Asn-L-Gln-L-Pro-D-Asn-L-Ser

Primary structure of the peptide moiety Decapeptide with a lactone ring between carboxy-terminal group of Ile10 and OH group of Tyr3 L-Glu-D-Orn-D-Tyr-D-a Thr-L-Glu-D-Ala-L-Pro-L-Gln-L-Tyr-L-Ile L-Glu-D-Orn-D-Tyr-D-a Thr-L-Glu-D-Val-L-Pro-L-Gln-L-Tyr-L-Ile L-Glu-D-Orn-D-Tyr-D-a Thr-L-Glu-D-Ala-L-Pro-L-Gln-D-Tyr-L-Ile L-Glu-D-Orn-D-Tyr-D-a Thr-L-Glu-D-Val-L-Pro-L-Gln-L-Tyr-L-Ile Heptapeptide closed by a lactam ring with the b-NH2 group of the acid chain L-Asn-D-Tyr-D-Asn-L-Pro-L-Glu-D-Ser-L-Thr L-Asn-D-Tyr-D-Asn-L-Gln-L-Pro-D-Asn-L-Thr L-Asn-D-Tyr-D-Asn-L-Ser-L-Glu-D-Ser-L-Thr L-Asn-D-Tyr-D-Asn-L-Gln-L-Pro-D-Asn-L-Ser L-Asn-D-Tyr-D-Asn-L-Gln-L-Pro-D-Asn-L-Ser

C13 C13, C14, C15 iC13, aC13, nC14 iC15, aC15 aC15, iC15, nC16, iC16, aC17, iC17 iC14, nC14 iC15, aC15

iC11, nC12, iC12, iC13 b-OH fatty acids

nC14, iC15, aC15 nC16, iC16, aC17 b-OH fatty acid chain

Schneider et al Schneider et al Nishikiori et al Nishikiori et al

aC15, iC16, nC16 aC15, iC16, nC16, C17 nC16, aC17 nC16, aC17 b-NH2 fatty acids nC14, iC15, aC15 nC16, iC17, aC17 nC14, iC15, aC15 nC14, iC15, aC15 nC16, iC16

1999 1999 1986 1986

Lee et al 2007 Thomas and Ito 1969 Lin et al 1994 Naruse et al 1990 Peypoux et al 1999

Hathout et al 2000

Peypoux et al 1981 Peypoux et al 1985 Volpon et al 2007 Peypoux 1978 Winkelmann et al 1983 Peypoux et al 1986 Peypoux et al 1986

References

Main fatty acid chains b-OH fatty acids

b

Or bacillopeptin D forms of amino acid residue are deduced from NRPS modular structure (Bumpus et al. 2009; Abderrahmani et al. 2010) c L and D forms are not specified d The b-carboxyl of Asp5 is engaged in the lactone e Or halobacillin With XE7 ¼ Leu or Val, XL1 ¼ Gln or Glu; XL2 ¼ Leu or Ile; XL4 and XL7 ¼ Val or Ile; XP7 ¼ Val or Ile; XS2 ¼ Val, Leu or Ile; XS4 ¼ Ala, Val, Leu or Ile; XS7 ¼ Val, Leu or Ile

a

Bamylocin Ac Esperind Lichenysine Pumilacidin Surfactin

Kurstakin Surfactin family

b

Iturin C Mycosubtilin Kurstakin family

Fengycin A Fengycin B Plipastatin A Plipastatin B Iturin family Bacillomycin D Bacillomycin F Bacillomycin L or Lca Iturin A Iturin AL

Name Fengycin family

Table 1 Peptide sequence and fatty acid chains of the different variants of each lipopeptide family from Bacillus spp.

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compounds that involve non-ribosomal peptide synthetases (NRPS) (see Sect. 3). In such megaenzymes, the amino acid residue, which will be incorporated into the peptide, is first specifically recognised and activated by an adenylation domain. It was shown in several NRPS that the adenylation domains are not always specific and can accept some amino acid residues with similar structure. Interestingly, the presence of D-Leu seems specific, which could result from a higher specificity of the involved adenylation domain or of the epimerisation domains responsible for the transformation of the L-Leu to D-Leu. In position 1, two different amino acids are observed, Glu and Gln. It is stated in literature that lipopeptide with a Glu in position 1 are named surfactin, while those with a Gln are named lichenysin as they were discovered in the supernatant of Bacillus licheniformis culture (Horowitz et al. 1990). The presence of an Asn in position 4 was first mentioned in the structure described for lichenysin A by Yakimov et al. (1995) but the use of fast atom bombardment mass spectrometry allowed the same authors to definitively confirm the presence of an Asp as in surfactin (Yakimov et al. 1999). A surfactin-like compound was isolated from Bacillus pumilus culture supernatants and was called pumilacidin (Morikawa et al. 1992). Structural analysis shows that they correspond to Leu4, Val7 or Ile7 surfactin. Esperin differs from the surfactin by a lactone ring involving the b-carboxyl of Asp in position 4 instead of the a-carboxyl of the terminal Leu. The b-hydroxy fatty acid chain linked to these different peptide moieties can contain 12 16 C atoms and show n, iso and anteiso configurations. The main fatty acid chains are usually C14 and C15. Recently a new lipopeptide, bamylocin A, was isolated from Bacillus amyloliquefaciens (Lee et al. 2007). The peptide chain of this molecule is Glu-Leu-Met-Leu-Pro-Leu-Leu-Leu. The molecular weight of the C13 form differs by less than 0.1 mass unit from the standard surfactin C14 isoform. This new result demonstrates the need of a precise MS-MS analysis of surfactin products to confirm their primary structure. Surfactin-O-methyl ester was isolated from purified culture broth of different strains. Originally, it was considered to be the result of chemical modification, which occurs during the extraction procedure by methanol. The biological origin of such a methyl ester has been, however, confirmed by Liu et al. (2009) by using acetonitrile instead of methanol during the purification procedure of surfactin compounds produced by Bacillus subtilis HSO121.

2.2

The Family of Iturins

Iturin A, which is the main studied lipopeptide of the iturin family, is a heptapeptide interlinked with a b-amino acid fatty acid with a length from C14 to C17 (Peypoux 1978). Six other members of the iturin family were then described: iturin C, bacillomycin D, F, L and Lc and mycosubtilin (Bonmatin et al. 2003). All iturins have the same LDDLLDL chiral sequence with a common part of the peptide cycle: b-amino acid L-Asx D-Tyr D-Asn. Except for Iturin C, the first amino acid of the peptide chain is L-Asn. Recent work from Volpon et al. (2007) confirms the presence

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of L-Asn in position 1 of bacillomycin L instead of L-Asp initially described and thus demonstrates bacillomycin Lc and bacillomycin L to display the same structure. Contrary to most of the different variants of surfactin, which are synthesised by the same enzymes accepting different substrates, these different members of the iturin family probably results from different synthetases (see Sect. 3). In addition, the main length of the fatty acid chain differs from one member to another. Iturin A and C and bacillomycin D, L, each display a fatty acyl chain with a length of 14 and 15 C, while the C16 and C17 forms are the main representative fatty acid of the bacillomycin F and the mycosubtilin. A form of Iturin A with long fatty acyl chain (C16) was identified by Winkelmann et al. (1983) and called Iturin AL.

2.3

Fengycin or Plipastatin, Who’s Who?

This family of lipopeptides includes fengycins A and B, which are also called plipastatins. These molecules are lipodecapeptides, which differ by their amino acid residue in position 6 that can be Ala (form A) or Val (form B). They display an internal lactone ring in the peptidic moiety between the carboxyl terminal amino acid (Ile) and the hydroxyl group in the side chain of the tyrosine residue in position 3. Different bhydroxy fatty acid chains (C14 to C18) are linked with an amide bond to the Nterminal amino acid residue (Glu) (Nishikiori et al. 1986; Vanittanakom et al. 1986). The main representative fatty acid chains are C15, C16 and C17. They are saturated, except in the case of a single lipopeptide isolated from the supernatant of Bacillus thuringiensis, which has a fatty acid chain with one double bond between carbons 13 and 14 (Kim et al. 2004). Two differences were initially identified between fengycin and plipastatin: a Gln instead of a Glu in position 8 and the L and D forms of tyrosine, which are in position 3 and 9, respectively, for plipastatins and 9 and 3 for fengycins. In the different works describing fengycin structure since its discovery, the presence of a Glu in position 8 was never mentioned. However, we confirmed the existence of fengycin molecule with a D-Tyr in position 3 from the supernatant of Bacillus subtilis S499 (Schneider et al. 1999). This result cannot be correlated with the structure of the synthetases described in other fengycin- or plipastatin-producing strains (see Sect. 3) (Tosato et al. 1997; Steller et al. 1999; Koumoutsi et al. 2004).

2.4

Other Lipopeptide Compounds, the Kurstakins

Kurstakins are a novel class of lipopeptides composed of several lipoheptapeptide with the same amino acid sequence: Thr-Gly-Ala-Ser-His-Gln-Gln (Hathout et al. 2000). Different fatty acyl chains (isoC11, nC12, isoC12 and isoC13) are linked via an amide bond to the N-terminal amino acid residue. Each lipopeptide has a lactone linkage between the carboxyl terminal amino acid and the hydroxyl group on the side chain of the serine residue. The L and D forms or the amino acid residues are

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not yet characterised. However, the recent identification of the genes involved in the biosynthesis of such or similar compounds indicate that amino acids in 1 and 6 positions could be in the D-form (Bumpus et al. 2009; Abderrahmani et al. 2010) (see Sect. 3.5).

2.5

Conclusion

Each family of lipopeptides is constituted of several variants, which can differ in their fatty acid chain and their peptide moiety. The resulting wide diversity of molecules can be used to study the relationships between the structure and function of the lipopeptides. However, the existence of several different compounds with the same molecular weight shows that it is essential to precisely characterise their structure by using, for example, LC-MS-MS techniques.

3 Catalytic Assembly Lines for the Biosynthesis of Lipopeptides: From the Genes to the Biomolecules In this section, we briefly discuss an overview of the NRPS, which is responsible for the biosynthesis of lipopeptides from Bacillus spp. We will then focus on the precise organisation of the catalytic assembly lines involved in the biosynthesis of surfactin and lichenysin and especially the spatial arrangement of the termination module of surfactin synthetase, a recently determined structure. A precise description of the other NRPS, which catalyse iturin A, mycosubtilin, bacillomycin D, fengycin, plipastatin and kurstakin, will be then given. Figure 2 summarises the modular organisation of all these multifunctional proteins.

3.1

Modular Enzymes: A Complex Catalytic Machinery Dedicated to the Biosynthesis of Secondary Metabolites

Here, we describe the enzymatic machinery for the non-ribosomal synthesis of peptides.

3.1.1

Discovery of the Non-ribosomal Peptide Synthesis

In 1968, Gevers et al. demonstrated for the first time using cell extracts of the producer strains that biosynthesis of gramicidin (a peptide antibiotic) is possible in the presence of RNases or inhibitors of the ribosomal machinery. It was

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AL C KS C AMT C C C

A

P

P

P

P C C P

P

A

A

M C T

A

Asn

fenF Mycosubtilin

P C E C P

P C E C P

A

Asn

Tyr

P C C P

P C C P

A

A

P C E C P

Ser

Pro

Gln

mycA Tyr

Asn

P C TE P

mycC Ser

Asn

Pro

Gln

ituA

A

Asn

mycB Asn

ituD

A

ituB

ituC

Iturin Asn

Asn

Tyr

Pro

Thr

Ser

Glu

bmyB

bmyA

bmyD

bmyC

Bacillomycin

C

P C C P

A

Glu

A

P C C P

Leu

P C E P

A

P C C P

C A

Val

Leu

srfA-A

A

P C C P

Asp

A

P C E P

A

C

Leu

P TE/ C TE AT P

Leu

srfA-B

srfA-C

srfA-D

Surfactin Leu

Gln

Leu

Val

IchAA

Asp

Ile

Leu

IchAB

IchAC

IchA-TE

Lichenysin A

P

P

P

P

C A CC A CE

Glu

P

P

P

P

P

P

P

P

P

P

P

P

P

P

C A CC A CE C A C C A CE C A CC A CC A CE

Orn

Tyr

ppsA/fenC

A-Thr

Glu

ppsB/fenD

Pro

Ala/Val

Gln

ppsC/fenE

Tyr

ppsD/fenA

P

C A CTE P

Ile

ppsE/fenB

Plipastatin/Fengycin P

C A CE P

Thr ORF1(GrsC)

P

P

P

P

P

P

P

P

P

P

P

P

C A C C A C C A C C A C C A C E C A CTE

Gly

Ala

ORF2(NRPS)

Ser

His

Gln

Gln

ORF3(Bac1)

Kurstakin

Fig. 2 Operons responsible for lipopeptide biosynthesis in Bacillus spp. Domains are described in the text

a key experiment to show that another biosynthetic pathway exists for the biosynthesis of the peptides. Since this discovery, many works were carried out to describe in detail the non-ribosomal peptide biosynthesis (Finking and Marahiel 2004). It is responsible for the synthesis of more than 1,000 active biomolecules that can be gathered in about 200 families. A recent database (NORINE) compiles most of the non-ribosomal peptides (NRPs) (Caboche et al. 2008).

Surfactin and Other Lipopeptides from Bacillus spp.

3.1.2

67

The Main Catalytic Domains

Non-ribosomal peptide synthesis involves large multienzymatic proteins called non-ribosomal peptide synthetases (NRPS), which are organised in modules (Sieber and Marahiel 2005). Each module is responsible for the incorporation of one building block into the growing polypeptide chain and can be subdivided in a defined section of the protein called a domain involved in a specific enzyme activity. Four main domains are present in most of the NRPS: adenylation (A), thiolation (T), condensation (C) and thioesterase (TE) domains. The adenylation domain selects the cognate amino acid and activates it as amino acyl adenylate. Several crystal structures of A-domains have been solved to date. The first two structures were the phenylalanine-activating A-domain of the gramicidin Ssynthetase A (GrsA) from Bacillus brevis (Conti et al. 1997) and the 2,3-DHBactivating A-domain DhbE from B. subtilis (May et al. 2002). These crystal structures facilitate the assignment of ten amino acid residues that play a decisive role in the coordination of the substrate. The so-called non-ribosomal code can be used to predict an A-domain selectivity on the basis of its primary sequence (Stachelhaus et al. 1999). The activated amino acid is then transferred to the Tdomain, also called peptidyl-carrier protein (PCP), as it is the transport unit of the activated intermediate. In such a domain, the activated amino acid residue is covalently tethered to its 40 phosphopantheteinic (40 -PP) cofactor as thioester. This cofactor is post-translationally transferred to a serine of the PCP. This reaction is catalysed by a phosphopantetheinyl transferase (encoded in Bacillus subtilis by the sfp gene), which is thus essential to transform apoform of NRPS in its holoform (Mofid et al. 2004). The 40 PP cofactor acts as a flexible arm to allow the bound amino acyl and peptidyl substrate to travel between different catalytic centres. The C-domain catalyses the formation of the peptide bond between amino acyl substrate bound to PCPs of adjacent modules. The termination of synthesis is catalysed by the terminal enzyme of the last module. In most cases and for lipopeptide synthesis, this reaction is performed by a thioesterase domain (TE). This allows the release of the peptide and is also frequently involved in the formation of a macrocyclic product (lactones and lactams) or the oligomerisation of peptide units (Kopp and Marahiel 2007). Other alternative release mechanisms can be achieved by the reduction of the peptidyl-S-PCP final product to generate a linear aldehyde or alcohol.

3.1.3

Secondary Catalytic Domains

Additional enzymes can be involved in the biosynthesis of the peptide to modify the structure of the monomer involved in the primary structure or to add some external compounds to the peptide. Some of these are integral parts of the NRPS and act in cis whereas others are distinct enzymes acting in trans before full maturation of the NRPS product. Among these tailoring domains, which include cyclisation (Cy), methylation (Me), oxidation (Ox), glycosylation, epimerisation (E) and addition of fatty acid chain, the final two are involved in lipopeptide biosynthesis in Bacillus

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spp. The E-domain catalyses the epimerisation of the PCP-bound L-amino acid of the growing polypeptide chain. The addition of the fatty acid chain to the first amino acid of the peptide moiety is catalysed by a first specific condensation domain, also called starter condensation domain. The added fatty acid chain can itself be partially synthesised by another main group of modular enzymes, the polyketide synthases (PKS). In this last case, a hybrid PKS/NRPS is required for the synthesis of the biomolecules (Du et al. 2001).

3.1.4

Protein–Protein Interactions

In most multienzyme complexes, several proteins are involved in the complete assembly lines. The biosynthesis of the right NRP requires the proper protein protein interaction between partner enzymes and concomitantly prevents undesired interactions between non-partner enzymes. Short terminal structures, referred to as NRPS communication-mediating (COM) domains are responsible, at least for the most part, for the correct channelling of reaction intermediates along the assembly line (Hahn and Stachelhaus 2004). A donor COM domain COMD X, situated at the C terminus of an aminoacyl- or peptidyl-donating NRPS “X”, and an acceptor COM domain COMA Y, located at the N terminus of the accepting partner enzyme “Y”, form a compatible (cognate) pair that is crucial for establishing the productive interaction between both enzymes. In contrast, within a hypothetical assembly line “X Y-Z”, the COM domains COMD X and COMA Z of the nonpartner NRPSs “X” and “Z” are considered incompatible (non-cognate), preventing their futile contact. Accordingly, the establishment of a defined assembly line and synthesis of a distinct NRP product is ensured by the grouping of exclusively cognate pairs of COM domains. Donor and acceptor NRPS COM domains comprise 15 30 amino acid residues.

3.2

Non-ribosomal Peptide Synthesis of Surfactin and Lichenysin

The structure of the enzymatic complexes participating in the synthesis of surfactin and lichenysin has been described in detail and is presented in this section.

3.2.1

The Surfactin Operon

Three large Open Reading Frames (ORFs) coding for surfactin synthetases are designated srfA-A, srfA-B and srfA-C (Galli et al. 1994). They present a linear array of seven modules (one module per residue), three modules are present in the products of SrfA-A and SrfA-B, respectively, and the last one in SrfA-C. A high specificity was observed for adenylation domains of modules 1, 3, 5 and 6, which recognised L-Glu, L-Leu, L-Asp and L-Leu amino acid residues, respectively. Modules 3 and 6

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69

contain an epimerase domain (E), which transforms the incorporated L-Leu in a DLeu. In vitro studies of the specificity of adenylation domains of modules 2, 4 and 7 show that they are able to accept several aliphatic amino acid residues as substrates. The b-hydroxylated fatty acid chain is added to the amino acid activated in the first module by the way of a starter condensation domain. A first thioesterase fused with the carboxy-terminal end of the final activation PCP domain is responsible for the release of the synthesised product from the enzymatic template. This enzyme also catalyses the lactone bond formation between the carboxylate group of the last amino acid and the hydroxyl group of the fatty acid chain. Chemoenzymatic approaches were used to characterise the specificity of the thioesterase domain. Different heptapeptides were synthesised by chemical means, and a thioesterase domain was cloned and produced as an isolated enzyme. The presence of Glu in position 1 and the two Leu in positions 6 and 7 are essential for substrate recognition by the thioesterase domain. A second thioesterase/acyltransferase (TE/At-domain) encoded by a fourth gene, srfA-D, stimulates the initiation of the biosynthesis. Two cognate-pairs of communication-mediating (COM) domains, COMDSrfA-A/COMASrfA-B; COMDSrfA-B/ COMASrfA-C, facilitate the selective interaction between partner enzymes (Chiocchini et al. 2006). A small gene, designated comS, is located within the coding region of the fourth amino acid-activation domain of srfA and thus co-expressed with the srfA operon. This gene is required for competence development in Bacillus subtilis but not directly involved in the biosynthesis of surfactin (Hamoen et al. 1995).

3.2.2

The Lichenysin Operon

The same organisation in three ORFs was described for lichenysin operon (Konz et al. 1999). Only small differences were observed in the primary sequences of the corresponding synthetases, especially in the ten amino acids representing the Nonribosomal code, present in the active site of adenylation domains of module 1 and 7. In the module 1, the modification of a Lys to Glu led to the incorporation of a Gln instead of a Glu in position 1 of the lipopeptide. In the module 7, the replacement of an Ala by a Gly and a Cys by a Val in the active site of the adenylation domain favours the incorporation in the peptide formed of an Ile instead of a Leu.

3.2.3

Structure of an Entire Termination Module

A variant of the termination module SrfA-C of the surfactin synthetase, in which the active site serine of PCP was changed into alanine by site-directed mutagenesis, provided diffracting crystals (Tanovic et al. 2008). The structure of the SrfA-C variant was solved at 2.6 resolution and covered the entire module with its four catalytic domains and linker regions in between. This is the first example of the resolution of the complete structure of a NRPS module. The SrfA-C structure provided unique information on how the essential catalytic

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domains are orientated and spatially arranged. Overall, the structural core of the module is a compact rectangular catalytic platform mainly built by the intimate association of the C-domain and the large N-terminal part of the A-domain. Both active sites of the A- and C-domains are arrayed on the same side of the platform. The C-terminal lid region of the A-domain (100 residues of the Cterminal part) and the PCP domains are tethered to each other on top of the platform and connected to the large N-terminal region of A by a flexible linker of 15 residues. In this arrangement, PCP and A-lid region can easily move relative to the static C A-platform. In contrast to the compact C A-PCP domain association, the terminal TE domain builds an independent fold connected through a short (9 residues) linker to PCP and essentially shows an independent a/b-fold, identical to that of the dissected TE-domains (Marahiel 2009).

3.3

The Hybrid PKS/NRPS Complex Involved in Iturin Biosynthesis

Contrary to surfactin and fengycin, iturin derivatives are synthesised by a PKS NRPS hybrid complex. The operon consists of four ORFs called fenF, mycA, mycB and mycC for mycosubtilin (Duitman et al. 1999), ituD, ituA, ituB and ituC for iturin (Tsuge et al. 2001a) and bmyD, bmyA, bmyB and bmyC for bacillomycin D (Hofemeister et al. 2004; Koumoutsi et al. 2004). The last three genes encode for the three NRPSs, which are responsible for the incorporation of the first residue (Asn for MycA, ItuA and BmyA), the following four residues (Tyr, Asn, Gln, Pro for MycB and ItuB; Tyr, Asn, Pro, Glu for BmyB) and the two last residues (Ser, Asn for MycC; Asn, Ser for ItuC and Ser, Thr for BmyC). The thioesterase present in the last module catalyses the release of the peptide and the formation of an amide bond between the carboxylic group of the last amino acid and the amino group of the fatty acid chain. Epimerisation domains were identified in modules 2, 3 and 6. D-form of the corresponding amino acid residues is observed in the final product. The difference between structures of iturin A and mycosubtilin in which the last amino acids are inverted can be explained by an intragenic domain exchange between mycC and ituC. The synthetase of bacillomycin D is similar to that of iturin A except for the amino acid residues activated by modules 4, 5 and 7, which are Pro, Glu and Thr, respectively. FenF (ituD) encodes a malonyl-CoA transacylase (MCT-domain), and the mycA also contains genes related to PKS. These genes are responsible for the last steps of the biosynthesis of the fatty acid chain [last elongation and b-amination (Aron et al. 2005)] before its transfer to the first amino acid of the peptidic moiety [acyl-CoA ligase (AL-domain), acyl carrier protein (ACPdomain), b-keto acyl synthetase (KS-domain) and amino transferase (AMT domain)]. Hansen et al. (2007) have shown that the AL-domain is able to activate

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free fatty acids through an acyl-adenylate intermediate and loaded on the adjacent thiolation domain independently of co-enzyme A.

3.4

Non-ribosomal Peptide Synthesis of Fengycin and Plipastatin

Fengycin or plipastatin are synthesised by five NRPSs (Fen1 to Fen5) encoded by an operon with five ORFs fenA-E (or ppsA-E) (Tosato et al. 1997; Steller et al. 1999; Koumoutsi et al. 2004). The first three enzymes, Fen1, Fen2 and Fen3 contain two modules, the fourth contains three modules and the last enzyme consists of one module. Fen1 activates and incorporates glutamate in position 1 and ornithine in position 2, which form the side chain of the peptidic moiety of fengycin. Fen2 is responsible for the activation and incorporation of tyrosine in position 3 and allothreonine in position 4. Fen3 activates and incorporates glutamate in position 5 and alanine or valine in position 6. Fen4 is a three modular enzyme, which catalyses the activation and incorporation of proline in position 7, glutamine in position 8 and tyrosine in position 9, and Fen5 allows the incorporation of the last amino acid residue: isoleucine in position 10. Like in surfactin biosynthesis, the b-hydroxylated fatty acid chain is added to the amino acid activated in the first module by the way of a starter condensation domain. The thioesterase present in the last module catalyses the release of the peptide and the formation of an ester bond between the carboxylic group of the last amino acid (Ile) and the hydroxyl group of a tyrosine in position 3. Epimerisation domains were identified in modules 2, 4, 6 and 9. D-form of the corresponding amino acid residues is observed in the final product. This depicts the structure of the plispastatin; however, it does not correlate with fengycin produced by Bacillus subtilis S499. In the latter case, a D-Tyr is present in position 3 of the peptide and an L-Tyr in position 9 (Schneider et al. 1999). However, the organisation of the fengycin operon, which was not yet described for this strain could be different.

3.5

The Recent Discovery of the Biosynthesis Mechanism of Kurstakin

Two new approaches were recently developed to detect new NRPS genes responsible for the biosynthesis of lipopeptide compounds. The first being a PCR approach using degenerated primers based on the intraoperon alignment of adenylation and thiolation nucleic acid domains of all enzymes implicated in the biosynthesis of each lipopeptide family (Tapi et al. 2010). For the second approach (Bumpus et al. 2009), the authors took advantage of the size of the NRPS enzymes and the presence of unique marker ions derived from the common phosphopantetheinyl cofactor to adapt mass spectrometry-based proteomics to selectively detect NRPS and PKS gene clusters in microbial proteomes without requiring genome sequence

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information. In both cases, authors discovered in Bacillus thuringiensis (Abderrahmani et al. 2010) and Bacillus cereus the genes involved in the biosynthesis of the kurstakin. The exponential growth of genome sequences and the development of such interesting tools to detect or predict potentially novel NRPs highlight the interest of a database, such as NORINE, which collects all the known structures of NRPs and provides software for efficient structure comparison (Caboche et al. 2009). A good example is the pseudo-discovery of bacillorin, the synthetase sequences of which have been deposited in GenBank since 2008. The peptide was assumed to be a novel compound synthesised by NRPS. However, it was demonstrated to be the bacillomycin L, based on predictive amino acid incorporation followed by structural pattern comparison with all peptides annotated in the NORINE database (Lecle`re, personal communication). Bacillorin and bacillomycin L should then be considered as synonymous names for a single molecule.

3.6

Conclusion

Up to now, all the lipopeptides produced by Bacillus spp. are synthesised by NRPS. These modular enzymes have been especially well studied in the case of surfactin. These “megaenzymes” exhibit a broad spectrum of activities, conferring them an extraordinary potential for the development of bioprocesses, which lead to the biosynthesis of novel compounds for pharmaceutical, agricultural and biotechnological sectors.

4 A Complex Regulation of the Biosynthesis In this section, the molecular mechanism of regulation of the expression of the operon involved in the biosynthesis of surfactin and other lipopeptides from Bacillus are first described. Figure 3 shows the cascade of regulation, which controls surfactin operon expression. The environmental factors, which mainly influence the production of the lipopeptides, are also discussed.

4.1

Quorum Sensing and Surfactin Efflux

Surfactin production is quorum sensing-dependent. Its regulation involves several pheromones ComX, PhrC, PhrF, PhrG and PhrH together with several pleiotropic regulators such as CodY, DegU and AbrB. These form a complex cascade governing multiple differentiation pathways, such as sporulation and competence (Hamoen et al. 2003; Hayashi et al. 2006). Cosby et al. (1998) also showed that surfactin synthetase expression is pH dependent. A surfactin-susceptible mutant

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Fig. 3 Regulatory cascade of surfactin operon expression. Bacteria are able to communicate with each other through the production of “quorum sensing” pheromones. Accumulation of these pheromones in the growth medium signals the presence of a sufficient number of congeners (a quorum) and triggers various cell density dependent processes. Expression of the surfactin operon is one of these processes. Two types of pheromones are involved in the regulation of srfA operon. The first pheromone is ComX, which is secreted with the help of ComQ in the culture medium. ComP senses the accumulation of the ComX, and at critical ComX concentration, phosphorylates itself. Subsequently, autophosphorylated ComP phosphor ylates ComA. ComA P activates expression of surfactin operon and ComS, which is an ORF encoded within de srfA mRNA. ComS leads to the autoactivation of ComK. In addition, several activators such as ComK itself and DegU and repressors including CodY, AbrB and Rok are involved in ComK activation. The second type of pheromones is the group of Phr peptides: PhrC, PhrF, PhrG and PhrH. They are probably first synthesised as small proteins, which are secreted, processed in pentapeptides and then internalised by an oligopeptide permease (Opp). The phrH gene constitutes an operon with a rapH gene. Both genes are thus activated by ComK. Products of phr and rap genes are interacting regulator factors that modulate the phosphoryla tion state or DNA specific response regulators. The phrC, phrF and phrH genes need the sH (Spo0H) form of RNA polymerase to be transcribed. RapC, RapF and, in certain circumstances, RapG and RapH, are inhibitors of the DNA binding of ComA P. ComA P activates rapC and rapF. PhrC, PhrF, PhrG and PhrH inhibit their cognate Rap proteins. RapG and RapH inhibit DNA binding of DegU. RghR is a rapG, rapH and phrH repressor

was obtained by transposon mutagenesis. Genetic analysis revealed yerP, a gene that would be involved in surfactin self resistance. It has homology with resistance, nodulation and cell division (RND) family proton motive force-dependent efflux pumps. It should be responsible for surfactin efflux. Its expression is highest at the end of logarithmic phase (Tsuge et al. 2001b). Duitman et al. (2007) have shown recently that mycosubtilin synthetase expression is under the influence of a regulatory cascade depending on AbrB, one of the main transition state regulators in B.

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subtilis. In contrast to surfactin, the expression of mycosubtilin synthetase is lowest in minimal medium and highest in rich medium. We recently demonstrated that mycosubtilin synthetase expression is oxygen-dependent (Guez et al. 2008). To our knowledge, little information is available about the regulation mechanism of fengycin or plipastatin biosynthesis. However, the introduction of a pleiotropic regulator DegQ in Bacillus subtilis 168 results in a tenfold increase in the production of plipastatin (Tsuge et al. 1999).

4.2

Influence of Environmental Factors

A lot of studies have pointed out different environmental factors for their effect on surfactin production (Peypoux et al. 1999; Akpa et al. 2001). Several experiments performed in our laboratory have shown that this effect can be strain dependent. Regarding the carbon source, glucose was the most used substrate. Saccharose and fructose have also been mentioned as efficient carbon sources contrary to glycerol and hexadecane. The two main culture media mentioned in literature for surfactin production are Cooper’s medium, which contain mineral nitrogen source (NH4NO3) (Cooper et al. 1981), and Landy’s medium (Landy et al. 1948), with glutamic acid as a nitrogen source. Studies of the mineral requirement clearly established the need and the stimulatory effect of iron and manganese (Wei et al. 2004). For continuous operation, a critical nitrogen/iron/manganese molar ratio of 920:7.7:1.0 was determined and was found to sustain surfactin production for at least 36 generations (Sheppard and Cooper 1991). Oxygen and temperature are also considered as important parameters. Higher temperatures (37 C) favoured surfactin synthesis of Bacillus subtilis RB14 isolated from compost (Ohno et al. 1995a) and ATCC6633 but not of Bacillus subtilis S499 (Jacques et al. 1999). The replacement of Cooper’s nitrogen source and the introduction of oxygen limitation, which redirects the energy flux into product synthesis, have led to the highest productivity mentioned for surfactin production (7 g l 1) by Bacillus subtilis C9 (Kim et al. 1997). Such a process appears not to be adapted to Bacillus subtilis S499, which produced higher surfactin yield in better aeration conditions (Hbid et al. 1996; Jacques et al. 1999). Production of lichenysin by Bacillus licheniformis was conducted in anaerobic conditions and mainly at higher temperatures (up to 45 C) than for surfactin. Presence of salt (NaCl 5%) was also sometimes required (Yakimov et al. 1995). However, production yields were in average, five to ten times lower than with surfactin (Yakimov et al. 2000). A 30-fold increase in mycosubtilin production was observed when the temperature was decreased from 37 to 25 C. This was observed both for strain ATCC6633 and its derivative BBG100, which is a constitutive mycosubtilin over-producer. However, no significant difference in either the expression of the mycosubtilin synthetase encoding genes or in the intracellular synthetase concentration could be

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found, suggesting that the observed phenotype originated from a higher mycosubtilin synthetase turnover at lower temperature (Fickers et al. 2008). Recently, our laboratory demonstrated that environmental conditions can drastically modify the ratio between surfactin and fengycin produced by a derivative strain of Bacillus subtilis ATCC21332. Presence of polypropylene carriers in a batch culture (Gancel et al. 2009) or use of a polypropylene membrane (Coutte et al. 2010a) to aerate bioreactors (bubbleless process see Sect. 6) increases the biosynthesis of fengycin.

4.3

Conclusion

Many of the physico-chemical parameters influence the biosynthesis of lipopeptides from Bacillus spp. This is certainly due to the complex regulation of lipopeptide operon expression involving several pleiotropic regulators. However, intracellular pools of synthetase substrates (amino acid residues and fatty chain) and turn-over of the enzymes have also to be taken into account. To our knowledge, a limitation of lipopeptide secretion was never described in literature.

5 Physico-chemical Properties and Biological Activities: A Strong Relationship The content of this section is a summary of the main physico-chemical properties of surfactin, iturin and fengycin compounds, biological activities and their possible relationships. The mechanisms of action, where known, are also described. To end with, the multifunctional role of lipopeptide in plant protection against phytopathogen is also discussed.

5.1

Surfactin: A Potent Biosurfactant, Which Combines High Effect on Surface Tension and Low Critical Micellar Concentration

Surfactins are powerful biosurfactants with exceptional emulsifying and foaming properties (Razafindralambo et al. 1996). They are able to reduce surface tension of water to 27 mN m 1 and show a low critical micellar concentration (CMC) of about 10 mg l 1. Due to their amphiphilic nature, surfactins can also readily associate and tightly anchor into lipid layers. It can thus interfere with biological membrane integrity in a dose-dependent manner. The effect of surfactin on artificial membrane was studied in detail on different models using atomic force microscopy (Deleu et al. 1999a, 2001;

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Francius et al. 2008), light scattering, differential scanning calorimetry, small-angle neutron scattering and cryo-electron microscopy (Kell et al. 2007). Studies on lipid vesicles suggest that at low concentration (surfactin-to-lipid mole ratio Rb lower than 0.04 in the membrane), surfactins insert exclusively in the outer leaflet of the membrane inducing only limited perturbation. At intermediate concentration (Rb 0.05 0.1), it provokes a transient permeabilisation, but membranes re-anneal. Irreversible pore formation then occurs at higher ratio (Rb 0.1 0.2) due to the insertion of surfactinrich clusters in the membrane. Further addition of surfactins to reach the CMC leads to complete disruption and solubilisation of the lipid bilayer with formation of mixed micelles (Rb 0.22) (Heerklotz and Seelig 2007; Carrillo et al. 2003). Interestingly, the presence of cholesterol in the phospholipid layer attenuates the destabilising effect of surfactins, which suggests that the susceptibility of biological membranes may vary in a specific manner, depending on the sterol content of the target organisms. This could explain why surfactins display haemolytic, antiviral (Kracht et al. 1999), antimycoplasma (Vollenbroich et al. 1997) and antibacterial activities but, intriguingly, no marked fungitoxicity. However, recent studies have shown that surfactin could induce plant systemic resistance (Ongena et al. 2007; Jourdan et al. 2009). The molecular mechanism of this induction is not yet known but could also involve interaction with the membrane of plant cells. The lipid bilayer destabilisation process, observed with membrane models, is facilitated by the tri-dimensional form of the surfactin molecule. In solution, the peptide moiety of surfactin shows a “horse-saddle” topology. Its two negatively charged amino acid residues (Asp and Glu) form a claw and function as binding sites for mono- and divalent cations, while its fatty acid chain is extending at the opposite side of the ring. In membranes, charged side chains are protruding into the aqueous phase and apolar moieties reaching into the hydrophobic core of the membrane (Deleu et al. 2003). The use of chemically modified or biosynthetic variants has revealed prominent roles for some sub-structures (Morikawa et al. 2000). The esterification of Glu residue in surfactin increases its surface active power. The association properties of the lipopeptide increase with the diminution of their anionic charges. Surfactin mono methyl-ester has higher haemolytic activity than surfactin. But linear surfactin is less haemolytic (Dufour et al. 2005). The emulsification properties of surfactin were also used to solubilise xenobiotic compounds (Lai et al. 2009) and enhance their biological degradation (Whang et al. 2009) or to facilitate oil recovery from carbonate reservoirs (Zhang et al. 2000). When linear surfactin was prepared by saponification of the lactone ring, its oil displacement activities decreased to one-third of their respective original value (Morikawa et al. 2000). Several studies have also shown the role played by surfactin in pellicle formation of the producing Bacillus strain at the air-water interface (Hofemeister et al. 2004; Chollet-Imbert et al. 2009), in swarming (Julkowska et al. 2005; Debois et al. 2008) or in biofilm formed on roots (Bais et al. 2004). At concentrations between 5 and 50 mg l 1, surfactin inhibits Salmonella biofilm formation in microtiter plates and urethral catheter (Mireles et al. 2001),and Pseudomonas syringae growth on roots (Bais et al. 2004). The adsorbtion of surfactin on Stainless steel or Teflon substrata deeply modified the hydrophobicity of the surface and the Bacillus cereus spore attachment

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(Shakerifard et al. 2009). Surfactin can also modify the surface hydrophobicity of the producing strain (Ahimou et al. 2000). Neutron reflectometry was recently used to study the structure of surfactin at the air-water and hydrophobic solid/water interfaces and on surfaces. Surfactin was found to adopt a ball-like structure with a thickness of ˚ and an area per molecule of 147  5 A ˚ 2 (Shen et al. 2009). The structure of 14  1A surfactin micelles has been examined by means of small-angle neutron scattering. The aggregation number was found to be unusually small at 20  5. Furthermore, anti-inflammatory (Kim et al. 1998), anticancer activity (Kameda et al. 1974) and immunomodulatory effects (Park and Kim 2009) were also determined for surfactin. Cao et al. (2009) showed that surfactin induces apoptosis and G2/M arrest in human breast cancer MCF-7 Cells.

5.2

Iturin: A Strong Antifungal Compound

Iturin is also a biosurfactant but less potent than surfactin (Deleu et al. 1999b). It reduces the surface tension of water to 43 mN m 1 and forms at CMC (about 20 mg l 1) micelles with a Stokes radius of 1.3 nm and an aggregational number of 7. At concentrations slightly higher than CMC, iturin probably forms a fully interdigitated bilayer where each hydrocarbon tail spans the entire hydrocarbon width of the bilayer, resulting in multilamellar vesicles with an average size of 150 nm (Grau et al. 2001). Though they are also strongly haemolytic, the biological activity of iturins differs from surfactins: they display a strong in vitro and in vivo antifungal action against a large variety of yeast and fungi but have only limited antibacterial and no antiviral activities (Lecle`re et al. 2005; Mizumoto et al. 2007; Romero et al. 2007; Fickers et al. 2009). This fungitoxicity of iturins almost certainly relies on their membrane permeabilisation properties. However, the underlying mechanism is based on osmotic perturbation due to the formation of ion-conducting pores and not membrane disruption or solubilisation as caused by surfactins (Aranda et al. 2005). In addition to the direct activity against fungi, iturin derivatives enhance the invasive growth of the producing strain, and thus, by these two mechanisms participate in plant protection against phytopathogens (Lecle`re et al. 2005, 2006). The length of the fatty acid chain, the presence of an Asp in position 1 and tyrosine residue in position 2 are important structural traits for the antifungal activity of the lipopeptide (Bonmatin et al. 2003; Fickers et al. 2009).

5.3

Fengycin and Plipastatin: Immunomodulators in Plants and Animals

Fengycins are less haemolytic than iturins and surfactins but retain a strong fungitoxic activity more specifically against filamentous fungi. Mechanistically,

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the action of fengycins is less known compared to other lipopeptides, but they also readily interact with lipid layers and somewhat retain the potential to alter cell membrane structure (packing) and permeability in a dose-dependent way. Immunosuppressive activity of plipastatin was also described and patented (Umezawa et al. 1988). Lipopeptides are known to act in a synergistic manner as suggested by several studies, on surfactin with iturin (Maget-Dana et al. 1992; Razafindralambo et al. 1997), surfactin with fengycin (Ongena et al. 2007) and iturin with fengycin (Romero et al. 2007).

5.4

Lipopeptides: Versatile Weapons for Biocontrol of Plant Diseases

In the context of biocontrol of plant diseases, the three families of Bacillus lipopeptides (surfactins, iturins and fengycins) were at first mostly studied for their antagonistic activity against a wide range of potential phytopathogens including bacteria, fungi and oomycetes. Recent investigations have shed light on the fact that these lipopeptides can also influence the ecological fitness of the producing strain in terms of root colonisation and thereby persistence in the rhizosphere, also playing a key role in the beneficial interaction with plants by stimulating host defence mechanisms (Ongena et al. 2005, 2007). The different structural traits and physico-chemical properties of these effective surface- and membrane-active amphiphilic biomolecules explain their involvement in most of the mechanisms (Ongena and Jacques 2008).

5.5

Conclusion

The large panel of physico-chemical properties and biological activities of lipopeptides from Bacillus has to be correlated to their remarkable structure diversity. They offer them various potential applications. Their complementarity and the observation of some synergistic effects entice interest to further study the different families.

6 New Strategies for an Optimal Production of Novel or Existing Lipopeptidic Compounds In this section, we first discuss the molecular strategies used to achieve higher yield of lipopeptide biosurfactants or to obtain modified compounds. Then we give a brief overview of the different techniques used to produce, purify and quantify the lipopeptide from Bacillus subtilis.

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79

Surfactin Synthetases Re-engineering

Surfactin synthetase was the first NRPS for which a genetic re-engineering experiment was reported. It concerns the terminal module, which incorporates leucine in the natural system and exhibits the domain composition C-A-PCP-TE. The activating and covalent attachment domains (A-PCP) were exchanged by A-PCP units from bacterial and fungal origin with various amino acid specificities. Novel surfactin variants with aliphatic (Val), charged (Orn) and aromatic (Phe) residues at position 7 were created and their structure was confirmed by mass spectrometry (Stachelhaus et al. 1995). All these new variants displayed the same haemolytic activity as the native surfactin. However, low yields of the peptide products (0.1 0.5% in comparison to the parent strain) were observed probably due to the high selectivity of C domains in the acceptor site for the cognate amino acid substrate. Different results were obtained with deletion of the Leu incorporating module 2. Initially unsuccessful, a second trial yielded the predicted surfactin deprived of the second Leu residue in about 10% yield in comparison to the parent strain (Mootz et al. 2002). This highlighted the importance of precise linker surgery. The recombination of whole modules represents a rather drastic intervention in NRPS biosynthesis, which usually results in reduced catalytic efficiency and product yield. However, Yakimov et al. (2000) report the entire replacement of module 1 and 5 of surfactin synthetase by those of lichenysin synthetase, to create a fully active hybrid enzyme that forms lichenysin-like biosurfactant in high yields. A more conservative strategy involves manipulating the A domain’s specificity through point mutations affecting the substrate-coordinating amino acid residues according to the “non-ribosomal code” (see A-domain section). For example, the substrate specificity of the Glu-activating module 1 of surfactin synthetase was rationally altered in this way. A single point mutation changed the specificity from Glu to Gln without a decrease in catalytic efficiency. A second mutation changing the specificity of module 5 from Asp to Asn yielded the expected surfactin derivative in vivo (Eppelmann et al. 2002).

6.2

Combinatorial Synthesis of Lipopeptides

Up until now, corresponding approaches were generally limited to module swaps, which: (1) require major manipulations of the biosynthetic template, (2) were mostly connected with significant decrease in product titer, and (3) constitutionally lead to the synthesis of only one product per experiment. Chiocchini et al. (2006) show that a heterologous COM domain pair can replace the native COM domain pair. Indeed, the replacement of COMDSrfA-A/COMASrfA-B by COMDTycB-A/COMATycC; a cognate COM domain pair facilitating the interaction between the two NRPSs, TycB and TycC, involved in the biosynthesis of tyrocidine in Bacillus brevis only caused a minor decrease in surfactin production. Using different modifications of COM

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domain pairs, the authors also show that it was possible to skip the srfA-B domain and to generate in majority a lipotetrapeptide instead of the natural lipoheptapeptide.

6.3

Directed Biosynthesis and Molecular Optimisation

Bacillus strains frequently co-produce different families of lipopeptides (Jacques et al. 1999). Several strategies were thus proposed to selectively overproduce a family or only a variant of a family. Firstly obtained by random mutagenesis (Lin et al. 1998; Yoneda et al. 2002), overproducing mutants were then constructed by promoter exchange. The four main promoters used are PrepU (Tsuge et al. 2001a; Lecle`re et al. 2005; Coutte et al. 2010a), Pxyl (Fickers et al. 2009), Pspac (Sun et al. 2009) and PamyQ (Ongena et al. 2007). Disruption mutants were also obtained by introducing resistance genes into synthetase operons (Coutte et al. 2010a). Lin et al. (1998) isolated a Bacillus licheniformis mutant derived by random mutagenesis with N-methyl-N9-nitro-N-nitrosoguanidine, which produces 391 mg l 1 of lichenysin. Using the same approach, Yoneda et al. (2002) described in a patent, a production yield of up to 50 g l 1 of surfactin. The replacement of the native promoter of iturin (Tsuge et al. 2001a) and mycosutilin (Lecle`re et al. 2005) operons by PrepU led to an overproduction of respectively 3 and 15 times of the lipopeptide to reach a concentration of about 200 mg l 1 in both cases. Performed on surfactin synthetase, the same promoter exchange did not allow the improvement of the surfactin production (Coutte et al. 2010a). The native promoter Psrf of Bacillus subtilis 168 is considered very efficient, frequently allowing the concentration of surfactin in the culture supernatant to be higher than 1 g l 1 (Duitman et al. 2007). The horizontal transfer of iturin operon in Bacillus subtilis 168 together with a functional sfp gene and the pleiotropic regulator degQ allowed conversion of this strain to an efficient iturin producer (Tsuge et al. 2005). Recently, the combined use of low temperature, isoleucine addition and overproducing mutant strains with Pxyl instead of Pmyc driving the mycosubtilin operon allowed us to synthesise more than 900 mg l 1 of the most active mycosubtilin isoform against Candida sp. (Fickers et al. 2009).

6.4

New Bioprocesses for Continuous Production and Extraction of Lipopeptides

The different strategies that have been used for the fermentation process and extraction methods for the production of lipopeptides are discussed. 6.4.1

Quantitative Evaluation of Surfactin Concentrations

Several indirect techniques based on surfactin properties, such as measurement of surface tension or haemolytic activities (Jacques et al. 1999), are used for

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quantitative evaluation of surfactin. These techniques are mainly dependent on the degree of purity of the tested samples and can thus only give a qualitative idea about the presence of surfactant or relative idea about their concentrations. Analysis by high performance liquid chromatography is the main accurate technique for surfactin evaluation, but it needs to be carefully conducted (Razafindralambo et al. 1993). As surfactin precipitates at pH below 6, samples should be stored at higher pH. In several cases, high salt or organic compound concentrations can also lead to precipitation during sample freeze-thawing. If pre-step purification by solid-phase extraction is recommended, its working conditions depend of the nature of the precolumn used, and it has to be carefully tested before being routinely used to avoid the partial loss of the surfactin present in the samples.

6.4.2

Purification of Surfactin

Surfactin was first extracted from culture broth by acidic precipitation (HCl) followed by extraction with methanol or other organic solvents. Purification was completed by chromatographic procedures (on silica gel or by reversed phase chromatography). Other methods were then proposed to purify and concentrate the lipopeptide: a two-phase extraction (Drouin and Cooper 1992), an ultrafiltration method (Mulligan and Gibbs 1990; Isa et al. 2008), a solid-phase extraction (Razafindralambo et al. 1993; Montastruc et al. 2008), a liquid membrane extraction (Dimitrov et al. 2008) or different combined methods (Chen et al. 2007, 2008).

6.4.3

Bioprocesses for Lipopeptide Production

Three main types of bioprocesses were developed for lipopeptide production: solidstate fermentation, foam fractionation and membrane bioreactors. Solid state fermentation has been suggested for a long time by Japanese researchers, as Bacillus subtilis can easily grow on different food processing wastes. They showed a high level of surfactin production (2 g kg 1 of wet weight) (Ohno et al. 1995b). The technique of foam fractionation was first suggested by Cooper et al. (1981). It offers the double advantage of continuous in situ removal of produced surfactin from the fermentation broth together with the prevention of any possible feedback inhibition. It was then developed by Davis et al. (2001). We also applied this strategy for the production of mycosubtilin. An overflowing exponentially fed batch process (OEFBC) was so defined, which allowed a continuous extraction of the mycosubtilin with high efficiency (Guez et al. 2007). With the same idea of developing a continuous process for surfactin and fengycin extraction, a membrane bioreactor was tested. The use of a membrane to ensure the oxygen transfer in bubbleless conditions allowed to avoid foam formation, leading to the development of a membrane bioreactor with cell recycling by microfiltration and lipopeptide concentration by ultrafiltration (Coutte et al. 2010b).

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6.5

Conclusion

The actual level of knowledge of the biosynthesis of lipopeptide from Bacillus subtilis and its regulation mechanism allows to develop different strategies to overproduce the main active compounds and to reach yields that are compatible with industrial development of such compounds.

7 Industrial Applications: Dream and Reality In this section, we summarise the main potential applications of surfactin developed in literature and the commercially existing products. Figure 4 gives an overview of application sectors of surfactin.

7.1

Main Applications

Several recent reviews summarise the high interest of biosurfactant for different application fields (Singh et al. 2007): food (Nitschke and Costa 2007), petroleum recovery (Sen 2008), environmental (Cameotra and Makkar 2010; Mulligan 2009), biomedical (Rodrigues et al. 2006) and cosmetics (Kanlayavattanakul and Lourith 2010). During its long history, surfactin has been first studied for its potential BIOTECHNOLOGY Antiviral

PHARMACEUTICALS

PHYTOSANITARY Inducer of Systemic Resistance Antibacterial Antiviral Spreading-biofilm

Antiviral Antimycoplasma Antibacterial Anticancer Hypocholesterolemic Antithrombotic

SURFACTIN COSMETICS

FOOD Emulsifyer Biosurfactant

Emulsifyier Antibacterial

PETROLEUM INDUSTRY ENVIRONMENT

Oil extraction

Bioremediation of polluted soils

CHEMICALS Biosurfactant

Fig. 4 Application sectors and exploited properties of surfactin

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pharmaceutical applications (antibacterial, antitumoral and hypocholesterolemic activities). Its biosurfactant properties were also highlighted, and its application for example in petroleum industry was considered. The discovery at the end of 1990s of its antimycoplasma and antiviral properties led to the proposal of its use to ensure the safety of pharmaceutical or biotechnological products. The presence of several lipopeptides in different Asian (detection of several mg of surfactin/100 g of Natto wet weight, Sumi et al. 2000) or African (presence of mycosubtilin in Netetu, N’dir et al. 1994) fermented food products or its secretion by strains isolated from these products (Cho et al. 2009) allows for consideration of their application in the food sector. More recently, its ability to induce systemic resistance in plants and its implication in the spreading of the cells and thus the rhizosphere colonisation could open new fields of applications as phytopharmaceutical products (Ongena and Jacques 2008).

7.2

Conclusion

More than 50 publications published in 2009 are referenced by Scopus with the keyword “surfactin” showing the high interest in this compound, which remains the most studied lipopeptide produced by Bacillus spp. Despite this high number of scientific publications and patents, industrial surfactin applications still remain quite limited. Sold by SIGMA and SHOWA DENKO Co for analytical or laboratory purposes, the compound is also available in several Japanese cosmetics products. Production costs and also suspected toxicity (From et al. 2007) are probable reasons why these compounds are not yet commonly used. Recent development of overproducing strains and upscalable bioprocesses could solve the problem of cost efficiency. Toxicological and ecotoxicological studies are now needed to confirm the low toxicity (Hwang et al. 2009) of a compound which is probably consumed every day by thousands of people. Acknowledgements The author thanks Dr Vale´rie Lecle`re, Dr Max Bechet, Dr Franc¸ois Coutte, Ursula Collins and Deirdre Hallinan for their kind re reading of the manuscript and Isabelle Schack and Damien Jacques for help in the reference list and figure preparation. ProBioGEM is supported by the Universite´ des Sciences et Technologies de Lille, the Region Nord Pas de Calais, the Ministere de la Recherche Scientifique (ANR) and the European Funds for the Regional Development.

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Ahimou F, Jacques P, Deleu M (2000) Surfactin and iturin A effects on Bacillus subtilis surface hydrophobicity. Enzyme Microb Technol 27:749 754 Akpa E, Jacques P, Wathelet B, Paquot M, Fuchs R, Budzikiewicz H, Thonart P (2001) Influence of culture conditions on lipopeptide production by Bacillus subtilis. Appl Biochem Biotechnol 91 93:551 561 Aranda FJ, Teruel JA, Ortiz A (2005) Further aspects on the hemolytic activity of the antibiotic lipopeptide iturin A. Biochim Biophys Acta Biomembr 1713:51 56 Arima K, Kakinuma A, Tamura G (1968) Surfactin, a crystalline peptidelipid surfactant produced by Bacillus subtilis: isolation, characterization and its inhibition of fibrin clot formation. Biochem Biophys Res Commun 31:488 494 Aron ZD, Dorrestein PC, Blackhall JR, Kelleher NL, Walsh CT (2005) Characterization of a new tailoring domain in polyketide biogenesis: the amine transferase domain of MycA in the mycosubtilin gene cluster. J Am Chem Soc 127:14986 14987 Athukorala SNP, Fernando WGD, Rashid KY (2009) Identification of antifungal antibiotics of Bacillus species isolated from different microhabitats using polymerase chain reaction and MALDI TOF mass spectrometry. Can J Microbiol 55:1021 1032 Bais HP, Fall R, Vivanco JM (2004) Biocontrol of Bacillus subtilis against infection of Arabi dopsis roots by Pseudomonas syringae is facilitated by biofilm formation and surfactin. Plant Physiol 134:307 319 Besson F, Peypoux F, Michel G, Delcambe L (1977) The structure of bacillomycin L, an antibiotic from Bacillus subtilis. Eur J Biochem 77:61 67 Bonmatin JM, Labbe´ H, Grangemard I, Peypoux F, Maget Dana R, Ptak M, Michel G (1995) Production, isolation and characterization of [Leu4] and [Ile4] surfactins from Bacillus sub tilis. Lett Pept Sci 2:41 47 Bonmatin JM, Lapre´vote O, Peypoux F (2003) Diversity among microbial cyclic lipopeptides: iturins and surfactins. Activity structure relationships to design new bioactive agents. Comb Chem High Throughput Screen 6:541 556 Bumpus SB, Evans BS, Thomas PM, Ntai I, Kelleher NL (2009) A proteomics approach to discovering natural products and their biosynthetic pathways. Nat Biotechnol 27:951 956 Caboche S, Pupin M, Lecle`re V, Fontaine A, Jacques P, Kucherov G (2008) NORINE: a database of nonribosomal peptides. Nucleic Acids Res 36:D326 D331 Caboche S, Pupin M, Lecle`re V, Jacques P, Kucherov G (2009) Structural pattern matching of nonribosomal peptides. BMC Struct Biol 9:15 Cameotra SS, Makkar RS (2010) Biosurfactant enhanced bioremediation of hydrophobic pollu tants. Pure Appl Chem 82:97 116 Cao X, Wang AH, Jiao RZ, Wang CL, Mao DZ, Yan L, Zeng B (2009) Surfactin induces apoptosis and G(2)/M arrest in human breast cancer MCF 7 cells through cell cycle factor regulation. Cell Biochem Biophys 55:163 171 Carrillo C, Teruel JA, Aranda FJ, Ortiz A (2003) Molecular mechanism of membrane permeabi lization by the peptide antibiotic surfactin. Biochim Biophys Acta 1611:91 97 Chen HL, Chen YS, Juang RS (2007) Separation of surfactin from fermentation broths by acid precipitation and two stage dead end ultrafiltration processes. J Membr Sci 299:114 121 Chen HL, Chen YS, Juang RS (2008) Recovery of surfactin from fermentation broths by hybrid salting out and filtration process. Sep Purif Technol 59:244 252 Chiocchini C, Linne U, Stachelhaus T (2006) In vivo biocombinatorial synthesis of lipopeptides by COM domain mediated reprogramming of the surfactin biosynthetic complex. Chem Biol 13:899 908 Cho KM, Math RK, Hong SY, SMd AI, Mandanna DK, Cho JJ, Yun MG, Kim JM, Yun HD (2009) Iturin produced by Bacillus pumilus HY1 from Korean soybean sauce (kanjang) inhibits growth of aflatoxin producing fungi. Food Control 20:402 406 Chollet Imbert M, Gancel F, Slomianny C, Jacques P (2009) Differentiated pellicle organization and lipopeptide production in standing culture of Bacillus subtilis strains. Arch Microbiol 191:63 71

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Serrawettins and Other Surfactants Produced by Serratia Tohey Matsuyama, Taichiro Tanikawa, and Yoji Nakagawa

Contents 1 2

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94 Detection and Analysis of Serratia Biosurfactants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94 2.1 S. marcescens Wetting Activity and the Ways to Identify It . . . . . . . . . . . . . . . . . . . . . . . . . 95 2.2 Other Serratia Wetting Agents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 3 Production Characteristics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 3.1 Association of the Production of Serratia Surfactants with Extracellular Vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 3.2 Budding of Vesicles Filled with Serratia Surfactants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 4 Chemical Structure of Serrawettins and Rubiwettins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 4.1 Serrawettin W1, W2, and W3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 4.2 Rubiwettin R1 and RG1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 5 Physiological Functions of Biosurfactants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 5.1 Flagellum Independent Bacterial Spreading Growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 5.2 Flagellum Dependent Bacterial Spreading Growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 5.3 Other Biological Activities of Serrawettins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 6 Genetics of Biosynthesis and Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 6.1 Genes Involved in Serrawettin W1 Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 6.2 Quorum Sensing Regulation of Serrawettin Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . 113 6.3 Global Regulator Genes Concerned with Exolipid Production . . . . . . . . . . . . . . . . . . . . . 114 7 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117

Abstract Serrawettins are nonionic biosurfactants produced by Serratia marcescens. Three molecular species, serrawettin W1, cyclo(D-3-hydroxydecanoyl-L-seryl)2; W2, D-3-hydroxydecanoyl-D-leucyl-L-seryl-L-threonyl-D-phenylalanyl-L-isoleucyl lactone; and W3, cyclodepsipeptide composed of five amino acids and one T. Matsuyama (*) Niigata University Graduate School of Medical and Dental Sciences, Niigata 951 8510, Japan e mail: hy5s mtym@asahi net.or.jp T. Tanikawa and Y. Nakagawa Faculty of Agriculture, Niigata University, Niigata 950 2181, Japan

G. Sobero´n‐Cha´vez (ed.), Biosurfactants, Microbiology Monographs 20, DOI 10.1007/978 3 642 14490 5 4, # Springer Verlag Berlin Heidelberg 2011

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dodecanoic acid, have been reported. Serratia rubidaea produces rubiwettin R1, linked D-3-hydroxy fatty acids and RG1, b-glucopyranosyl linked D-3-hydroxy fatty acids. These biosurfactants are produced mainly at 30
C, but not at 37
C, and secreted through extracellular vesicles on solid media. The contribution of the biosurfactants to spreading growth in surface environments has been determined, and it is prominent under nutrient-poor conditions. Analyses of S. marcescens mutants revealed the involvement of three novel genes for serrawettin W1 production. The gene pswP encodes a phosphopantetheinyl transferase group enzyme, swrW encodes a unimodular synthetase belonging to the nonribosomal peptide synthetase (NRPS) family, and hexS encodes a LysR-type transcriptional regulator working as a downregulator of Serratia exolipids and some exoenzymes. Autoinducer-dependent serrawettin W2 production has been elucidated by the finding of SwrI/SwrR (homolog of LuxI/LuxR) and N-acyl homoserine lactones in the study on quorum-sensing controlled-swarming of S. marcescens.

1 Introduction Surfaces in nature are the favorite environment for most microbes. In addition, various novel kinds of surfaces are being developed as part of medical and industrial activities and these are becoming niches for microbes (Costerton and Lewandowski 1995). Although solid surfaces provide a stable foothold for microbes to generate an active population with intimate cell-to-cell interactions, microbes may be inactive on completely dry surfaces. Every organism on earth depends on water for its existence. However, the surface tension of water is strong enough to restrict microorganisms, and it seems quite difficult for microorganisms to escape beyond the air water interface. On the other hand, microbes easily adsorb on to wet surfaces, which may be convenient in establishing a nonspecific microbial lodging on various surfaces. In this chapter, the general feature of serrawettins, one of the main types of the surfactants produced by Serratia, will be described. Then, the working mode of these biosurfactants on surface environments and the genetics of their biosynthesis in response to bacterial population behavior will be presented. For the coexistence of human beings with microbes, it is important for us to understand microbial strategies that arise in the bacterium-scale surface world.

2 Detection and Analysis of Serratia Biosurfactants Biosurfactant production seems to be of general importance for different types of bacteria, especially when these bacteria colonize surfaces. This is one of the reasons for the initial studies on serrawettins, the biosurfactants produced by Serratia

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marcescens. Serrawettins can be detected and analyzed by different techniques. The description of these methodologies is the aim of this section.

2.1

S. marcescens Wetting Activity and the Ways to Identify It

The rapid and spontaneous development of a dendritic smear pattern on the surface of a glass slide prepared for Gram staining was the first sign of wetting activity of S. marcescens (Fig. 1a). In contrast, such a dynamic phenomenon was not observed with Escherichia coli (Fig. 1b). The phenomenon seemed to suggest the production of biosurfactants by Serratia (Matsuyama et al. 1985). Thereafter, for the detection of the bacterial wetting activity, a bacterial suspension was made on a glass slide as the routine procedure. Thereafter, wetting activities of S. marcescens on various surfaces in nature were examined, for example, on surfaces of cotton flowers and a pea pod (Fig. 2). These results may support the indication with phytopathogenic Corynebacterium by Akit et al. (1981) with respect to its broader accessibility to water-shedding surfaces. For quantitative examination of the wetting activity, the glass surface was inadequate due to too delicate and complicated results to measure the wetted area (Fig. 1a). To avoid this problem, a polystyrene surface was used for measuring contact angles of a still droplet of a bacterial suspension on the hydrophobic surface. Table 1 shows quantitative wetting activities, which were consistent with the morphological findings shown in Figs. 1 and 2 (Matsuyama et al. 1986). This method was widely used for the rapid detection of bacterial biosurfactants

Fig. 1 Wetting activity of a bacterial suspension on a glass slide (high grade cleanness). (a) Spreading wetting by S. marcescens NS 38; (b) Nonspreading wetting by E. coli K 12S (from Matsuyama et al. 1985)

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Fig. 2 Suspension of S. marcescens placed on a water repelling plant surface. (a) On cotton flowers, NS 38 wild type grown at 37
C (1 left) and at 30
C (1 middle), NS 38 09 serrawettin deficient mutant grown at 30
C (2). (b) On a pea pod, NS 38 (1) and NS 38 09 (2) grown at 30
C (from Matsuyama and Nakagawa 1996a, b)

Table 1 Contact angles of droplets of cell supernatants on a polystyrene surface

Growth temp. (
C)a Contact angle (y)b 82.0  1.6 30 76.8  2.2 37 76.0  1.1 S. marcescens NS 38 30 19.8  0.4 37 69.4  1.0 S. marcescens NS 25 30 21.8  0.5 37 69.6  0.9 S. marcescens NS 45 30 17.9  0.5 37 66.6  0.9 a The strains were grown for 3 days on a PG agar medium at the indicated temperature. Supernatants (2 ml) of the bacterial sus pensions (10 mg/ml) were spotted on a polystyrene surface b Obtained by the method of Mack (1936); values are means of 10 determinations  SEM From Matsuyama et al. (1986) Bacterial strain Saline (control) E. coli K12 S

(Bar-Ness et al. 1988) and was also referred to as the “drop-collapsing test” (Jain et al. 1991). Conventional practical assays with a canvas sheet indicated that the wetting activity of S. marcescens was bacterial concentration- and cultivation temperature-dependent (Table 2).

2.1.1

Analysis of Serratia Lipids by Thin-Layer Chromatography

Lipid extracts from bacterial masses of Serratia strains grown on agar media were examined by thin-layer chromatography (TLC). As seen in Fig. 3, many spots of characteristic Serratia lipids appeared in addition to the usual bacterial phospholipid spots (Matsuyama et al. 1986). Among these specific spots, a spot of the waterinsoluble pigment prodigiosin, characteristic of Serratia, was identified easily by its red color. With regards to other specific lipids of S. marcescens, three dense spots (named W1, W2, and W3) on the upper part of the chromatograms were novel and

Serrawettins and Other Surfactants Produced by Serratia Table 2 Canvas sheet wetting activity of bacterial suspensionsa

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S. marcescens strain NS 38 NS 38 09 37
C 30
C Cultivation temperature 30
C Cell concentration (mg/ml) 1.0 0.5 1.0 1.0 Sinking time (min) 9.1 17.1 >60 >60 Wetted area height (mm) 11.2 4.0 <0.5 <0.1 a This activity was examined by a standard method at 30
C. Briefly, a canvas disk was floated on the surface of a bacterial suspension, then the time until sinking was measured, or a canvas strip was held vertically and its lower end was touched on to the surface of the bacterial suspension, and then the height of the wet area was measured after 5 min

Fig. 3 Thin layer chromatograms of lipids extracted from a 30
C culture of S. marcescens strains NS 33 (lane 1), NS 45 (lane 2), NS 25 (lane 3), NS 13 (lane 4), ATCC 8100 (lane 5), ATCC 13880 (lane 6), NS 12 (lane 7), NS 38 (lane 8), and 933 (lane 9); S. liquefaciens strains NCTC 10442 (lane 10) and CDC 1284 54 (lane 11); and S. rubidaea strains ATCC 27593 (lane 12) and CDC 299 72 (lane 13). Each sample (50 mg) was developed with a solvent of chloroform/methanol/5 M ammonia (80:25:4, by vol.). P, prodigiosin; W1, serrawettin W1; W2, serrawettin W2; W3, serrawettin W3; R1, rubiwettin R1; RG1, rubiwettin RG1; PLs, phospholipids; OL, ornithine containing lipid; O, origin (from Matsuyama et al. 1986)

considered as candidates of wetting agents. With Serratia rubidaea, two spots were recognized as novel specific lipids (named R1 and RG1). For identification of the active bacterial product responsible for Serratia wetting activity, a temperature-dependent phenotype of this activity was used as a key indicator. As shown in Fig. 2a and Tables 1 and 2, bacterial suspension from a 37
C

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Fig. 4 Thin layer chromatograms of lipids extracted from S. marcescens NS 25 (lanes 1 and 2), NS 38 (lanes 3 and 4), ATCC 13880 (lanes 5 and 6); S. rubidaea ATCC 27593 (lanes 7 and 8). Odd numbers, 30
C culture; even numbers, 37
C culture. Abbreviations are the same as in legend to Fig. 3

culture did not show wetting activity. Wetting activity on the glass surface was also absent with a 37
C culture. Consequently, Serratia lipids produced at 30
C, but not at 37
C, were examined by TLC. A thin-layer chromatogram shown in Fig. 4 shows the spots of such characteristic lipids (P, W1, W2, R2, and RG1). P is a spot of the red pigment prodigiosin. Thus, W1 and W2 lipids were identified as candidate wetting agents produced by S. marcescens. An additional lipid, W3, was also selected as a candidate by the same approach (Matsuyama et al. 1985, 1992). In examinations of purified lipids using a glass slide, the three specific lipids of S. marcescens, W1, W2, and W3, showed strong wetting activity in contrast to prodigiosin. These three S. marcescens-specific lipids were produced from different strains and differed in their Rf values and considered as different wetting agents named serrawettin W1, W2, and W3.

2.1.2

Surface Activities of Isolated Serrawettins

The identified Serratia wetting agents are listed in Table 3 with their surface activities and representative producer strains. Serrawettin W1 was a common wetting agent produced by pigmented S. marcescens strains (ATCC 13380, 274, NS 38, etc). Serrawettin W2 and W3 were produced by nonpigmented

Serrawettins and Other Surfactants Produced by Serratia Table 3 Surface activities of purified Serratia wetting agents Wetting agent Source Surface tensiona (mN/m) Serrawettin W1 S. marcescens NS 38 32.2 S. marcescens 274 S. marcescens ATCC 13380 Serrawettin W2 S. marcescens NS 25 33.9 Serrawettin W3 S. marcescens NS 45 28.8 Rubiwettin R1 S. rubidaea ATCC 27593 25.5 Rubiwettin RG1 S. rubidaea ATCC 27593 25.8 a Means of six determinations, 10 mg/ml saline (from Matsuyama et al. 1986, 1990) b Means of ten determinations on a polystyrene surface (from Matsuyama 1993)

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Contact angleb (y) <29.3 <11.6 <9.4 ND <6.0

S. marcescens strains. S. marcescens strains producing more than one kind of serrawettin have not been recognized so far.

2.2

Other Serratia Wetting Agents

As already mentioned, among other Serratia spp., S. rubidaea strains were found to have wetting activity due to the production of two characteristic lipids (R1 and RG1), which were different from serrawettins in their Rf values as shown in Fig. 3. In addition, these two lipids were detected in the same lane of a thin-layer chromatogram, and RG1 was shown to be a glycolipid (Matsuyama et al. 1990). It was noteworthy that production of R1 and RG1 was also temperature-dependent. Therefore, R1 and RG1 were considered as candidate of wetting agents as was the case for serrawettins. Purified R1 and RG1 showed distinct wetting activities. Thus, these lipids were called rubiwettin R1 and RG1, and surface activities and representative source strains are described in Table 3.

3 Production Characteristics In this section, different aspects of the production of serrawettins and other surfactants produced by Serratia are discussed.

3.1

Association of the Production of Serratia Surfactants with Extracellular Vesicles

For the preparation of Serratia wetting agents, peptone-glycerol (PG) agar medium (Matsuyama et al. 1985) is routinely used. This solid medium guaranteed proficient production of the surfactants serrawettins and rubiwettins and of the pigment

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Fig. 5 Phase contrast micrographs of unstained S. marcescens NS 38. (a) 30
C culture. (b) 37
C culture. Bars, 5 mm (from Matsuyama et al. 1986)

prodigiosin. Liquid medium lacked such an ability to promote the production of these lipids. However, silica gel dispersed in the aqueous phase of PG liquid medium remarkably enhanced productions of serrawettins and prodigiosin (Yamashita et al. 2001), presumably by providing numerous solid microsurfaces in the culture fluid. Serrawettins were produced proficiently at 30
C (as high as 17% [serrawettins] and 8% [rubiwettins] of dry weight of the bacterial mass) but not at 37
C. Prodigiosin in both species (S. rubidaea was also producer) was also produced in the same temperature-dependent manner. In parallel with this lipid production, the temperature-dependent generation of extracellular vesicles from S. marcescens and S. rubidaea was recognized (Matsuyama et al. 1986, 1990). In particular, 30
C-grown bacterial masses contained many extracellular vesicles of various sizes (diameter, less than 3.0 mm) in contrast to 37
C-grown bacteria (Fig. 5). It was noteworthy that the extracellular vesicles of pigmented strains were red, but the bacterial cells themselves were colorless when examined using a bright-field microscope. The extracellular vesicles of prodigiosin nonproducing strains were colorless. These findings suggested that Serratia wetting agents and red pigment are exolipids and the main lipid components of the extracellular vesicles.

3.2

Budding of Vesicles Filled with Serratia Surfactants

Scanning and transmission electron microscopy showed that the extracellular vesicles may be budding from Serratia cells and are encapsulated with bacterial outer membrane (Matsuyama et al. 1986). Thin-section electron microphotography visualized the presumed budding process of extracellular vesicles full of amorphous material (Fig. 6). Amorphous drops presumably before budding were also observed in bacterial cells. Production of extracellular vesicles has not been recognized with Serratia strains defective in the production of biosurfactants. For examinations of the chemical composition of extracellular vesicles, the vesicles were separated from bacterial cells by differential centrifugation. The isolated vesicle fraction demonstrated strong wetting activity. Then, the chemical compositions of isolated vesicles of pigmented strain NS 38 and nonpigmented strain NS 25 were examined by TLC. The result showed that the main components

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Fig. 6 Thin section electronmicrograph of S. marcescens NS 38 grown at 30
C. The arrow indicates an extending outer membrane from a bacterial cell. Bar, 0.5 mm (from Matsuyama et al. 1986)

of the vesicles were serrawettin W1 or W2 (Matsuyama et al. 1986). In the case of pigmented strain, red pigment prodigiosin was an additional component in the extracellular vesicles. Rubiwettin R1 and RG1 were also secreted from bacterial cells as extracellular vesicles (Matsuyama et al. 1990).

4 Chemical Structure of Serrawettins and Rubiwettins Chemical analyses of purified serrawettins and rubiwettins were performed by standard methods, such as degradation analyses, gas chromatography, infrared spectroscopy, chiral column high-performance liquid chromatography, mass spectrometry, and proton magnetic resonance spectroscopy as described in the original articles (Matsuyama et al. 1985, 1990, 1992; Nakagawa and Matsuyama 1993; Matsuyama and Nakagawa 1996b). Chemical analyses of the biosurfactant from Serratia liquefaciens MG1 were also carried out and serrawettin W2 was identified (Lindum et al. 1998).

4.1

Serrawettin W1, W2, and W3

Degradation analyses demonstrated that serrawettins were composed of fatty acids and amino acids. By infrared spectra analysis, serrawettins were shown to have amide and ester-linkages. Further studies indicated that serrawettin W1 was composed of L-serine and D-3-hydroxydecanoic acid (molar ratio, 1:1). The mass spectrum of serrawettin W1 clearly indicated the molecular ion at m/z 658 and

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Fig. 7 Chemical structures of serrawettins. Serrawettin W1 (a). Serrawettin W2 (b). Space filling model of serrawettin W2 (c)

characteristic patterns of fragment ions. Consequently, mass spectrometric analysis suggested a symmetrical molecule consistent with the results of degradation analyses and infrared spectroscopy. The proposed chemical structure of serrawettin W1 is cyclo(D-3-hydroxydecanoyl-L-seryl)2 as shown in Fig. 7a. These results indicate that serrawettin W1 is identical to serratamolide discovered previously by Wasserman et al. (1961, 1962) as an antibiotic produced by S. marcescens. This symmetrical dilactone molecule has two free hydroxyl groups of serines as hydrophilic parts and has no ionic hydrophilicity. Thus, serrawettin W1 is a nonionic biosurfactant. Serratamolide (serrawettin W1) has been chemically synthesized through a large number of synthetic steps (Shemyakin et al. 1965; Iliev et al. 2006). Recently, 65 mg of pure serratamolide has been synthesized by a novel and robust methodology combining solid-phase reactions with reactions in solution (Teixido´ et al. 2007), for the study of the therapeutic activity (Escobar-Dı´az et al. 2005). Serrawettin W2 was composed of D-3-hydroxydecanoic acid and five different amino acids in equal molar ratios. The amino acid sequence and the location of ester linkage were examined by tandem mass spectroscopy and proton NMR (Matsuyama et al. 1992). Figure 7b shows the proposed chemical structure of serrawettin W2 as a novel cyclodepsipeptide, D-3-hydroxydecanoyl-D-leucyl-Lseryl-L-threonyl-D-phenylalanyl-D-isoleucyl lactone with a molecular weight of 731. Figure 7c is a space-filling model of serrawettin W2 exposing two nonionic hydroxyl moieties (at the opposite position to the hydrophobic heptyl moiety).

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Fig. 8 Chemical structures of rubiwettin R1 (a) and rubiwettin RG1 (b)

Serrawettin W3 was composed of dodecanoic acid, threonine, serine, valine, leucine, and isoleucine (molar ratio, 1:1:2:1:0.5:0.5), and its molecular weight indicated by secondary ion mass spectrometry was 683, which was consistent with the result of the addition and reduction of molecular parts. Thus serrawettin W3 may be mixture of two types of condensation products containing leucine or isoleucine (Matsuyama et al. 1992). Although serrawettin W3 was shown to be a novel nonionic cyclodepsipeptide, structural details have not been elucidated.

4.2

Rubiwettin R1 and RG1

Rubiwettins produced by S. rubidaea seemed to be quite different from serrawettins in their chemical structures, because of the tailing of the R1 spot on a TLC plate developed with a neutral or alkaline solvent system, and positive reactivity of the RG1 spot with anthrone and a-naphthol reagents (Matsuyama et al. 1990). These findings suggested that rubiwettin R1 was an anionic lipid, and RG1 was a glycolipid with Rf value close to that of the rhamnolipid. Further investigations (Matsuyama et al. 1992; Nakagawa and Matsuyama 1993) revealed the chemical structures of rubiwettin R1 and RG1 (Fig. 8). Rubiwettin R1 is a mixture of linked D-3-hydroxy fatty acids belonging to 3-(3-hydroxyalkanoyloxy)alkanoic acids (HAAs). The major components were D-3-(D-30 -hydroxytetradecanoyloxy) decanoate and D-3-(D-30 -hydroxyhexadecanoyloxy) decanoate. In addition, the presence of minor variants such as D-3-(D-30 -hydroxytetradecenoyloxy)decanoate was indicated. On the other hand, rubiwettin RG1 was b-D-glucopyranosyl D-3(D-30 -hydroxytetradcanoyloxy)decanoate and contained minor fatty acid isomers. Specifically, rubiwettin RG1 is rhamnolipid-like glycolipid having glucose moiety instead of rhamnose. Thus, R1 and RG1 were indicated to be novel surface active exolipids. Although rubiwettin R1 (HAAs) will be a precursor of rubiwettin RG1 as in the case of rhamnolipid synthesis (Burger et al. 1963), it is noteworthy that rubiwettin R1 was secreted in massive quantities (Matsuyama et al. 1990) and has a simple chemical structure and potent surface activity. Rubiwettin R1 itself seemed to work independently as an anionic biosurfactant in the physiology of S. rubidaea, as suggested by De´ziel et al. (2003), on rhamnolipid precursor of Pseudomonas aeruginosa. Recently, a Serratia sp. ATCC 39006 strain with the ability to produce

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biosurfactant has been reported to have a gene-sharing homology with the rhlA, the gene involved in the synthesis of HAAs in P. aeruginosa (Williamson et al. 2008).

5 Physiological Functions of Biosurfactants Although serrawettin W1 was first reported as an antibiotic serratamolide, its activity was slight in comparison to the clinically useful antibiotics (Wasserman et al. 1962). Antibacterial activity of serrawettin W2 was also slight (Matsuyama et al. 1992). On the other hand, Staphylococcus aureus coated with serratamolide in vitro showed resistance to phagocytosis by polymorphonuclear leukocytes (Miyazaki et al. 1993). Recently, usefulness of serratamolide for protection of plants from Oomycete pathogens has been shown (Strobel et al. 2003). In terms of the physiological functions as bacterial surfactants, rhamnolipid has been reported as emulsifiers or solubilizers for efficient uptake of water-insoluble nutrients or signal substances (Hisatsuka et al. 1971; Calfee et al. 2005). Although such a function of serrawettins and rubiwettins could be expected, there are no studies with such aspects on serrawettins. Most studies were focused on their functions in dynamic behaviors of bacteria on solid surface environments. In experimental studies with living bacteria for elucidation of the physiological functions of biosurfactants, mutants defective in the production of the corresponding biosurfactants were indispensable. However, routine procedures for finding mutants defective in the ability to produce bacterial lipids are laborious, requiring numerous independent extractions of lipids from mutagenized isolates. In the mutant screening novel method (Matsuyama et al. 1987), a bacterial mass (ca. 1 mg) from a single isolated colony grown on an agar plate, which was streaked with mutagenized bacterial culture, was placed on a Silica Gel G TLC plate and predeveloped in chloroform methanol for 10 min. After 15 min of drying, the shrunken bacterial mass remaining on the plate was removed. Bacterial extracts remaining on the plate were developed and clearly indicated the presence of a specific lipid-deficient mutant. Because the chromatograms obtained by the direct-colony TLC and by the ordinary method using independently prepared lipid extract gave similar spot-development, this method has been used routinely in mutant screenings (Matsuyama et al. 1995; Sunaga et al. 2004).

5.1

Flagellum-Independent Bacterial Spreading Growth

After point inoculation of bacteria onto the surface of hard agar medium, growth behaviors of S. marcescens NS 38 was examined. In contrast to wild-type and a flagellum-less mutant, the mutant defective in the production of serrawettin W1 was unable to exert spreading growth during the 10-day period of culture (Fig. 9). Because these cultivation conditions on a hard agar (1.4 1.5%) medium were not

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Fig. 9 Colonies of S. marcescens NS 38 (1), serrawettin deficient mutant NS 38 09 (2), and flagellum less mutant NS 38 45 (3) on a hard agar (1.5%) medium in a Petri dish (inner diameter, 85 mm) cultivated at 30
C for 10 days

a

b 100000 Df = 1.77 (1 - 87)

N(p)

10000 1000

100

10

1

10

p

100

Fig. 10 (a) A giant colony of S. marcescens NS 38 on a Davis hard agar medium after 20 day culture at 30
C. (b) Fractal analysis of the colony by the box counting method. p, pixel size; N(p), number of boxes; Df, fractal dimension in the box size range shown (from Matsuyama and Matsushita 1993)

permissible to flagellum-dependent swarming of S. marcescens (Harshey 2003), and a serrawettin W1-producing flagellum-less mutant displayed spreading growth similar to the wild-type, it was evident that serrawettin W1 has a critical role in slow but steady spreading growth. This type of function for bacterial spreading behavior was also confirmed for serrawettin W2 and W3 (Matsuyama et al. 1989, 1992). Flagellum-independent slow-spreading growth was mostly observed with bacteria point-inoculated on a hard agar (1.4% Eiken agar) medium with low nutrients (e.g., minimal synthetic Davis medium). After longer incubation periods of more than 2 weeks, a flower-like giant colony developed in a Petri dish. In contrast to small, simple, round colonies usually observed after a few days of incubation, such a characteristic morphology with a complex outline (Fig. 10a) seemed to suggest the

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presence of some special mechanisms. Therefore, geometric analysis of the giant colony morphology was carried out. Figure 10b shows the result of analysis by the box counting method using a computerized image analyzer (Matsuyama and Matsushita 1992, 1993). Briefly, a number N(p) of boxes (with a pixel size, p) covering a colony pattern was determined for each covering. A log log plot of obtained values gave the straight line, indicating that the giant colony morphology was a typical self-similar fractal (fractal dimension, Df ¼ 1.77). Statistical self-similar fractals are ubiquitous in nature (Mandelbrot 1983), for example, bronchial trees in the lung. Although fractal morphology is everywhere in nature, it was quite difficult to find growing fractal objects appropriate for experimental analyses. Discovery of bacterial fractal colonies (Matsuyama et al. 1989) opened the way for in vitro experimental systems for analysis of morphogenesis processes of other familiar patterns in nature (Ben-Jacob et al. 1994; Rauprich et al. 1996; Wakita et al. 1998). Thus, by experimental studies with bacterial fractal colonies of Bacillus subtilis and Salmonella, nutrient-diffusion-limited bacterial multiplication was shown to be a critical factor for the generation of the selfsimilar fractal colony (Matsushita and Fujikawa 1990; Matsuyama and Matsushita 1992), which was consistent with computer simulations based on a diffusion-limited aggregation model (Witten and Sander 1981). It was also noteworthy that the characteristic morphology (Fig. 10a) was not genetically designed. Fundamental processes of bacterial fractal colony morphogenesis are a reflection of basic physical principles (Matsushita 1997). B. subtilis also developed a surfactindependent giant fractal colony. Biosurfactants were considered as enhancers for the fractal colony growth by loosening the water containment force acting on bacteria at the expanding colony front (Matsuyama et al. 1989; Matsuyama and Matsushita 1992). Some bacterial species unable to produce biosurfactants (e.g., Salmonella) were shown to form fractal colonies by simple expansion of the growing cell population under poor nutrients conditions. In such cases, special cellular factors seem to be working (Toguchi et al. 2000; Wang et al. 2005; Chen et al. 2007). S. marcescens or B. subtilis seems to be adapted to enhanced spreading growth by use of their biosurfactants. When S. marcescens was point-inoculated onto the surface of a low-agar (0.35%) medium, the semisolid surface was covered by spreading bacteria in a day. This rapid spreading growth was observed even with a flagellum-less mutant but not with flagellum-less and serrawettin-deficient double mutant (Matsuyama et al. 1995). Thus, serrawettins also contributed to this rapid surface spreading of bacteria on a low-agar medium. P. aeruginosa is well-known as a producer of biosurfactant rhamnolipid. Rhamnolipid also enabled marked spreading growth of P. aeruginosa on a solid (1.5 2.0% agar) medium (Nozawa et al. 2007). Furthermore, a giant colony (Fig. 11a) was formed in a rhamnolipid-dependent process by a flagellum-less or type IV pilus-less mutant (Fig. 11b). However, as shown in Fig. 11a, the smooth outline of a round colony of a parent strain was different from the fractal colony made by S. marcescens. Because a fiord-like outline appeared with a mutant lacking type IV pili, the pili of P. aeruginosa seemed to contribute to the formation of a nonfractal colony.

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a

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70 60 50 40 30 20 10 0 T

TP2

TF2

TR

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Fig. 11 (a) A giant spreading colony of P. aeruginosa PAO1 T on a modified Davis 1.5% agar medium after 5 day culture at 30
C under O2 depleted 8% CO2 condition. An arrowhead indicates the margin of the giant colony. (b) Colony diameters of P. aeruginosa PAO1 T, its mutants, and transformants with plasmids cultured under the same condition as described above. T, wild type; TP2, type IV pilus less mutant; TF2, flagellum less mutant; TR, rhamnolipid deficient mutant; pME, control vector pME6032; pMR2, pME carrying rhlAB. **P < 0.001; n  6 (from Nozawa et al. 2007)

Thus, biosurfactants (serrawettins, surfactin, rhamnolipid) enabled surface spreading growth of bacteria having no motile organs (flagella or pili). It was noteworthy that surfactant-dependent spreading growth occurred in nutrient-poor environments. On the nutrient-rich agar medium, bacteria generally made raised round colonies after long period of culture. On the other hand, the spreading colony on a nutrient-poor medium grew generally as a thin film. Bacteria under the nutrient-poor conditions sought to occupy broader surfaces in order to seek nutrient-rich areas.

5.2

Flagellum-Dependent Bacterial Spreading Growth

Bacterial swarming has been known as flagellum-dependent surface-restricted migration of bacteria on a solid or semisolid medium. Swarming of pigmented S. marcescens NS 38 wild-type and NS 38-09 (serrawettin W1-deficient mutant) strains on a semisolid agar medium is shown in Fig. 12. Although both strains formed swarming colonies, characteristic differences in their spreading patterns developed (Matsuyama and Matsushita 1993). Dense but thin branching morphology was evident with the swarming colony of S. marcescnes wild-type. On the other hand, rarely branching rough morphology was observed with the mutant. Serrawettin W1 seemed to promote thin branching of the colony by weakening the surface tension (containment force) at the margin of the colony. Weakened containment force seemed to permit many thin protrusions by swarming bacterial groups (recognized using a microscope). Devoid of such containment-force weakening, the serrawettin W1-deficient mutant was unable to develop dense branching

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Fig. 12 Swarming colonies of S. marcescens NS 38 (a) and serrawettin deficient mutant NS 38 09 (b). Bacteria were point inoculated onto the center of semisolid nutrient agar (0.5%) and cultured at 30
C for 12 h (from Matsuyama and Nakagawa 1996a)

Fig. 13 (a) Extracellular complementation by serrawettin producing live cells. Serrawettin W1 producer S. marcescens NS 38 45 (flagellum less mutant) was point inoculated at the center of 0.5% agar nutrient medium and incubated at 30
C for 12 h. Three different S. marcescens mutants only defective in the production of serrawettins (NS 38 09, NS 25 09, and NS 45 09) were point inoculated nearby and away from the center. Compare inside colonies (near the central colony) to outside colonies. The inside colonies are modified and extending thin branches. (b) Effect of purified serrawettin W2 on bacterial swarming growth on a semisolid medium. B. subtilis JH 642 (nonproducer of surfactin) were point inoculated at sites away from (right of the plate) and nearby (center) the paper disk (left) containing 100 mg of serrawettin W2. Culture was at 37
C for 24 h (from Matsuyama and Matsushita 1996)

and formed thick branches with a remarkable tendency to bend to the right (Matsuyama and Matsushita 1996). As shown in Fig. 13a, the serrawettin W1producing S. marcescens NS 38-45 (flagellum-less mutant) strain point-inoculated near to W1-deficient, W2-deficient, and W3-deficient S. marcescens mutants performed extracellular complementation and enabled the characteristic dense branching by these mutants (Matsuyama and Matsushita 1996). Purified serrawettins soaked into paper disks also induced the dense branching of swarming colonies

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of serrawettin-deficient S. marcescens mutants and biosurfactant-deficient other species of bacteria (Matsuyama et al. 1993). The effect of purified serrawettin W2 on to the swarming of B. subtilis JH 624 (nonproducer of surfactin) is shown in Fig. 13b. Swarming of P. aeruginosa on a semisolid (0.5% agar) medium was also reported to be rhamnolipid (and its precursor)-dependent (K€ohler et al. 2000; De´ziel et al. 2003; Heurlier et al. 2004). Recently, rhamnolipid-dependent swarming of P. aeruginosa on a hard agar (1.5%) medium has been reported (Takahashi et al. 2008). With regards to the P. aeruginosa swarming on the hard agar medium, it was noteworthy that bacterial cells swarmed without differentiation into elongated hyperflagellates. In the well-known swarming of Proteus mirabilis (Williams and Schwarzhoff 1978) or Vibrio parahaemolyticus (McCarter 1999) on a hard agar (1.5%) medium, cellular differentiation into elongated hyperflagellates was an essential requirement for coordinated efficient cellular translocations (Allison and Hughes 1991). Even in the swarming of S. marcescens, E. coli, and Salmonella typhimurium on softer agar (0.75%) medium, these bacterial species exerted similar cellular differentiation (Alberti and Harshey 1990; Harshey and Matsuyama 1994). However, P. aeruginosa swarming cells were shown to maintain their unipolar and normal sized rod shape with one or two flagella (Takahashi et al. 2008). It is also noteworthy that this rhamnolipid-dependent swarming by P. aeruginosa occurred on nutrient-poor medium and not on a nutrient-rich medium (Takahashi et al. 2008). The giant colony that developed after 2 days culture was composed of a thin film-like sheet broadly occupying the agar medium surface. Such bacterial behavior was quite similar to that of flagellum-independent but rhamnolipiddependent slow-spreading growth of P. aeruginosa on a hard agar (Fig. 11a).

5.3

Other Biological Activities of Serrawettins

The nematode Caenorhabditis elegans living in soil is a bacterivorous organism. When C. elegans worms were placed on a lawn of E. coli grown in a Petri dish, the worms remain on the lawn until all bacteria were eaten. However, when the worms were put in a lawn of bacteria (S. marcescens) pathogenic to C. elegans, the worms migrated out the lawn after a few hours. Thus, C. elegans seemed to have the ability to avoid pathogenic bacterial populations by sensing specific bacterial products. S. marcescens mutant JESM267, which was unable to induce an avoidance reaction by C. elegans, was found to be serrawettin W2-deficient mutant. Therefore, purified serrawettin W1, W2, and W3 were examined for C. elegans repellent activity by spotting on the E. coli lawn. Because C. elegans showed avoidance reaction to these spotted E. coli lawns, but not from a lawn spotted with surfactin, serrawettins (not the virulence factor of S. marcescens in infection to C. elegans) were considered to be specific repellent sensed by C. elegans (Pradel et al. 2007). This case seems to

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indicate an unexpected biological role of serrawettins functioning as a chemical signal in predator prey relationships in addition to the physicochemical activity of the biosurfactant. With regards to antitumor activity, serratamolide (AT514) has been reported to be a potent inducer of apoptosis of several cell lines derived from various human tumors and B-chronic lymphocytic leukemia cells (Escobar-Dı´az et al. 2005). Consequently, biological studies of AT514 using human B-lymphocytes are now in progress for clinical applications of AT514 in the field of medical oncology.

6 Genetics of Biosynthesis and Regulation Serrawettins are secondary metabolites with cyclic aminolipid structures. Production of such molecules seems to require systematic and energy-consuming processes for linking specific components in a defined sequence. In addition, generous production of serrawettins on surfaces or at 30
C and scarce production in liquid or at 37
C suggested distinct adaptability of these organisms to variable environments. How do these processes occur and how are they regulated? The responsible molecular mechanisms were partly brought to light through analyses of serrawettin-deficient or -overproducing mutants and swarming defective mutants.

6.1

Genes Involved in Serrawettin W1 Biosynthesis

Among serrawettin W1-deficient mutants due to a single transposon insertion, four mutants of S. marcescens 274 were also defective in the production of prodigiosin. Although serrawettin W1 and prodigiosin were secreted together as exolipids in vesicles and produced in parallel in the same temperature-dependent manner, they have no common molecular constituents in their structures. In particular, prodigiosin is only composed of three pyrrole rings linked in a linear manner. Initially, this finding was mysterious. However, sequencing of the DNA fragment carrying the inserted transposon from such serrawettin- and prodigiosin-deficient mutants indicated the presence of an essential step in the biosynthesis of serrawettin W1. Briefly, the genetic study revealed an open reading flame (ORF) of a novel gene pswP (DDBJ Accession Number AB163428) encoding 40 -phosphopantetheinyl transferase (PPTase), which is the activator of peptidyl carrier protein (PCP). Thus, PPTase activity by PswP in S. marcescens was shown to be indispensable for the biosynthesis of serrawettin W1, presumably in the incorporation reaction of L-serine as a molecular component of the whole molecule (Sunaga et al. 2004). With regards to the biosynthesis of prodigiosin, one pyrrole ring in a prodigiosin precursor molecule has been reported to be an oxidized product of a pyrrolidine ring of proline tethering to PCP through a 40 -phosphopantetheinyl moiety (Thomas et al.

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2002). Specifically, PswP seemed to be required for the initial transfer of the 40 -phosphopantetheinyl moiety from coenzyme A to apo-PCP to make activated PCP (holo-PCP). Consequently, mutation of the gene pswP will result in the generation of a prodigiosin-deficient mutant and mutants defective in the production of other secondary metabolites composed of amino acids or amino acid-derivatives. All four mutants with a combined failure of the production of serrawettin W1 and prodigiosin completely restored their defects by transformation with a low copy-number plasmid carrying the single pswP gene (Sunaga et al. 2004). Consistent with experimental findings noted above, recent studies on the biosynthesis of secondary metabolites (e.g., antibiotics) have disclosed the systematic functioning of nonribosomal peptide synthetases (NRPSs) (Trauger et al. 2000) in which specific components (e.g., amino acids) that are components of final products are tethered to the phosphopantetheinyl moiety of each thiolation domain in the multimodular synthetic enzyme (Cosmina et al. 1993). It was possible that serrawettin W1 synthetase may also belong to the NRPS family, as indicated with a partially sequenced swrA gene involved in serrawettin W2 biosynthesis (Lindum et al. 1998). In addition, in spite of intensive searches for accumulating precursors of serrawettin W1 in the serrawettin W1-deficient mutants, it was not possible to find serratamic acid (D-3-hydroxydecanoyl-L-serine; the most probable precursor of the dilactone serrawettin W1) in lipid extracts of these mutants. Alkaline extracts of Serratia group wild-type were reported to contain the serratamic acid (Cartwright 1955), but they should now be considered as hydrolytic products of serrawettin W1. Remaining seven serrawettin W1-deficient mutants were not defective in the production of prodigiosin. Genes other than the gene pswP were shown to be inactivated in these mutants by a single transposon insertion. Sequencing of the DNA flanking mini-Mu transposon indicated the presence of a remarkably large ORF named swrW (DDBJ Accession No. AB193098). The putative 1,310 aminoacid sequence deduced from the ORF (from base No. 481 to 4,413 in Fig. 14a) showed high homology with the NRPS family (Li et al. 2005). The presumed product SwrW was composed of condensation, adenylation, thiolation, and thioesterase domains in functional order, as observed in the NRPS family (Marahiel et al. 1997). It seemed evident that one of these domains, the thiolation domain, must be activated by acquiring the phosphopantetheinyl moiety by the action of PswP (PPTase) for the proper functioning of serrawettin W1 synthetase SwrW. The presumed biosynthetic processes by SwrW and acyl carrier protein (ACP) after the activation by PPTase (e.g., PswP) are indicated in Fig. 14b. In the first step, the adenylation domain adenylates L-serine, then the activated L-serine will bind as thioester to the thiolation domain, which has been phosphopantetheinylated beforehand by PswP. To the amino group of the tethered L-serine on the thiolation domain, the 3-D-hydroxydecanoyl moiety tethered to ACP will react to create an amide linkage by detaching from ACP. Thus, serratamic acid tethered to the thiolation domain will be formed. A mutant lacking ACP has not been isolated from S. marcescens, which is a situation similar to the study of surfactin biosynthesis (Cosmina et al. 1993). It is possible that ACP is required for cellular fatty acid

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Fig. 14 (a) Schematic representation of the swrW gene. Sections corresponding to condensation (C), adenylation (A), thiolation (T), and thioesterase (TE) domains of a putative NRPS are bordered by thin vertical lines. Conservative motifs (e.g., C1 C7) in the four domains are indicated with thick vertical bars. (b) NRPS dependent biosynthesis of serrawettin W1. Presumed steps for the synthesis of serrawettin W1 are indicated by sequential numbers from (1) to (10). The zigzag line extending from the ACP (acyl carrier protein) or T domain represents part of a phosphopan tetheinyl moiety (from Li et al. 2005)

synthesis, so a mutation of such a fundamental gene will be lethal. The thiolation domain is freed by transferring the first serratamic acid to the neighboring thioesterase domain. The next step for tethering the second serratamic acid to the free phosphopantetheinylated thiolation domain will follow. Thus, two serratamic acids

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will be tethered to the thiolation and thioesterase domains side by side. Thereafter, processes (No. 8 10 in Fig. 14b) similar to the ones proposed for the biosynthesis of gramicidin S (Trauger et al. 2000) will occur, and the final product serrawettin W1 will be released from SwrW. In contrast to multimodular NRPSs such as SyrE (syringomycin synthetase has eight modules), as reported by Guenzi et al. (1998), SwrW is the simplest unimodular NRPS and producer of rotationally symmetric aminolipid with no peptide bonds. Complementation with DNA carrying the swrW gene completely restored the ability to produce serrawettin W1 in all of the seven serrawettin-deficient mutants of S. marcescens 274. In the construction process of plasmid carrying the swrW gene for the marker rescue, we used E. coli JM 109 as a plasmid host strain. However, the E. coli strain harboring this swrW gene carrying plasmid did not produce serrawettin W1. The native PPTase (EntD) of E. coli seemed to fail to phosphopantetheinylate the thiolation domain of SwrW, presumably reflecting its low homology (37% identity) with PswP of S. marcescens. Therefore, E. coli JM 109 transformed with the swrW and pswP genes was prepared and examined for serrawettin W1 production. In the examination of wetting activity of bacterial mass, the transformed E. coli clearly exhibited wetting activity. In TLC examination, the production of serrawettin W1 by E. coli was confirmed. Thus, for the production of serrawettin W1 by the synthetase SwrW in heterologous hosts, specificity of PPTase should be considered (Tanikawa et al. unpublished data). In addition, it was noteworthy that serrawettin W1 production in E. coli occurred at 30
C but not at 37
C. Thus, the temperature-dependent production mode of the serrawettin was also reproduced in transformed E. coli (Tanikawa et al. unpublished data). This was an unexpected result, because no thermoregulator genes of S. marcescens were considered to be transferred to E. coli. To examine this finding, another E. coli transformant with a cluster of S. marcescens genes (pigA etc) for prodigiosin biosynthesis (Harris et al. 2004) and the pswP gene was prepared and examined for prodigiosin production. The results indicated that temperaturedependent production of prodigiosin was evident in the E. coli transformant. Is the pswP gene responsible for thermoregulation in the production of serrawettin W1 and prodigiosin? This possibility seemed to be unlikely, because pswP-defective S. marcescens mutants receiving a high copy-number plasmid carrying the pswP gene exhibited no change in its temperature-dependent production of serrawettin W1 and prodigiosin (Sunaga et al. 2004).

6.2

Quorum-Sensing Regulation of Serrawettin Biosynthesis

Genes involved in serrawettin W2 biosynthesis were elucidated through studies on the multicellular behavior of S. liquefaciens MG1 (Eberl et al. 1996; Lindum et al. 1998). Swarming motility on surfaces has been shown as a representative multicellular behavior of bacteria (Allison and Hughes 1991). S. liquefaciens wild-type MG1 is capable of swarming motility and has cell-density-dependent signaling mechanisms known as “quorum sensing.” In particular, following the positive

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bioluminescent test with sensor plasmid pSB403, the production of signal molecules named autoinducers, such as N-acyl homoserine lactones (AHLs), was confirmed by defining N-butanoyl-L-homoserine lactone [BHL] and N-hexanoylL-homoserine lactone [HHL] in strain MG1. Then, a S. liquefaciens gene (swrI) involved in AHLs synthesis was identified and the presumed protein product SwrI showed strong homology to members of the LuxI family (Eberl et al. 1996). Along with this work, an ORF (located downstream of swrI) encoding a polypeptide (SwrR) with similarity to members of the LuxR family was also identified (Givskov et al. 1997). Mutant MG44 defective in the production of AHL was selected by insertion of a streptomycin-resistance marker into the swrI gene. It was noteworthy that the swarming behavior was defective in mutant MG44, and this defect was restored by supplementation of synthetic BHL to the swarming medium. This finding became the first example of autoinducer controlled-swarming behavior of bacteria (Eberl et al. 1996). In the subsequent examinations, the swrI mutant MG44 was shown to be negative in the drop-collapsing test in contrast to surfactantproducing wild-type, and this negative result was restored by supplementation of BHL. Because BHL has no surfactant activity, this result seemed to indicate that the quorum-sensing mechanism of S. liquefaciens MG1 controls the production of biosurfactant. Thereafter, as a target gene of the quorum-sensing system (i.e., BHL-dependent genes), a novel gene swrA (GenBank accession no. AF039572, partial ORF coding for more than 991 amino acids), encoding a putative peptide synthetase with high homology to the surfactin synthetase SfrA of B. subtilis (Cosmina et al. 1993) was identified (Lindum et al. 1998). Isolation of the biosurfactant produced by S. liquefaciens MG1 was also performed, and this was identified as serrawettin W2 by Lindum et al. (1998). Thus, serrawettin W2-dependent swarming of S. liquefaciens MG1 (recently reclassified as a S. marcescens strain) (Rice et al. 2005) was shown to be regulated by the quorum-sensing system. With regards to the serrawettin W1-producing strain 274, the absence of AHLs-dependent quorum-sensing system has been reported (Coulthurst et al. 2004, 2006; Tanikawa et al. 2006). The smaI/smaR (sharing high sequence similarity with swrI/swrR ) locus from S. marcescens 12I (smaI disrupted strain) was introduced into serrawettin W1-producing strain 274 by general transduction. Swarming motility of the resultant transductant was impaired, but this behavior was restored by BHL supplementation of the medium (Coulthurst et al. 2006). This result seemed to show that serrawettin W1-dependent swarming of strain 274 was also sensitive to the quorum-sensing regulation.

6.3

Global Regulator Genes Concerned with Exolipid Production

A mutant defective in the regulation of serrawettin W1 and prodigiosin production was isolated from mini-Tn5 inserted S. marcescens 274. Although the mutant Tan1 developed pigmented colonies after cultivation at 37
C, it seemed to overproduce prodigiosin and serrawettin W1 at 30 and 37
C. Therefore, instead of the disruption

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of thermoregulation, disruption of parallel downregulation in the production of two exolipids was considered as a possible defect. Comparative time-lapse TLC analysis of exolipids demonstrated earlier and greater production levels of prodigiosin and serrawettin W1 in the Tan1 mutant (Fig. 15). Bacterial biosurfactants, in contrast to synthetic surfactants, have many advantages for technological applications in human life. However, improvement of the low level production in industrial cultivation would be an important requirement (Das et al. 2008). Enhanced production of serrawettin W1 in the mutant Tan1 seemed to be a valuable example for further analyses. In the sequencing of a putative 945 ORF responsive for the defect, the gene named hexS (DDBJ Accession No. AB164082) was identified (Tanikawa et al. 2006). Site-directed mutagenesis of wild-type S. marcescens 274 by insertion of a Cmr gene produced several mutants with the same phenotype as Tan1. The aminoacid sequence deduced from the ORF indicated high homology with a LysR-type transcriptional regulator HexA of Erwinia carotovora ssp. carotovora (73% identity and 81% similarity) and other LysR-type regulators. LysR-type proteins have been reported as transcriptional regulators. Therefore, the mRNA transcription level was compared between 274 and Tan1 by RT-PCR analysis of three genes pigA, swrW, and pswP. The results clearly showed overtranscription of pigA and swrW in Tan1 (Tanikawa et al. 2006). In contrast, the transcription level of pswP in Tan1 was similar to that in 274. Thus, a target-specific repressive function of HexS in mRNA transcription was suggested, and there was no involvement of PswP in this parallel downregulation. Then, a large quantity of His6-tagged HexS protein was prepared in E. coli, and its molecular size was shown

b Time (hr)

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274 11 Tan1

log CFU/ml

10 9

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Tan1

Fig. 15 Uncontrolled overproduction of prodigiosin and serrawettin W1 by a hexS mutant (Tan1) of S. marcescens cultured in LB broth at 30
C. (a) Growth curves of S. marcescens parent strain 274 (open circle) and Tan1 (open triangle). Levels of prodigiosin (b) and serrawettin W1 (c) productions by 274 and Tan1 examined by TLC at intervals. Prodigiosin and serrawettin W1 spots appearing zones in specific parts of the developed TLC plates are shown. TLC plates (a) were not heated (from Tanikawa et al. 2006)

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to be 34.8 kDa. Thereafter, differential direct binding of the purified His6-tagged HexS protein to the promoter region of the downregulated genes (pigA and swrW) was confirmed by gel mobility shift analyses using the promoter region of the pswP gene and the coding regions of these three genes as negative controls (Tanikawa et al. 2006). In the hexS gene DNA transformants of several S. marcescens wild-types, overdownregulation of specific exolipid (prodigiosin, serrawettin W1, W2, and W3) productions was recognized. In addition, several exoenzymes (protease, chitinase, and DNase) except phospholipase C showed specifically reduced activity in the hexS gene DNA transformant of S. marcescens 274. Thus, unlike most LysR-type regulators, which work as transcriptional activators, HexS was a transcriptional negative regulator. The presence of such a gene means that Serratia is ready for situations requiring more serrawettins, prodigiosin, and exoenzymes. For what sort of situation is the organism preparing? Although knockout of this downregulator gene may generate useful mutant strains for microbial industries, elucidation of specific conditions resulting in the switch-off of the hexS gene may also lead to deeper insights into the bacterial life.

7 Concluding Remarks In the studies on serrawettins, we realized that microbes are strongly influenced by the surface tension of water. Water is indispensable for bacteria and most bacterial cells, even those multiplying on a solid agar surface, are wet, i.e., covered with a thin water film. Bacteria may be unable to escape from the surface of water by their own power except as spores released into the air. Consequently, bacteria that move along a surface must carry a water film with them. However, water will resist broadening its surface area and bacteria will be limited to water-covered areas. To overcome such difficulty in migration to new surface environments and to break out of the dilemma of maintaining a water-dependent life, bacteria produce surface active agents. Biosurfactants may loosen containment of expanding bacterial population and enable the spreading growth of bacteria with water film. Synthetic surfactants (e.g., Tween 80) also exerted the same effect for bacterial spreading growth (Nozawa et al. 2007; Takahashi et al. 2008). The precise mechanisms of such bacterial spreading growth, however, have not yet been clarified. For example, by dropping a water droplet on a surface of solid medium, we recognized that the high water wettability of a solid medium surface (e.g., medium with Difco rather than Eiken agar) before the inoculation of bacteria was inhibitory to surfactant-dependent spreading growth (Matsuyama et al. 1989; Matsuyama and Matsushita 1996). It was possible to make the medium surface more wettable by preparing Tween 80-containing medium. However, on such medium, P. aeruginosa failed in rhamnolipid-dependent spreading growth. In spreading growth, bacteria need to secrete biosurfactant with wetting activity, but spreading growth of S. marcescens or P. aeruginosa was generally observed on

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the surface with low wettability, (Takahashi et al. 2008; Nozawa et al. unpublished results). So far, there are no plausible explanations for these contradictory findings. Spreading growth with biosurfactants seems to be passive translocation depending on the expansion pressure caused by the population-volume increase resulting from bacterial multiplication, and the easy rupture and sealing of weakened water surfaces with surfactant molecules. Such passive translocation seems to work with low energy cost. Thus, it seems natural that surfactant-dependent thin spreading growth occurs on the surface of a nutrient-poor medium. Bacteria in low-nutrient environment may have to grow across a considerable distance in search of nutrientrich places. Hunger may be the strongest motivation for the migration of every organism. In this situation, surfactant-dependent translocation may be reliable in contrast to energy-consuming flagellum- or pilus-dependent spreading growth. Acknowledgments The authors are indebted to all their coworkers at Niigata University and Chuo University. This work was supported in part by Grants in Aid for Scientific Research from the Ministry of Education, Science and Culture of Japan, and by a grant from the Urakami Foundation.

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Rice SA, Koh KS, Queck SY, Labbate M, Lam KW, Kjelleberg S (2005) Biofilm formation and sloughing in Serratia marcescens are controlled by quorum sensing and nutrient cues. J Bacteriol 187:3477 3485 Shemyakin MM, Antonov VK, Shkrob AM, Shchelokov VI, Agadzhanyan ZE (1965) Activation of the amide group by acylation; hydroxy and aminoacyl incorporation in peptide systems. Tetrahedron 21:3537 3572 Strobel GA, Morrison SI, Cassella M (2003) Methods for protection of plants from Oomyocyte pathogens by use of Serratia marcescens and isolates. US Patent Appl US2003/0049230 A1 Sunaga S, Li H, Sato Y, Nakagawa Y, Matsuyama T (2004) Identification and characterization of the pswP gene required for the parallel production of prodigiosin and serrawettin W1 in Serratia marcescens. Microbiol Immunol 48:723 728 Takahashi C, Nozawa T, Tanikawa T, Nakagawa Y, Wakita J, Matsushita M, Matsuyama T (2008) Swarming of Pseudomonas aeruginosa PAO1 without differentiation into elongated hyper flagellates on hard agar minimal medium. FEMS Microbiol Lett 280:169 175 Tanikawa T, Nakagawa Y, Matsuyama T (2006) Transcriptional downregulator HexS controlling prodigiosin and serrawettin W1 biosynthesis in Serratia marcescens. Microbiol Immunol 50:587 596 Teixido´ M, Caba JM, Prades R, Zurita E, Martinell M, Vilaseca M, Albericio F, Giralt E (2007) Does the solid phase synthesis of a tetrapeptide represent a challenge at the onset of the XXI century? The case of cyclo [(3R) 3 hydroxydecanoyl L seryl (3R) hydroxydecanoyl L seryl]. Int J Pept Res Ther 13:313 327 Thomas MG, Burkart MD, Walsh CT (2002) Conversion of L proline to pyrrolyl 2 carboxyl S PCP during undecylprodigiosin and pyoluteorin biosynthesis. Chem Biol 9:171 184 Toguchi A, Siano M, Burkart M, Harshey RM (2000) Genetics of swarming motility in Salmonella enterica serovar Typhimurium: critical role for lipopolysaccharide. J Bacteriol 182:6308 6321 Trauger JW, Kohli RM, Mootz HD, Marahiel MA, Walsh CT (2000) Peptide cyclization catalyzed by the thioesterase domain of tyrocidine synthetase. Nature 407:215 218 Wakita J, Ra`fols I, Itoh H, Matsuyama T, Matsushita M (1998) Experimental investigation on the formation of dense branching morphology like colonies in bacteria. J Phys Soc Jpn 67: 3630 3636 Wang Q, Suzuki A, Mariconda S, Porwollik S, Harshey RM (2005) Sensing wetness; a new role for the bacterial flagellum. EMBO J 24:2034 2042 Wasserman HH, Keggi JJ, Mckeon JE (1961) Serratamolide, a metabolic product of Serratia. J Am Chem Soc 83:4107 4108 Wasserman HH, Keggi JJ, Mckeon JE (1962) The structure of serratamolide. J Am Chem Soc 84:2978 2982 Williams FD, Schwarzhoff RH (1978) Nature of the swarming phenomenon in Proteus. Annu Rev Microbiol 32:101 122 Williamson NR, Fineran PC, Ogawa W, Woodley LR, Salmond GPC (2008) Integrated regulation involving quorum sensing, a two component system, a GGDEF/EAL domain protein and a post transcriptional regulator controls swarming and RhlA dependent surfactant biosynthesis in Serratia. Env Microbiol 10(5):1202 1217. doi: 10.1111/j. 1462 2920. 2007. 01536.x Witten TA, Sander LM (1981) Diffusion limited aggregation, a kinetic critical phenomenon. Phys Rev Lett 47:1400 1403 Yamashita M, Nakagawa Y, Li H, Matsuyama T (2001) Silica gel dependent production of prodigiosin and serrawettins by Serratia marcescens in a liquid culture. Microbes Environ 16:250 254

Trehalolipids Zongze Shao

Contents 1 2 3

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 Trehalolipid Producing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 Chemical Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 126 3.1 Cord Factor: A Trehalose Diester from Mycobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127 3.2 Trehalose Lipids from Rhodococcus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129 4 Physicochemical Property . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 5 Biological Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 5.1 Trehalose Lipids in Mycobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136 5.2 Trehalose Lipids from Rhodococcus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136 6 Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 7 Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 7.1 Substrates Used for Trehaloselipids Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138 7.2 Cell Wall Association . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 7.3 Growth Associated Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 140

Abstract Trehalose-containing glycolipids are mainly produced by Gram-positive, high GC content bacteria of Actinomycetales. Their structures are quite diverse in hydrophobic moiety, varying from short simple to long complex fatty acids. Correspondingly, functions and physiochemical properties vary upon structures. From the view of practical use as a biosurfactant, the trehalose lipids from Rhodococcus and the genera other than Mycobacterium are of high potential in application. While, like other kinds of biosurfactants, their relative low productivity limits practical use. And yet, the biosynthesis mechanism of trehalose lipids has been less exploited and needs further investigations. Z. Shao Key Laboratory of Marine Biogenetic Resources, Third Institute of Oceanography, State of Oceanic Administration, Daxue Lu 178, 361005 Xiamen, China e mail: [email protected]

G. Sobero´n‐Cha´vez (ed.), Biosurfactants, Microbiology Monographs 20, DOI 10.1007/978 3 642 14490 5 5, # Springer Verlag Berlin Heidelberg 2011

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Z. Shao

1 Introduction Trehalolipids, or trehalose lipids, are glycolipids containing trehalose hydrophilic moiety. They are among the best known biosurfactants but distinguished from rhamnolipids and sophorolipids in both composition and activity. They are active to lower water surface and interface tension. In addition, they function uniquely in many other ways. The occurrence of trehalose in lipids was first noticed in 1933 in fat extracts from tubercle bacilli (Anderson and Newman 1933). At that time, the authors did not pay attention to the property as a biosurfactant but found intriguing that trehalose was in place of glycerol as the alcoholic component of a fat. However, the pure compound of trehalose lipid was first successfully purified in 1956 from the lipids of Mycobacterium tuberculosis and identified as 6, 60 -dimycoloyl-a0 , a0 -D-trehalose (cord factor). Later, several groups of trehalose-containing lipids were isolated, mainly from Mycobacteria, Nocardia, and Corynebacteria, as reviewed by Asselineau and Asselineau (Asselineau and Asselineau 1978). Interest in trehalose lipids as a general surfactant can be traced back to the discovery that the emulsion layer of Arthrobacter paraffineus culture broths contained trehalose dimycolates when the cells were grown on hydrocarbon substrates (Suzuki et al. 1969). Various trehalose lipids varying in fatty acid part have been found. The glycolipids from mycobacteria usually have branched long-chain hydroxyaliphatic acids as their hydrophobic moiety; while some bacteria choose short-chain fatty acids, such as some cases of Rhodococcus. Trehalose lipids are quite diverse in chemical structure and attractive in applications as other biosurfactants.

2 Trehalolipid-Producing Bacteria Production of trehalolipids mainly occurs in the following bacteria of Actinomycetales, including the genera of Rhodococcus, Mycobacterium, Micrococcus, Nocardia, Gordonia, Corynebacterium, Brevibacteria, Arthrobacter, etc. (Table 1). However, yeast and other fungus have seldom been reported as a producer of trehalose lipids. Among the reported trehalolipid-producing bacteria, most are alkane-degrading bacteria. Surfactive compounds are generated only when bacteria feed on hydrophobic substrates such as n-hexadecane (listed in Table 1), such as Rhodococcus erythropolis, Rhodococcus opacus, Rhodococcus wratislaviensis, Gordonia amarae, Arthrobacter parafineus, Micrococcus luteus, Mycobacterium paraffinicum, Corynebacteria, etc. They have different isolation origins, some of which are from marine environments (Passeri et al. 1991; Schulz et al. 1991; Yakimov et al. 1999; Satpute et al. 2010); some are from alkaline soils or polar soils. These bacteria seem to be tolerant with high salt concentrations (up to 6% salinity).

Rhodococcus fascians

Rhodococcus isolate Q

Trehalose-containing lipids (not fully characterized)

Arthrobacter sp.4301; Brevibacteria sp.; Corynebacterium spp.; Nocardia spp. Arthrobacter sp. EK 1

An anionic 2,3,4,20 -trehalose tetraester; containing succinate A novel trisaccharide glycolipid; containing trehalose bears ester-linked hexanoate, succinate, and acyloxyacyl moieties

–

a-branched-b-hydroxy fatty acid trehalose ester

Arthrobacter parafineus KY4303

32

–

–

–

–

Trehaloselipid-o-dialkyl monoglycerides

–

Pseudomonas fluorescence

Corynebacterium matruchotii

Cord-factor (6,60 -diester of trehalose containing two moles of smegma mycolic acids) Trehalose 6-mono- and 6,60 di-corynomycolates –

–

2,3-di-O-acyltrehalose

Mycobacterium tuberculosis H37Rv Mycobacterium smegmatis

–

–

–

–

–

–

–

–

–

–

Surface tension Interfacial tension (mN m1) (mN m1)a –

Trehalose lipids

Mycobacterium fortuitum Mycoside F

Bacteria

Table 1 Trehalose lipids: producing bacteria and physiochemical properties

–

–

–

–

–

–

–

–

–

–

CMC (mg l1)

n-alkanes

n-hexadecane; definitely not glucose

Crude oil degrading

n-paraffin

n-paraffin

Alkanes

–

–

–

–

Substrates

–

–

–

–

Growth association

Significant amounts of cell-associated

2 (crude extract)

–

Perhaps

–

High yield with gasoline; – depending upon hydrocarbon source 1.3. Penicillin reduced Yes the yield in emulsion layer; but stimulated extracellular accumulation 0.5–1.9 Yes

–

–

–

–

Production

(continued)

Passeri et al. (1991), Schulz et al. (1991) Esch et al. (1999)

Suzuki et al. (1969)

Suzuki et al. (1969)

Datta and Takayama (1993) Desai et al. (1988)

(Mompon et al. 1978)

Gautier et al. (1992) Besra et al. (1992)

References

Trehalolipids 123

2,3,4,20 -trehalosetetraesters containing succinic acid

Two main succinoyl trehalose 19 (STL-1) lipids (STL-1 and STL-2); MW STL-1 ¼ 1,019

Rhodococcus erythropolis SD-74

Rhodococcus wratislaviensis BN38

A mixture of glucolipids and trehalolipids

Rhodococcus erythropolis 3C-9

28.6 (mixture) 24.4 (purified)

33.4 (culture)

–

Novel trehalose dinocardiomycolates of unsaturated fatty acid chains

Rhodococcus opacus 1CP

5.3

–

–

–

5

5 (STL-1)

–

–

0.02 (against 1.5 decane)

15

<1

26

–

4

0.7

14

18

CMC (mg l1)

32

43

Surface tension Interfacial tension (mN m1) (mN m1)a

A mixture with octaacyltrehalose as main component

Trehalose monocorynomycolates Trehalose-2,20 ,3,4-tetraester

a mixture of trehalose lipids Trehalose dicorynomycolates

Trehalose lipids

Rhodococcus sp. H13-A

Rhodococcus erythropolis DSM43215

Bacteria

Table 1 (continued)

C8–C17, best with nhexadecane

n-alkanes; definitely not glucose n-hexadecane

n-alkane

n-alkanes (best with C10) n-alkanes

n-alkanes

Substrates

3.1 g l1

Extracellular accumulation

From 247 mg l1 for n-decane up to 420 mg l1 for n-dodecane Growth on C14 and C16 gave yields between 333 and 315 mg l1 –

Extracellularly accumulated

32

Both extracellular and cell-bound –

Production

Yakimov et al. (1999) Kretschmer et al. (1982), Kretschmer and Wagner (1983), Kim et al. (1990)

References

Yes

–

Yes

Yes

No

Uchida et al. (1989), Tokumoto et al. (2009) Tuleva et al. (2008)

Peng et al. (2007)

Singer and Finnerty (1990), Singer et al. (1990) Niescher et al. (2006)

No. nitrogen Kim et al. limited (1990)

Yes

Growth association

124 Z. Shao

1.7

–

–

Trehalose esters of the C8–C11 fatty acids (depending on the carbon source) Two main products of trehalose tetraesters, MW ¼ 876, 848 24.1

5

27.9

A trehalose tetraester

Water against hexadecane

a

Micrococcus luteus BN56

Rhodococcus erythropolis 51T7 Corynebacteria sp.51 T7 ¼ R erythropolis 51T7 25

–

37

n-hexadecane

Tetradecane (2%,V/V) n-alkanes In first 80 h, mainly associated with the cell wall; afterwards, mainly extracellular –

0.48–1.12

Yes

–

Yes

Tuleva et al. (2009)

Marques et al. (2009) Martin et al. (1991)

Trehalolipids 125

126

Z. Shao

Among them, isolates of Rhodococcus are the most potential in biosurfactant production (Lang and Philp 1998). Rhodococcus bacteria usually produce trehalose lipids when they grow with alkanes as carbon sources (Espuny et al. 1995; Rapp et al. 1979; Singer et al. 1990). Among the bacteria of this genus, isolates of R. erythropolis are most frequently reported as a producer of trehalose lipids (Lang and Philp 1998; Peng et al. 2007). On the other hand, glycolipids other than trehalose lipids have been also found recently in bacteria of Rhodococcus, e.g., a glucolipid was found in R. erythropolis isolate 3C-9 in addition to a trehalolipid (Peng et al. 2007); for another example, rhamnose glycolipid was found as the main biosurfactant in the case of Rhodococcus fascians isolate A-3 grown with glucose or kerosene as a substrate (Gesheva et al. 2010). R. erythropolis 51 T7 is a relatively intensively investigated bacterium. It was once named Corynebacterium sp. 51 T7 at the beginning (Martin et al. 1991), then renamed as Rhodococcus sp. 51 T7 (Espuny et al. 1996), and now named as R. erythropolis 51 T7 (Marques et al. 2009). This bacterium has been confirmed to produce tetraesters of trehalose with short fatty acid chains (C8 C11). Mycobacterium is another group of trehalolipid-producing bacteria. But the trehalolipids produced by Mycobacterium are seldom reported as a biosurfactant, interests to them are mainly attributed to their role in pathogenicity and molecular immunology (Imasato et al. 1990; Ryll et al. 2001; Hunter et al. 2006; Ortiz et al. 2008, 2009; Zaragoza et al. 2009). Many species of this genus have been reported to produce trehalose-containing lipids, such as Mycobacterium tuberculosis, Mycobacterium bovis, Mycobacterium paraffinicum, Mycobacterium phlei, Mycobacterium flavescens, and Mycobacterium avium (Mompon et al. 1978; Batrakov et al. 1981; Fujita et al. 2005). In addition to their effects in health, bacteria of this genus also frequently reported as degraders of alkanes and polycyclic aromatic hydrocarbons (Hallas and Vestal 1978; Berekaa and Steinbuchel 2000; Kotani et al. 2006; Churchill et al. 2008; Vila and Grifoll 2009). The trehalolipids of mycobacteria might also involve in dissolving hydrophobic substrates. One example supporting this assumption is that a paraffin-oxidizing bacterium of M. paraffinicum can produce at least five trehalose lipids including the cord factor or two analogs (Batrakov et al. 1981).

3 Chemical Structures Like other glycolipids, trehalose lipids are composed of a carbohydrate group in combination with fatty acids groups. Differently, their hydrophobic moieties are more diverse, including aliphatic acids and hydroxylated branched-chain fatty acids (mycolic acids) of varied chain lengths. The numbers of hydrophobic chain in each molecule of trehalose lipids are usually 1, 2, and 4, forming mono-, di-, and tetraesters, correspondingly. Triesters can also be the main glycolipid component in some cases, such as in Mycobacteriurn fortuitum (Gautier et al. 1992). Among

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the trehalose lipids, the trehalose esters produced by R. erytropolis have been studied most extensively, in addition to the cord factors of Mycobacteria. Trehalose is a nonreducing sugar formed from two glucose units joined by a 1 1 alpha bond, giving it the name of a-D-glucopyranosyl-(1 ! 1)-a-D-glucopyranoside (Fig. 1). The bonding makes trehalose very resistant to acid hydrolysis, and therefore is stable in solution at high temperatures, even under acidic conditions. Trehalose is the carbohydrate group of cell wall glycolipids in Mycobacteria and Corynebacteria. Mycolic acids were first identified in an unsaponifiable lipid extract isolated from M. tuberculosis. They are complex hydroxylated branched-chain fatty acids with 60 90 carbon atoms, while those from other species (Corynobacterium, Nocardia) are shorter and named corynomycolic (22 36 carbons) or nocardomycolic (44 60 carbons) acids. They may also contain diverse functional groups such as methoxy, keto, or epoxy ester group and cyclopropane ring (Fig. 2). The structure, physiological function, and biosynthesis of mycolic acids have been reviewed by Barry et al. (1998).

3.1

Cord Factor: A Trehalose Diester from Mycobacteria

Trehalose lipids in mycobacteria have been extensively exploited in chemical composition (Mompon et al. 1978; Batrakov et al. 1981; Fujita et al. 2005). Of all HO O OH OH

• 2H2O

OH

HO O OH

Fig. 1 Trehalose structure

OH

O OH

OH COOH OH COOH

Fig. 2 Mycolic acids of the unsaturation and cyclopropane chains (cited from web page www. cyberlipid.org/fa/acid0006.htm)

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trehalose lipids, cord factor is the best known. It has two mycolic acids of variable length, esterified to the 6-hydroxyl group of each glucose to give trehalose 6,60 dimycolate (TMD) (Ryll et al. 2001) (Fig. 3). TDM is the most prominent and beststudied mycolic acid-containing glycolipid of mycobacteria. The structure varies greatly among mycobacterial species, and the mycolyl moiety is related with toxicity and antigenicity, thereby constituting potential virulence or immunostimulating mechanisms (Ryll et al. 2001; Hunter et al. 2006; Guidry et al. 2007; Ishikawa et al. 2009). With MALDI-TOF mass spectrometry, cord factors were characterized by Fujita et al.(2005) from nine species of human-virulent and nonvirulent mycobacteria as follows, M. tuberculosis H37Rv (ATCC 27294), M. tuberculosis Aoyama B (ATCC 31726), M. bovis BCG Tokyo 172 (ATCC 35737), M. bovis BCG Connaught (ATCC 35745), Mycobacterium intracellulare serotype 4 (ATCC 35767), HO HO

O

OH

O HO O

O HO

OH OH

O O OH O

Fig. 3 Trehalose 6,60 dimycolate(TDM ) from Isolated from M. tuberculosis

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M. intracellulare serotype 16 (ATCC 13950), Mycobacterium kansasii (ATCC 12478), Mycobacterium phlei (ATCC 11758), and Mycobacterium flavescens (ATCC 14474). They found that a-mycolic acid was a ubiquitous component among the mycobacterial species, with carbon numbers ranging from C74 to C88 and with two cyclopropane rings or equivalent double bonds. Ketomycolates were also widely distributed, with carbon numbers ranging from C76 to C91, but the ranges of the carbon number of ketomycolates differed greatly among the species (Fujita et al. 2005).

3.2

Trehalose Lipids from Rhodococcus

The surface-active lipids in rhodococci have been reviewed by Lang S (Lang 1999). The biosurfactants produced by members of the genus Rhodococcus is dominated by trehalose-containing glycolipids. They are quite diverse in the fatty acid moiety, such as dicorynomycolates, monocorynomycolates, and 2, 20 , 3, 4-tetraester (Kim et al. 1990; Espuny et al. 1996; Rapp et al. 1979; Kretschmer et al. 1982; Kretschmer and Wagner 1983). Additionally, succinoyl-trehalose and octaacyl-trehalose have also been detected in R. erythropolis (Uchida et al. 1989; Singer and Finnerty 1990; Tokumoto et al. 2009).

3.2.1

Trehalose Diesters

Trehalose Dimycolates from R. erythropolis DSM 43215 The trehalose lipids produced by R. erythropolis DSM 43215 were characterized as a-D-Glucopyranosyl-a-D-glucopyranoside-6,60 -di-(2-alkyl-3-hydroxy)-carboxylic ester(Rapp et al. 1979; Kretschmer et al. 1982) (Fig. 4). The bacterium was grown on 2% (w/v) n-alkanes (chain length C12 C18). The glycolipid was extracted from the biomass with n-hexane and purified by repeated chromatography on silica gel. O O OH OH OH

O O OH HO

O

O OH

OH O

OH

Fig. 4 a, a trehalose 6,60 dicorynomycolates from R. erythropolis DSM 43215 (Rapp et al. 1979). The mycolic acids range from C32H64O3 to C38H76O3.

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NMR revealed that its lipid moiety was consisted of saturated long-chain a-branched b-hydroxy fatty acids (mycolic acids) ranging from C32H64O3 to C38H76O3, of which C34H38O3 and C35H70O3 predominated. Prior to this study and afterwards, similar trehalose dimycolates were found in Rhodococcus, Nocardia, and Rhodochrous grown on glycerol.

Novel Trehalose Dimycolate from Rhodococcus opacus 1CP Glycolipids were purified from culture of R. opacus 1CP growing with n-decane as sole carbon source. They are characterized to be trehalose dimycolate with 1H-NMR spectroscopy. The fatty acids are nocardiomycolic acids of total chain lengths between 48 and 54 carbons (Fig. 5). In addition, in each mycolic acid, two double bonds exist (Niescher et al. 2006). These features (longer chain and unsaturation) distinguish the trehalose dimycolates from those of R. erythropolis DSM43215 (Rapp et al. 1979; Kretschmer et al. 1982) and Rhodococcus ruber IEGM231 (Philp et al. 2002).

Trehalose Lipids of Rhodococcus ruber IEGM231 Three kinds of glycolipids were purified from R. ruber culture, namely C1, C2, and C3. Each has analogs with varied chain lengths (Philp et al. 2002). Their structures were determined by NMR and ESI-MS (Fig. 6). In common, they share trehalose as the hydrophilic moiety. Among them, C1 was a typical trehalose dicorynomycolate with chain a-branched-b-hydroxy fatty acids (the total carbon number ranging from 34 to 46, with C40 as a main component), which is longer than that in OR1 OH

6 O 1

OH

OH O

HO

O

OH

R2O

OH Sugar residue OH CO R 1/2 =

2 6 H2C

(CH2)x

CH

CH

(CH2)y

CH

CH

(CH2)z

CH3

3

CH2 4 (CH2)n CH3

Fatty acid residue

Fig. 5 Structure of the trehalose dinocardiomycolate of R. opacus 1CP (Niescher et al. 2006). The mycolic acid components of carbon number ranging from C 48 to C 54 (n + x + y + z from 37 to 43)

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131 6 CH2OR1 O

OH

1

HO HO

3

O O

OH

CH2OR2 O

C1

OH

HO

R1 / R2 =

C

OH α CH

CH β

(CH2)m

(CH2)n

CH3

(CH2)m

CH3

CH3

m + n = 29 – 41 (centered at 35) O C2

R1 / R2 =

C

m = 13 – 15 (i.e. probably 14 + 12 and 14 + 16 with main component 14 + 14) O C3

R1 =

C

(CH2)m

CH3

R2 = H m = 10 – 14 (main component 12)

Fig. 6 Trehalose lipids of Rhodococcus ruber IEGM231 (Philp et al. 2002)

R. erythropolis DSM 43215 (Rapp et al. 1979). In the case of C2 component, the major component was equivalent to a molecule carrying two C16-fatty acid units and other analogs of the C11 C14 acids. The major C3 component is C14glycolipids containing fatty acids of both the saturated and unsaturated.

Succinoyl Trehalose Lipids from R. erythropolis SD-74 The bacterium strain SD-74 can abundantly produce extracellular trehalose lipids from n-alkanes. The characteristic of this trehalose lipid is succinic acid containing. Two main components named STL-1 and STL-2 have been characterized from culture of strain SD-74 (Uchida et al. 1989; Tokumoto et al. 2009). They are not only powerful surfactants, but they also show versatile biochemical actions, such as induction of human cell differentiation (Isoda et al. 1995; Sudo et al. 2000). The STL structure is primarily determined by NMR and GC-MS. STL1 and STL2 were 2,3,4,200 -di-O-succinoyl-di-O-alkanoyl-a,a-trehalose and 2,3,4-mono-

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O-succinoyl-di-O-alkanoyl-a,a-trehalose, respectively(Uchida et al. 1989). This is the first report of succinoyl trehalose lipids (STLs) (Uchida et al. 1989). Recently, the structure of STL1 produced by Rhodococcus sp. SD-74 was subjected to further characterization (Tokumoto et al. 2009). The exact structure of STL-1 was depictured on the basis of NMR, MALDI-TOF/MS, and GC-MS analyses. The STL-1 in this report was produced from n-hexadecane and was the main component. It was identified to be 3,4-di-O-alkanoyl-2-O-succinoyl-a-Dglucopyranosyl-20 -O-succinoyl-a-D-glucopyranoside (Fig. 7). The major fatty acid of STL-1 was C16, indicating that n-hexadecane as the carbon source is possibly directly incorporated into the carbohydrate moiety via terminal oxidation.

Trehalose Diesters and Triesters of Mycobacteriurn fortuitum M. fortuitum produced a family of trehalose-containing glycolipids named mycoside F (Gautier et al. 1992). Three main glycoconjugates were detected and their structures established as 2, 3-diacyl, 2, 3, 4- and 2, 3, 6-triacyl trehalose. The nature of the fatty acyl substituents identified primarily as 2-methyl-octadecen-2-oyl. In another case, a trehalose-containing glycolipid was detected in several strains of M. fortuitum and characterized as 2,3-di-O-acyltrehalose (DAT) (Ariza et al. 1994). In the report, lipid constituents were identified as a mixture of straight-chain (14 18 carbon atoms) and methyl-branched-chain (17 21 carbon atoms) fatty acyl groups. DAT was further fractionated by reverse phase TLC into four fractions that were designated DAT- I to IV. DAT-I contained 70 75% straight-chain acyl substituents (hexadecanoyl and octadecanoyl predominating) and 25 30% 2-methyl branched substituents (mainly 2-methyl octadecadienoyl). DAT-II was composed of a mixture in which the acyl groups were almost exclusively 2-methyl branched, with 2-methyl octadecadienoyl and 2-methyl octadecen-2-oyl predominating. DAT-III, which was the major isolated fraction, consisted of compounds in which the ratio linear to branched acyl groups varied between 0.8 and 0.9, 2-methyl octadecen-2-oyl, hexadecanoyl and octadecanoyl being the most abundant. Finally, HO O O O HO

O

O

HO

O OH

Fig. 7 Succinoyl trehalose lipids (STL 1) from R. erythropolis SD 74

CH3 O

O

(CH2)n O

O

O

O (CH2)n

CH3

OH O n = 14,12,10 OH

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DAT-IV comprised a mixture of DAT molecules containing mostly 2-methyl octadecadienoyl, 2-methyl octadecen-2-oyl, 2-methyl eicosadienoyl, and 2-methyl eicosen-2-oyl groups. Actually, early in 1964, trehalose lipids of M. fortuitum had been described having hydrophobia moiety like above described, i.e., C16:0 and sometimes with C19:0 and C20:0 in fatty acids moiety (Vilkas and Rojas 1964).

3.2.2

Trehalose Tetraesters Produced by Rhodococcus and Related Bacteria

In this section, the structure of the trehalose lipids produced by Arthrobacter, Micrococcus, and Rhodococcus is presented.

Arthrobacter sp. EK 1 (Passeri et al. 1991) Arthrobacter sp. EK 1 is a marine n-alkane-utilizing bacterium. After purification by column and thick layer chromatography, the main fraction, an anionic 2,3,4,20 trehalose tetraester, was obtained. The chain lengths of fatty acids ranged from 8 up to 14; furthermore, succinate was detected. The exact position of succinate is confirmed at C 2 atom of trehalose. This is the first report of succinate on trehalose of a glycolipid (Passeri et al. 1991).

Rhodococcus wratislaviensis BN38 (Tuleva et al. 2008) Glycolipids were produced by R. wratislaviensis BN38 when grown on 2% nhexadecane. The glycolipids were isolated by chromatography on silica gel columns, and the main product was characterized as 2,3,4,20 -trehalose tetraester with molecular mass of 876, esterified with two decanoic, one octanoic, and one succinic acid (Tuleva et al. 2008). Trace amounts of another 2,3,4,20 -trehalose tetraester of molecular mass 849 were also detected.

Micrococcus luteus BN56 (Tuleva et al. 2009) The soil strain M. luteus BN56 can produce a mixture of two trehalose tetraester biosurfactants when grown aerobically on n-hexadecane. The biosurfactants were extracted and purified from whole cultures and characterized by NMR and mass spectrometry (Fig. 8). The two major components consisted trehalose tetraesters with molecular mass of 876 and 848. In both cases, a,a-trehalose was linked at C-2 or C-4 with a succinic and at C-20 with a decanoic acid. The trehalose lipid with molecular mass of 876 was esterified at C-2, C-3, or C-4 with an octanoic and a decanoic acid. The other one with MW 848, of lower molecular mass, was esterified at the same positions with two octanoic acids (Tuleva et al. 2009).

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Fig. 8 Chemical structure of trehalose tetraesters produced by M. luteus BN56 (n R2 succinic acid) (Tuleva et al. 2009)

6 11;

Rhodococcus erythropolis 51 T7 (Marques et al. 2009) The chemical structure of trehalose lipids produced by strain 51 T7 was further studied by LC-MS, with tetradecane as the carbon source (Marques et al. 2009). This analysis revealed that this surfactant is a mixture of at least six components, with the pseudomolecular ions being between m/z 905 and 821 (using negative ion mode). The most abundant component is the pseudomolecular ion 876, occupying 43.9% of the total lipids after 72 h in culture. This molecular weight may correspond to either trehalose succinic acid C9 C9 C10 or trehalose succinic acid C11 C10 C7. The remaining components had a lower and more constant concentration during growth. Worth to note, the trehalose tetraesters of R. erythropolis 51 T7 differed with the results of a previous report in the main component (Espuny et al. 1996). The authors postulated that this may have resulted from variations in the carbon source used.

3.2.3

Octaacyltrehalose from Rhodococcus sp. H13-A

Growing with hexadecane, Rhodococcus sp. H13-A produced a mixture of trehalose lipids with one major and ten minor components(Singer et al. 1990). The components of hydrophobic tail are relatively complex, constituting of normal C10 C22 saturated and unsaturated fatty acids, C35 C40 mycolic acids, hexanedioic and dodecanedioic acids, and 10-methyl hexadecanoic and 10-methyl octadecanoic acids. The major glycolipid was identified as 2,3,4,6,20 ,30 ,40 ,60 -octaacyltrehalose, plus minor components of di-, tetra- and hexa-acyltrehalose derivatives.

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4 Physicochemical Property Physicochemical properties of trehalose lipids have been mainly examined in the genus Rhodococcus, while the glycolipids from mycobacteria are seldom investigated in this aspect due to their long hydrophobic moiety tightly attached on cell wall and limitations as a practical biosurfactant. Recently, some trehalose lipids come to purification (Tuleva et al. 2008, 2009; Marques et al. 2009; Tokumoto et al. 2009; Peng et al. 2007; Niescher et al. 2006; Kretschmer et al. 1982; Kretschmer and Wagner 1983; Kim et al. 1990). Most of them showed strong surface activity by lowering water surface tension below 30 mN m 1 (ranging from 19 to 43 mN m 1), lowering the interfacial tension against hexadecane (decane or kerosene) to 5 mN m 1, even <1 mN m 1 (Table 1), while the CMC values can reach 0.7 mg l 1 in case of dicorynomycolates (Kretschmer and Wagner 1983), and the CMC values of most trehalose lipids varied from 0.7 to 37 mg l 1. Examples are as follows. The trehalose corynomycolates generated by R. erythropolis DSM43215 showed extremely low CMC in high-salinity solutions, and the interfacial properties were stabile in solutions with a wide range of pH and ionic strength(Kretschmer et al. 1982). The dicorynomycolates reduced interfacial tension from 44 to 18 mN m 1 and are less sensitive to salt concentrations than synthesized surfactants, therefore of potential application in enhanced oil recovery. Succinoyl trehalose lipids (STL) from R. erythropolis SD-74 are quite efficient to lower water surface tension. The estimated CMC and gCMC values for STL-1 were 5.6  10 6 M (equivalent to 5 mg l 1) and 19.0 mN m 1, and those for sodium salt (Na-STL-1) were 7.7  10 6 M and 23.7 mN m 1, respectively. Thus, STL-1 holds an excellent surface activity at remarkably low concentrations (Tokumoto et al. 2009). The glycolipid synthesized by Rhodococcus species H13-A (Singer et al. 1990) exhibited a CMC of 1.5 mg l 1 and minimum interfacial tension value of 0.02 mN m 1 against decane, even much lower in interfacial tension to 6  10 5 mN m 1 in the presence of pentanol. In the case of the glycolipids purified from M. luteus BN56, they own CMC value of 25 mg l 1 and can reduce the surface tension of water to 24.1 mN m 1 and lower the interfacial tension against hexadecane to 1.7 mN m 1. In addition, they have emulsion-stabilizing activity.

5 Biological Activity Biosurfactants play an essential role in bacterial swarming motility and participate in cell signaling and differentiation as well as in biofilm formation. In addition, they can increase the bioavailability of hydrophobic substrates, trap heavy metals, and function as an antibiotic (Singh and Cameotra 2004; Rodrigues et al. 2006). Trehalose lipids accept the general merits being a biosurfactant. Moreover, some particular functions also exist, and worthy to be noted.

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Trehalose Lipids in Mycobacteria

Trehalose lipids in mycobacteria draw attentions due to their role in pathogenicity and molecular immunology (Imasato et al. 1990; Ryll et al. 2001; Hunter et al. 2006; Ortiz et al. 2008, 2009; Zaragoza et al. 2009). TDM (cord factor) is the most abundant, granulomagenic, and toxic lipid in the cell surface of virulent M. tuberculosis(Hunter et al. 2006). TDM has been identified as the factor responsible for cord formation. It has been also detected in nontuberculous mycobacteria, including M. avium complex species (Fujita et al. 2005). In the mycobacterial cell envelope, together with arabinogalactan mycolate and trehalose monomycolate, TDM forms an integral part of the cell wall skeleton, resulting in highly hydrophobic cell surface properties and acid-fastness (Barry et al. 1998). In addition, it is believed that TDM is responsible for the low permeability of the membranes conferring appreciable drug resistance to the organisms. On the other hand, TDM and its stereoisometric derivatives (TDCMs) were evidenced to inhibit tumor metastasis, and TDCMs are more potential to suppress tumor growth and inhibit tumor metastasis than TDM (Watanabe et al. 1999).

5.2

Trehalose Lipids from Rhodococcus

Beside their known industrial applications, trehalose lipids from Rhodococcus recently attracted attention due to their functions in cell membrane interaction and the prospects as a therapeutic agent (Isoda et al. 1995; Zaragoza et al. 2009, 2010; Aranda et al. 2007; Harland et al. 2009; Ortiz et al. 2008, 2009; Imasato et al. 1990). l

l

l

The succinoyl trehalose lipid (STL-1) of R. erythropolis SD-74 shows effects on cell differentiation. Isoda et al. (1995) reported that STL-1 markedly inhibited the growth of a human monocytoid leukemic cell line and induced its morphological alteration along a monocyte-macrophage lineage. STL-1 markedly increased differentiation-associated characteristics in macrophage, such as phagocytic activities in U937. Furthermore, the U937 cells activated with STL-1 exhibited cytotoxic activity against human carcinoma cell line, while it has low cytotoxicity against normal human cells (Isoda et al. 1995). Hemolytic activity of a succinoyl-containing trehalose lipid: the trehalose lipid produced by Rhodococcus sp. was observed to cause the swelling of human erythrocytes followed by hemolysis at concentrations well below its CMC value. It works by a colloid-osmotic mechanism, most likely by formation of enhanced permeability domains, or “pores” enriched in the biosurfactant, within the erythrocyte membrane (Zaragoza et al. 2010). Interactions with phosphatidylethanolamine membrane: Ortiz et al. (2008) purified a trehalose lipid from Rhodococcus sp. and examined its effect on the thermotropic and structural properties of phosphatidylethanolamine membranes of different chain length and saturation. They find that the trehalose lipid

Trehalolipids

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l

137

presents good miscibility both in the gel and the liquid crystalline phases of the membrane and affects the gel to liquid crystalline phase transition. The trehalose lipid was suggested to incorporate into the membrane bilayers and produce structural perturbations, which might affect the function of the membrane. Similar conclusions were drawn based on the results with phosphatidylserine membranes (Ortiz et al. 2009): Mechanism of membrane permeabilization by the STL (Zaragoza et al. 2009): The partition constant of the trehalose lipid to palmitoyloleoylphosphatidylcholine membranes indicates that trehalose lipid behaves as a weak detergent, which prefers membrane incorporation over micellization. Addition of the trehalose lipid to large unilamellar vesicles results in a size-selective leakage of entrapped solutes to the external medium. Further studies support that the trehalose lipid incorporates into phosphatidylcholine membranes and segregates within lateral domains, which may constitute membrane defects or “pores.” Synthetic trehalose glycolipids confer desiccation resistance to supported lipid monolayers (Harland et al. 2009). Harland et al. (2009) presented the first synthetic trehalose glycolipids capable of providing desiccation protection to membranes of which they are constituents. They deduced that interactions between the trehalose headgroup and surrounding molecules are the determining factor in dehydration protection.

6 Biosynthesis The biosynthesis pathways of biosurfactant have been extensively studied in rhamnolipids and lipopeptides, and related genes have been determined in some cases (Nakano et al. 1991, 1992; Ochsner et al. 1994; Campos-Garcia et al. 1998). However, the corresponding researches on trehalose lipids are hard to be retrieved from literatures. With an aim to clarify the synthesis pathway of trehalose dicorynomycolates, Kretschmer and Wagner (1983) characterized biosynthetic intermediates of the trehalose lipids in R. erythropolis DSM 43215. Free corynomycolic acid intermediates were observed. The corynomycolic acid moiety and the trehalose moiety therefore are synthesized independently and are subsequently esterified. Trehalose is first esterified to form the monomycolate and then the dimycolate. Based on the intermediates, a biosynthetic pathway was suggested (Fig. 9) (Kretschmer and Wagner 1983).

7 Production The production of most trehalose lipids is growth-associated, whereas production is also observed under growth-limiting conditions or by resting cells. Compared to rhamnolipids, the yield of trehalose lipids reported in the literature is lower, usually below 3 g l 1, considering the production of purified products.

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Z. Shao N – TETRADECANE N – TETRADECANEOL N – TETRADECANE ALDEHYD N – TETRADECANOIC ACID FATTY ACID ELONGATION CIS – Δ9 – TETRADECENOIC ACID

N – EICOSANOIC ACID LONG FATTY ACID CONDENSATION β – OXIDATION

3 – OXO – 2 – DODECANYL – DOCOSANOIC ACID ACETYL – Co A GLUCONEOGENESE 3 – HYDROXY – 2 – DODECANYL – DOCOSANOIC ACID (= CORYNOMCOLIC ACID) GLUCOSE – 6 – p +UDP – GLUCOSE PHOSPHATE TREHALOSE – 6 – P

TREHALOSE – 6 – PHOSPHATE – 6 – CORYNOMYCOLATE PHOSPHATE TREHALOSE – 6 – CORYNOMYCOLATE TREHALOSE – 6,6′ – DICORYNOMYCOLATE

Fig. 9 Trehalose dicorynomycolate synthesis pathway from n tetradecane by Rhodococcus ery thropolis DSM 43215 (Kretschmer and Wagner 1983)

7.1 l

Substrates Used for Trehaloselipids Production

n-alkanes: In the case of oil degrading bacteria of Rhodococcus, n-alkanes are usually the optimal substrates for glycolipids production. Although bacteria of Rhodococcus can utilize alkanes of a wide length range, C14 and C16 alkanes

Trehalolipids

l

139

seem to be the best for high yielding (Niescher et al. 2006), especially for the trehalose lipids with medium length chains(Tokumoto et al. 2009). Nonalkanes: Trehalose dicorynomycolates have also been produced in the absence of n-alkanes or other lipophilic carbon sources by Brevibacterium vitarumen 12143 (Laneelle and Asselineau 1976), Corynebacterium diphtheriae (Adam et al. 1967), and different pathogenic Mycobacteriaceae (Asselineau and Asselineau 1978).

7.2

Cell Wall Association

The usual pattern of production of nonionic trehalose glycolipids is generally cell wall-associated (Suzuki et al. 1969; Kretschmer et al. 1982). Usually, the lipids of long-chain fatty acids tend to be cell wall-associated. Especially at early stage of cultivation, the products mainly attach to the cell wall. Growing on n-alkanes as a sole source of carbon and energy, R. fascians produced both an extracellular and cell-bound surface-active mixture of trehalose lipids that reduced the surface tension of water to 32 mN m (Yakimov et al. 1999). When R. erythropolis strain 51 T7 was cultivated with both alkanes and waste lubricant oil as a sole carbon substrate, it produced extracellular glycolipids with surface activity. Rhodococcus sp. strain 51 T7 produced trehalose lipids between 0.48 and 1.12 g l 1(Espuny et al. 1996). However, in a previous report of this strain (named Coryneform bacterium 51 T7 at first), trehalose lipids are associated mainly with the cell wall on incubation up to 80 h. After 80 h, the trehalose lipids are mainly extracellular (Martin et al. 1991).

7.3

Growth-Associated Production

Growth-associated production of trehalose lipids is popular (Table 1), such as the production of a trehalose dicorynomycolate in R. erythropolis DSM43215 (Rapp et al. 1979). The production of a trehalose dicorynomycolate in R. opacus 1CP is also growth-associated (Niescher et al. 2006) (Fig. 10). However, the formation of anionic trehalose ester by R. erythropolis DSM43215 seems to be uncoupled from growth and occurs in the stationary phase (Ristau and Wagner 1983; Kim et al. 1990). The growth-uncoupled production also was observed in a paraffin-oxidizing bacterium R. erythropolis cultivated in shake flasks on a mixture of C14 and C15 n-alkanes or kerosene. The growth-limiting conditions, such as nitrogen deficiency, caused the formation of a, a-trehalose-2,3,4,20 tetraesters. Contrarily, also being a trehalose 2,3,4,20 -tetraester, one of the Rhodococcus sp 51 T7 is produced in growth-associated mode. When grown on hydrocarbon, cells were highly segmented and accumulated lipid granules in the cytoplasm. Under

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Fig. 10 Correlation of cell growth (protein), alkane consumption, and glycolipid formation of R. opacus 1CP (Niescher et al. 2006). Batch cultures (20 ml), each containing mineral medium and 1% (w/v) n tetradecane, were inoculated with identical amounts of glucose grown R. opacus 1CP and incubated at 30 C with constant shaking (130 rpm)

optimal concentrations of sodium nitrate, potassium phosphate and iron (2.5 g l 1, 1.5 g l 1 and 0.01 g l 1, respectively), production was increased from 0.5 to 3 g l 1 (Espuny et al. 1996).

References Adam A, Senn M, Vilkas E, Lederer E (1967) Spectrome´trie de masse de glycolipides. Eur J Biochem 2:460 468 Anderson RJ, Newman MS (1933) The chemistry of the lipids of tubercle bacilli: xxxiii. isolation of trehalose from the acetone soluble fat of the human tubercle Bacillus. J Biol Chem 101:499 504 Aranda FJ, Teruel JA, Espuny MJ, Marques A, Manresa A, Palacios Lidon E, Ortiz A (2007) Domain formation by a Rhodococcus sp. biosurfactant trehalose lipid incorporated into phosphatidylcholine membranes. Biochim Biophys Acta 1768:2596 2604 Ariza MA, Martin Luengo F, Valero Guillen PL (1994) A family of diacyltrehaloses isolated from Mycobacterium fortuitum. Microbiology 140(Pt 8):1989 1994 Asselineau C, Asselineau J (1978) Trehalose containing glycolipids. Prog Chem Fats Other Lipids 16:59 99 Barry CE 3rd, Lee RE, Mdluli K, Sampson AE, Schroeder BG, Slayden RA, Yuan Y (1998) Mycolic acids: structure, biosynthesis and physiological functions. Prog Lipid Res 37:143 179 Batrakov SG, Bv R, Koronelli TV, Bergelson LD (1981) Two novel types of trehalose lipids. Chem Phys Lipids 29:241 266

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Mannosylerythritol Lipids: Microbial Production and Their Applications J. Arutchelvi and M. Doble

Contents 1 2

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146 Microbial Sources and Structural Diversity of MEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 147 2.1 MEL A Producers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 2.2 MEL B Producers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 2.3 MEL C Producers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 2.4 Effect of Addition of Different Sugars on the Type of MEL Produced . . . . . . . . . . . . 154 3 Significance of MEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154 4 Genetic Regulation and Biosynthesis of MEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 5 Evolutionary Relationship Among MEL Producers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 6 Bioprocesses Used for MEL Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 160 6.1 Factors Affecting MEL Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 7 Phase Behavior of MEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 7.1 Formation of Thermodynamically Stable Vesicles and Coacervates by MEL . . . . . 164 7.2 Multilamellar Vesicles and Large Unilamellar Vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . 165 7.3 Lyotropic Liquid Crystalline Phases of MEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166 7.4 Self Assembled Monolayer Structures of MEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 8 Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 8.1 Antimicrobial Activity of MEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 8.2 MEL Induces Cell Differentiation and Apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170 8.3 Purification of Glycoproteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170 8.4 Vehicles for Gene Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170 8.5 Inhibition of Ice Agglomeration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171 8.6 Cosmetic Applications of MEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171 9 Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171 10 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173

J. Arutchelvi and M. Doble (*) Department of Biotechnology, Indian Institute of Technology Madras, Chennai 600 036, India e mail: [email protected]

G. Sobero´n‐Cha´vez (ed.), Biosurfactants, Microbiology Monographs 20, DOI 10.1007/978 3 642 14490 5 6, # Springer Verlag Berlin Heidelberg 2011

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Abstract Mannosylerythritol lipids (MELs) are surface-active compounds synthesized by yeast strains of the genus Pseudozyma sp. or Ustilago sp., from soybean oil or n-alkane (over 100 g/L). The former strain produces it as the major while the latter as the minor component. They belong to the class of glycolipids. Although MELs have been known for more than five decades, they recently regained attention due to their environmental compatibility, mild production conditions, structural diversity, self-assembling properties, versatile biochemical functions, and high yield when compared to other biosurfactants. In this chapter, structural diversity of MELs, genetic diversity of MEL producers, various fermentation conditions, downstream steps involved in their separation, factors affecting the amount and type of MEL produced, and self-assembling properties and their applications are discussed. The biosynthetic pathways and the genetic regulation of the MEL production are also included.

1 Introduction Surfactants are amphiphilic molecules exhibiting surface and interfacial activity between solids, liquids, and gases. They have both polar head and nonpolar tail groups. They tend to aggregate at the interfaces and form organized molecular assemblies, monolayers, micelles, vesicles, and membranes (Kitamoto et al. 2009). Biosurfactants (BSs) are produced by a variety of microorganisms including bacteria, fungi, and yeast from various substrates including sugars, glycerol, oils, hydrocarbons, and agricultural waste (Lin 1996). BSs are required for the swarming and gliding motility of the microorganisms (Kearns and Losick 2003). In addition, BSs enhance microbial adhesion to hydrophobic surfaces, increase the bioavailability of water-insoluble substrates, reduce the heavy metal toxicity (Mulligan 2004), and often show antimicrobial activity (Ron and Rosenberg 2001). There are five main classes of BSs, namely lipopeptides; glycolipids; fatty acids including neutral lipids and phospholipids; polymeric; and particulate (Neu 1996; Desai and Banat 1997). Growing interest in the field of BSs can be attributed to their environmental compatibility, their preparation from renewable resources, production conditions, structural diversity, and variety of functions. Mannosylerythritol lipids (MELs) are representative of glycolipids. BS containing 4-O-b-D-mannopyranosyl-meso-erythritol as the hydrophilic group and a fatty acid and/or an acetyl group as the hydrophobic moiety is known as mannosylerythritol lipid (MEL) (Fig. 1) (Kitamoto 2008). Based on the degree of acetylation at C-4 and C-6 positions, and their order of appearance on the thin layer chromatography (TLC), the MELs (from Pseudozyma aphidis) are classified as MEL-A, -B, -C, and -D. MEL-A represents the diacetylated compound while MEL-B and MEL-C are monoacetylated at C-6 and C-4 positions, respectively. The completely deacetylated structure is known as MEL-D (Rau et al. 2005a). Mixtures of these four MELs, but predominantly MEL-A and MEL-B, were isolated from Candida antarctica strain T34 (Kitamoto et al. 1990b). MEL is a glycolipid produced by

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Fig. 1 Structure of MEL (MEL A: R1 R2 Ac; MEL B: R1 Ac, R2 H; MEL C: R1 H, R2 Ac: n 6 10)

Ustilago sp. (as a minor component along with cellobiose lipid (CL)) (Haskins et al. 1955), Schizonella melanogramma (shizonellin) (Deml et al. 1980), Pseudozyma sp. (as a major component) (Kitamoto et al. 1990a, b), Kurtzmanomyces sp. (Kakugawa et al. 2002), and different strains of the genus Candida. The synthesis of MEL is not growth-associated, and it can also be produced by using resting (stationary phase) cells of yeast. MEL acts as an energy storage material in the yeast cells similar to triacylglycerols (Kitamoto et al. 2002). MELs are shown to reduce the surface tension of water to less than 30 mN/m (Kim et al. 2006; Kitamoto et al. 1993b; Konishi et al. 2007b). Even though BSs have a wide range of applications, they are not yet commercially viable, mainly due to their low yield leading to high production cost. However, MEL (165 g/L) and sophorolipid (above 400 g/L) are produced in high quantity (Kitamoto et al. 2002) and hence may become economically attractive.

2 Microbial Sources and Structural Diversity of MEL MELs are secreted by several microorganisms and they are first noted as oily compounds in the cultured suspension of Ustilago maydis PRL-627 (Haskins et al. 1955). This microorganism was shown to produce MEL along with CL when glucose was used as the carbon source (Haskins et al. 1955; Fluharty and O’Brien 1969). Bhattacharjee et al. (1970) characterized MEL as a glycolipid, especially as a mixture of partially acylated derivative of 4-O-b-D-mannopyranosyl-D-erythritol, containing C2:0, C12:0, C14:0, C14:1, C16:0, C16:1, C18:0, and C18:1 fatty acids as the hydrophobic groups. S. melanogramma was the second microorganism identified as a MEL producer (schizonellin), but not in association with CL (Deml et al. 1980). Table 1 summarizes the structural variants of the MELs and the corresponding microbial sources. These variants arise due to the following reasons: l l l

Number and position of the acetyl group on mannose or erythritol or both Number of acylation in mannose Fatty acid chain length and their saturation

Sugar 4-O-b-D-mannopyranosyl -DerythritoI Candida (Pseudozyma) sp B7 4-O-b-D-mannopyranosyl-Derythritol Pseudozyma antarctica T-34 4-O-b-D-mannopyranosylerythritol Ustilago maydis DSM 4500 and ATCC 4-O-b-D-mannopyranosyl -DerythritoI 1482 Schizonella melanogramma 4-O-b-D-mannopyranosyl-Derythritol Kurtzmanomyces sp I-11 6-O-b-D-mannopyranosyl(1!4)-O-meso-erythritol Candida sp SY16 6-O-b-D-mannopyranosyl(1!4)-O-meso-erythritol Pseudozyma aphidis DSM 70725 4-O-b-D-mannopyranosyl(1!4)-O-meso-erythritol Pseudozyma rugulosa NBRC 10877 4-O-b-D-mannopyranosyl(1!4)-O-meso-erythritol Pseudozyma hubeiensis KM-59 4-O-b-D-mannopyranosyl(1!4)-O-meso-erythritol Pseudozyma shanxiensis 4-O-b-D-mannopyranosyl(1!4)-O-meso-erythritol P tsukubaensis JCM 10324T 1-O-b-D-mannopyranosyl(1!4)-O-meso-erythritol P fusiformata 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol Pseudozyma graminicola CBS 10092 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol P antarctica and P parantarctica 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol P antarctica and P rugulosa 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol Ustilago scitaminea NBRC 32730 (olive 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol oil) Ustilago cynodontis NBRC 7530 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol (soybean oil)

Microorganisms Ustilago nuda PRL-627,

Table 1 Microbial sources and structural diversity of MEL

Morita et al (2006b) Konishi et al (2007b, 2008) Fukuoka et al (2007c)

Morita et al (2008a) Fukuoka et al (2007c)

MEL A MEL C (65%) MEL C MEL B with New diastereomer of sugar MEL A MEL C (85%) Monoacylated MEL

C8:0 (28 09%), C10:0 (21 68%), C10:1 (22 94%) C6, C12, and C16 C16:0, C16:1, C16:2, and C14:1 C8, C12, C14

C6:0, C8:0, C12:0, C12:1, C14:0, and C14:1 C8:0, C10:0, C12:0, C14:0, C14:1

Morita et al (2008c)

C14:0 and C16:0

MEL C

Morita et al (2009a)

Konishi et al (2007a)

Fukuoka et al (2008), Morita et al (2007) Morita et al (2008a)

C18:1, C18:0, C10:0, C10:1, C16:0, and Triacylated MEL C8:0 C8:0 and C12:0 MEL B

C6 and C8

Rau et al (2005a)

MEL A

C10:0, C10:1, and C8:0

Kakugawa et al (2002)

Deml et al (1980)

Kitamoto et al (1990b, 2001a) Spoeckner et al (1999)

References Bhattacharjee et al (1970) Kawashima et al (1983)

Kim et al (1999)

MEL B7

Type of MEL (major component) –

MEL A (major component) and MEL C8:0 (27 26%), C10:0 (21 28%), B (it produce all 4 types) C10:1 (27 22%) C14:1 (43%), C6:0 (20%) and C16:1 MEL A (12%) C14:0, C16:1, C16:0, C18:0, C18:1 Schizonellin A and schizonellin B (similar to MEL A and B) C8:0 (36 4%), C12:0 (11 9%), C14:2 MEL I-11 (25 9%) C6:0, C12:0, C14:0 and C14:1 MEL A

Fatty acid C2:0, C12:0, C14:0, C14:1, C16:0, C16:1, C18:0 and C18:1 C7 – C14

148 J. Arutchelvi and M. Doble

P antarctica JCM 10317 (in the presence of sucrose) U maydis NBRC 5346 (in the presence of sucrose) Ustilago scitaminea NBRC 32730 (in the presence of sucrose) P Siamensis CBS 9960 (in the presence of sucrose) P Siamensis CBS 9960 (in the presence of Safflower oil) Pseudozyma parantarctica JCM 11752 (in the presence of mannitol)

4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol 4-O-b-D-mannopyranosyl(1!4)-O-meso-erythritol 4-O-b-D-mannopyranosyl(1! 4)-O-meso-erythritol 6-O-b-D-mannopyranosylD-mannitol (MML)

MEL A MEL A MEL B MEL C MEL C MEL A with different hydrophilic part and is more hydrophilic than MEL C

C10:0 and C12:0 C6:0, C14:0 and C16:0 C8:0, C10:0 and C12:0 C16:0 and C18:0 C14:2 and C16:0 C8:0 and C10:0 (fatty acid composition similar to MEL A)

Morita et al (2009b)

Rodrigues et al (2006)

Morita et al (2009c)

Morita et al (2009c)

Morita et al (2009c)

Morita et al (2009c)

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The acyl residues in MEL range from C7 to C14 and their proportions vary depending upon the carbon source used. For example, Candida sp. B-7 (currently known as Pseudozyma antarctica) produced 4-O-(20 ,60 -di-O-acyl-b-D-mannopyranosyl)-D-erythritol with high yield (25 36 g/L) when it was grown on n-alkane or triglycerides at 30 C for 7 days but not with carbohydrate as the carbon source (Kawashima et al. 1983). A MEL producer is identified using fluorescence microscopy. Figure 2 shows the photographs of Pseudozyma rugulosa NBRC 10877, a novel producer of MEL, stained with Nile Red as a lipid probe. The cells grown on soybean oil as the sole carbon source gave a strong orange-red fluorescence, while the cells grown on glucose gave a yellow fluorescence. Nile Red is known to emit a fluorescence of yellow-gold (582 nm) and orange-red (617 nm) in the presence of neutral lipids and polar lipids, respectively, when excited at 488 nm (Greenspan and Fowler 1985; Vejux et al. 2005). Nile red stains MEL and soybean oil, which gives orange-red and yellow colors, respectively. However, Nile red can stain lipids other than MEL also.

Nile red

Soybean oil

Glucose

DIC

Fig. 2 Fluorescence microscopic examination with Nile red staining (right panels). Differential interference contrast (DIC, left panels) (taken from Morita et al. 2006b)

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151

MEL-A Producers

Kurtzmanomyces sp. I-11 produces a novel MEL composed of a sugar moiety, 6-Ob-D-mannopyranosyl- (1!4)-O-meso-erythritol, and a mixture of fatty acids [C8:0 (36.4%), C12:0 (11.9%) and C14:2 (25.9)] (Kakugawa et al. 2002) as the lipophilic group. C. antarctica KCTC 7804, which is isolated from an oil-contaminated site, produced a MEL containing 6-O-acetyl-2,3-di-O-alkanoyl-b-D-mannopyranosyl(1!4)-O-meso-erythritol. The lipophilic group of this MEL is a mixture of hexanoic, dodecanoic, tetradecanoic, and tetradecanoic fatty acids. The acetyl group was linked at the C-6 position of the D-mannose (Kim et al. 1999). P. rugulosa NBRC 10877 is reported to produce from soybean oil, a mixture of MEL-A (68%), MEL-B (12%), and MEL-C (20%) consisting of C8 and C10 fatty acids as the hydrophobic moiety (Morita et al. 2006b).

2.2

MEL-B Producers

Pseudozyma tsukubaensis also produces an unusually different carbohydrate structure, namely 1-O-b-(20 ,30 -di-O-alka(e)noyl-60 -O-acetyl-D-mannopyranosyl)-D-erythritol (Fig. 3). The configuration of this erythritol moiety in this MEL is opposite to that of the known MEL-B or to other MELs reported (Fukuoka et al. 2008). A smut fungus, Ustilago scitaminea NBRC 32730 (Morita et al. 2009a), on sugar cane was found to produce large amount of glycolipids, and the major component of the mixture was characterized as MEL-B, 4-O-b-(-20 ,30 -di-O-alka(e)noyl-60 -O-acetylD-mannopyranosyl)-erythritol. The fatty acid chain length of MEL-B produced by the smut fungus from olive oil was C8 and C10, whereas P. tsukubaensis produced MEL-B with the chain length of C8 and C14. Hence, U. scitaminea is a novel MEL producer (Morita et al. 2009a).

Fig. 3 Diastereomer of MEL B from P. tsukubaensis; n 6 12

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MEL-C Producers

MEL-C is produced by Pseudozyma hubeiensis and the major fatty acids in it are C6, C12, and C16, which are different from those of the other MELs reported so far (Konishi et al. 2007b). Pseudozyma shanxiensis produces a hydrophilic MEL, consisting of a mixture of 4-O-[(20 , 40 -di-O-acetyl-30 -O-alka(e)noyl)-b-D-mannopyranosyl]-D-erythritol (Fig. 4) and 4-O-[(40 -O-acetyl-30 -O-alka(e)noyl-20 -O-butanoyl)-b-D-mannopyranosyl]-D-erythritol (Fig. 5). This has much shorter chain (C2 or C4) at the C-20 position of the mannose moiety when compared to other classes of MELs that comprise of a medium-chain length acid (C8 C14) at this position (Fukuoka et al. 2007c; Morita et al. 2008b). Pseudozyma siamensis CBS 9960 is found to accumulate higher amounts of glycolipid than its closely related species, P. shanxiensis, which is a known MEL-C producer. The former produces a mixture of different types of MEL-C, namely (>84% of the total) 4-O-[(20 ,40 -di-O-acetyl30 -O-alka(e)noyl)-b-D-mannopyranosyl]-D-erythritol and 4-O-[(40 -O-acetyl-30 -Oalka(e)noyl-20 -O-butanoyl)-b-D-mannopyranosyl]-D-erythritol. This MEL-C possesses a short-chain length acid (C2 or C4) at the C-20 position and a long-chain length acid (C16) at the C-30 position of the mannose moiety. Thus, the hydrophobic part is considerably different from that of the conventional MELs, which have two medium-chain length acids (C10) at the C-20 and C-30 positions (Morita et al. 2008a). P. antarctica and P. rugulosa, with soybean oil as the carbon source, produce the most hydrophobic triacylated MEL, namely 1-O-alka(e)noyl4- O-[(40 ,60 -di-O-acetyl-20 ,30 -di-O-alka(e)noyl)-b-D-mannopyranosyl]-D-erythritol

Fig. 4 Structure of 4 O [(20 ,40 di O acetyl 30 O alka (e)noyl) b D mannopyranosyl] D erythritol; R2 Ac; n 12 16

Fig. 5 Structure of 4 O [(40 O acetyl 30 O alka(e)noyl 20 O butanoyl) b D mannopyranosyl] D erythritol; R2 Ac; n 12 16

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(Fig. 6) (Fukuoka et al. 2007b). P. antarctica T-34 produces a hydrophilic monoacylated MEL, 4-O-(30 -O-alka(e)noyl-b-D-mannopyranosyl)-D-erythritol (Fig. 7), with glucose as the carbon source. This MEL reduces the surface tension of water to 33.8 mN/m at a critical micelle concentration (CMC) of 3.6  10 4 M, and it has a hydrophilic lipophilic balance of 12.15. The CMC is 100-fold higher than that of the other MELs reported (Fukuoka et al. 2007c). A strain of P. hubeiensis KM-59 produces a hydrophilic MEL-C, namely (4-O-[40 -O-acetyl-20 ,30 -di-O-alka(e)noylb-D-mannopyranosyl]-D-erythritol) (Konishi et al. 2008). Pseudozyma graminicola CBS 10092 also produces MEL-C. It has C6, C8, and C14 acids and is considerably different from the other MEL-C producing Pseudozyma strains including, P. antarctica and P. shanxiensis (Morita et al. 2008a). Ustilago cyanodontis NBRC 7530 (Morita et al. 2008c) was identified as a MEL producer, with fatty acid chain lengths of C14 and C16. It retains the same hydrophilic group as that of the conventional MEL-C. Different species of MEL-C producers show variation in their fatty acid chain length.

Fig. 6 Structure of Triacylated MEL; n m 6 16

Fig. 7 Structure of Monoacylated MEL; n 4 14

6 10;

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Effect of Addition of Different Sugars on the Type of MEL Produced

P. antarctica JCM 10317 and U. maydis NBRC 5346 produced MEL-A with the same hydrophilic group, when they are cultivated in the presence of simple sugars including glucose and sucrose (Morita et al. 2009c), as that of the hydrophilic group of MEL-A produced in the presence of vegetable oils. U. Scitaminea NBRC 32730 and P. siamensis CBS 9960 produced MEL-B and MEL-C, respectively, with glucose and sucrose as the carbon source. However, the major fatty acids of sucrose-derived MELs from the above mentioned organisms are quite different from those of the vegetable oil-derived MELs (Morita et al. 2009c). Pseudozyma parantarctica JCM 11752, P. antarctica JCM 10317, and P. rugulosa NBRC 10877 are reported to produce a novel type of glycolipid, which is more hydrophilic in nature than conventional MEL-C when they are cultivated in the presence of mannitol (Morita et al. 2009b). The glycolipid from P. parantarctica JCM 11752 is 6-O-[(40 ,60 - di-O-acetyl-20 ,30 -di-O-alka(e)noyl)-b-D-mannopyranosyl]-D-mannitol (Fig. 8), namely Mannosylmannitollipid (MML) (Morita et al. 2009b). Table 2 summarizes the surface activities of various homologs of MEL.

3 Significance of MEL The increasing interest in MELs could be attributed to their highest yield (of over 100 g/L) when compared to other classes of BSs, their pharmaceutical applications (Shibahara et al. 2000; Zhao et al. 2001) and versatile biochemical functions Table 2 Surface activities of various MEL homologs Type of MEL Critical micelle concentration (M) MEL A Sucrose derived MEL A from P. antarctica JCM 10317 Sucrose derived MEL A from U. maydis NBRC 5436 MEL B Sucrose derived MEL B from U. Scitaminea NBRC 32730 MEL C (from P. Hubeiensis KM 59) MEL C (from P. graminicola CBS 10092) MEL C (from P. siamensis CBS 9960) Sucrose derived MEL C from P. Siamensis CBS 9960 Monoacylated MEL MML

References

2.7  106 3.6  106

Surface tension (mN/m) 28.4 25.3

Kitamoto et al. (1993b) Morita et al. (2009c)

2.9  106

28.6

Morita et al. (2009c)

4.5  106 3.7  106

28.2 25.6

Kitamoto et al. (1993b) Morita et al. (2009c)

6.0  106

25.1

Konishi et al. (2008)

4.0  106

24.2

Morita et al. (2008a)

4.7  106

30.7

Rodrigues et al. (2006)

6.4  106

29.8

Morita et al. (2009c)

3.6  104 2.6  106

33.8 24.2

Fukuoka et al. (2007c) Morita et al. (2009b)

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Fig. 8 Structure of Mannosylmannitol lipid (MML) n 6 12

including antitumor and differentiation-inducing activities against human leukemia cells, rat peochromocytoma cells, and mouse melanoma cells (Isoda et al. 1997a, b; Isoda and Nakahara 1997; Wakamatsu et al. 2001; Zhao et al. 1999, 2000, 2001). They can be used in the treatment of schizophrenia or diseases caused by dopamine metabolic dysfunction (Vertesy et al. 2002; Zhao et al. 2000) and microbial infections (Kitamoto et al. 1993b). MELs are used in the purification of lectins (Immunoglobulin G) (Kitamoto et al. 2000; Im et al. 2001, 2003; Imura et al. 2007b;) and in the preparation of ice-slurry, as an antiagglomeration agent (Kitamoto et al. 2001b, 2002). In addition, MEL-A, acetylated at C-4 and/or C-6, markedly increases the efficiency of gene transfection mediated by cationic liposomes (Inoh et al. 2001, 2004). Self-assembling properties (Imura et al. 2004, 2005, 2006, 2007a) of MELs are of interest because this behavior could be leveraged in gene transfection and drug delivery (Inoh et al. 2001, 2004; Ueno et al. 2007a, b). The subsequent sections describe their production conditions, biosynthetic pathway, their physicochemical properties, and their applications.

4 Genetic Regulation and Biosynthesis of MEL Although glycolipids production in fungi has been known for a long time, the genetic basis of their production and their regulation are largely unknown. However, the genes involved in the biosynthesis of MEL are analyzed only for U. maydis by Hewald et al. (2005, 2006). Production of ustilipids [Mannosylerythritol lipid (MEL)] and ustilagic acid (cellobiose lipid) in U. maydis occurs readily under conditions of nitrogen starvation and can reach large values (up to 23 g/L). The yield and ratio of both the classes of glycolipids depend on the available carbon source and can be shifted towards either of these BSs (Spoeckner et al. 1999). Hewald et al. (2005) cloned two genes (emt1 and cyp1) involved in the biosynthesis of

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glycolipid in U. maydis. The gene cyp1 is essential for the production of cellobioselipid. A putative glycosyltransferase, Emt1, is required for the production of MELs. The expression of this particular protein is enhanced by nitrogen starvation. This protein showed high level of sequence similarity with glycosyltransferases and mycosamine transferases, which are involved in the production of actinobacterial macrolide antibiotic. Mycosamine transferases transfer mycosamine to the macrolide chain. This mycosamine is a derivative of mannose (Aparicio et al. 2003). This is generated by the activity of several enzymes that modify the activated GDPmannose. Based on this analysis, Hewald et al. (2005) suggested that the U. maydis Emt1 protein may use GDP-mannose for the generation of the mannosylerythritol moiety of ustilipids. Emt1 directly transfers mannose to the C-4 atom of the mesoerythritol. This reaction has to be stereospecific, since only mannosyl-D-erythritol is generated. Hewald et al. (2006) identified the first biosynthesis gene cluster (Fig. 9) for a fungal extracellular glycolipid. The U. maydis MEL cluster specifies five proteins, one glycosyltransferase, three acyltransferases, and one export protein of the major facilitator family. All the enzymes are most probably involved in the biosynthesis and export of mannosylerythritol. By mutation analysis, they demonstrated that the putative acyltransferases, Mac1 and Mac2, are both essential for the biosynthesis of MEL. The acetyltransferase Mat1 catalyzes the acetylation of MELs in U. maydis. The first step in the biosynthetic pathway (Hewald et al. 2006) (Fig. 10) would be the generation of mannosylerythritol by mannosylation of erythritol, which is most probably catalyzed by the previously described glycosyltransferase Emt1 (Hewald et al. 2005). The intermediate mannosylerythritol has been isolated in significant amounts from MEL-producing cells (Boothroyd et al. 1956). This disaccharide is subsequently acylated with fatty acids of various lengths by the putative acyltransferases Mac1 and Mac2 at positions C-2 and C-3, respectively. This acylation reaction appears to be essential for secretion because deletion of either mac1 or mac2 abolished MEL production completely. However, the order of activity of these two enzymes on their substrates or which acyltransferase is responsible for acylation of C-2 and C-3 is unclear. In general, MELs secreted by U. maydis carry a short fatty acid (C2 to C8) at position C-2 and a medium or long fatty acid (C10 to C18) at C-3 (Kurz et al. 2003; Spoeckner et al. 1999). This implies that Mac1 and Mac2 differ not only in their regioselectivity but also in their preference for the length of the acyl-CoA cofactor. MEL derivatives with an acetyl group at C-2 were identified, which implied that at least one of the acyltransferases, presumably the one which catalyzed the transfer of the short-chain fatty acid, also accepted acetyl-CoA as a donor. 2.3x

6.9x

5.4 x

25.5 x

5.0 x

mat 1

mmf 1

mac 1

emt 1

mac 2

um03114

um03115

um03116

um03117

um10636

Fig. 9 Biosynthetic gene cluster (adapted from Hewald et al. 2006)

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Fig. 10 Biosynthetic pathway of MEL (adapted from Hewald et al. 2006). mat1, Acetyltransfer ase gene; mmf1, member of the major facilitator family; mac1 and mac2, putative acyltrans ferases; emt1, glycosyltransferase gene; Induction of transcription under conditions of nitrogen limitation is indicated above the open reading frames. emt1 and mac1 contain single intron. The MUMDB entry numbers are shown

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For the MEL-producing fungus, C. antarctica, it has been previously shown that medium-length fatty acids are derived from longer fatty acids by partial peroxisomal b-oxidation (Kitamoto et al. 1998). U. maydis contains both mitochondrial and peroxisomal proteins for the degradation of fatty acid. The final step of MEL biosynthesis is the acetylation of the fully acylated MELs. Acetylation is not essential for glycolipid secretion, since strains deficient of mat1 secrete large amounts of deacetylated MEL-D. Mat1 displays a relaxed regioselectivity and is able to transfer acetyl groups to both positions (C-4 and C-6 positions) of the mannosyl moiety. Mat1 overexpressed in E. coli can perform this reaction in vitro, which indicates that no further enzymes are involved in this reaction. The detailed fatty acid biosynthetic pathway of MEL in basidiomycetous yeasts, including the genus, Pseudozyma, was studied by Kitamoto et al. (1993a). They showed that the fatty alcohols or acids of chain length of Cn were used and the products formed were with the chain length of Cn 2, Cn 4, Cn 6, etc. (Kitamoto et al. 2002). They also concluded that the products were the b-oxidation intermediates of the substrates supplied. 2-Bromooctanoic acid (Kitamoto et al. 1998) (strong inhibitor of the fatty acid b-oxidation) inhibited the production of MEL, and the degree of inhibition increased with the increase in the chain length of the substrate supplied. The cerulenin (strong inhibitor to de novo fatty acid synthesis) did not affect the fatty acid composition of MEL. Once the fatty acid entered the b-oxidation pathway, it was completely oxidized to an acetyl CoA, and the intermediates did not leak out from this cycle (Tanaka and Fukui 1989). Two types of b-oxidation cycles are present in mammals, namely peroxisomal partial b-oxidation and mitochondrial complete b-oxidation. The former is responsible for the shortening of long and very long-chain fatty acids, which are then converted into acetyl CoA in the latter cycle (Wander et al. 2001; Kitamoto et al. 2002). The biosynthetic “chain shortening” pathway of MEL is different from the three generally known pathways in microorganisms, namely de novo synthesis, chain elongation, and intact incorporation (Kitamoto et al. 2002). The genetic information on P. antarctica T-34 was obtained by constructing a cDNA [expressed sequence tags (ESTs)] library from P. antarctica T-34 after their cultivation in the presence of soybean oil, which was followed by identifying the genes involved in the biosynthesis of MEL under different culture conditions using real-time reverse transcriptase-PCR (RT-PCR) (Morita et al. 2006a). This study generated 398 ESTs assembled into 146 contiguous sequences. Similarity search using BLAST showed that 21.4% of all contigs were orphans, while 78.6% showed similarity to sequences in the protein database. 60.3% of all contiguous sequences share significant identities to hypothetical protein of U. maydis, which is a smut fungus and produces BS. Real-time reverse transcriptase-PCR results indicated that mannosyltranferase and acyltransferase were involved in the biosynthesis of MEL in cells that were grown in soybean oil (Morita et al. 2006a).

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5 Evolutionary Relationship Among MEL Producers MEL-producing microorganisms are placed under Ustilaginomycetous anamorphic yeast (Ustilaginales) along with the smut fungi Ustilago sp., (U. maydis produces MEL as well as cellobiose lipids) based on the rDNA sequence similarity (Sugita et al. 2003). These are isolated from plants. P. rugulosa NBRC 10877 was discovered by Morita et al. as MEL producer based on the sequence similarity of internal transcribed spacer1 (ITS1), 5.8S rRNA gene, and internal transcribed spacer (ITS2) with respect to other already known MEL producers (P. antarctica, P. aphidis, U. maydis, and Kurtzmanomyces sp. I-11). This was later confirmed as a MEL producer with more than 97% similarity to the above mentioned high MEL producers (Morita et al. 2006b). U. maydis was also closely related to the above strains. It has 85 and 86% identity to P. antarctica and P. aphidis, respectively. These facts indicate that the genus, Pseudozyma, is the BS-producing microorganism and that the genes involved in the biosynthesis of MEL might be conserved among the strains (Morita et al. 2006b). Molecular phylogenetic analysis of different strains of MEL producers demonstrated the three clusters in the phylogenetic tree. The high-level MEL producers (including P. antarctica, P. aphidis, P. rugulosa, and P. parantarctica) and efficient MEL-A producers (includes the strains KM-65, KM-73, KM-90, KM-99, KM-103, KM-105, KM-122, KM-125, KM-131, and KM-159) are positioned very close to each other in the phylogenetic tree (Fig. 11), except the strain KM-99 (Konishi et al. 2007b). MEL-C producer P. Shanxiensis (DQ008956)

MEL-C producer P. hubeiensis (DQ008954)

P. flocculosa (AB089364)

P. fusiformata (AB089366)

P. prolifica (AM160639)

MEL-B producer P. tsukubaensis (AB089372)

P. thailandica (AB089354)

MEL-A producer P. parantarctica (AB089356)

P. antarctica (AY641557) P. rugulosa (AB089370) P. aphidis (AB089362)

Fig. 11 Phylogenetic tree (adapted from Fukuoka et al. 2007a)

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This particular group produces MEL with the fatty acid chain lengths varying from C8 to C10. MEL-B producers (P. tsukubaensis, KM-160) are clustered together and are placed distant from MEL-A producers. These MELs possess fatty acid chain lengths of C8, C12, and C14 acids. Similarly, MEL C producers (P. hubiensis) represent the third main cluster, which is distantly placed from the other two producers. They possess fatty acid chain length of C6, C12, and C16 (Konishi et al. 2007b). In general, MEL-C is more hydrophilic when compared to the other conventional type of MELs (Konishi et al. 2007b). MEL-C from P. shanxiensis, P. graminicola, P. siamensis, and Ustilago cynodontis possesses much shorter chain length (C2 or C4 at C-20 position). This is reflected in the phylogenetic tree and moreover they are positioned a little distant from conventional MEL C producer, P. hubiensis (Morita et al. 2008c). Similarly, P. graminicola is also placed a little distant from the abovementioned MEL C producer and it possesses the fatty acid chain length of C6, C8, and C14. Pseudozyma fusiformata, which produces ustilagic acid and MEL, is placed very close to U. maydis and it is distant from MEL- A and B producers. Pseudozyma prolifica, Pseudozyma flocculosa, and Pseudozyma thailandica, which do not produce MEL, are also placed distant from the MEL producers (Morita et al. 2007; Konishi et al. 2007b). All these facts suggest that there is a high similarity in the biosynthesis of polar groups and the genes involved in it, whereas there is large diversity in the biosynthesis of hydrophobic chain and the genes involved in it (Konishi et al. 2007b). It is also indicated that the three groups physiologically differ in the fatty acid metabolism as well as in the manner of acetylation and/or acylation into the hydrophilic part, i.e., 4-O-b-D-mannopyranosyl-meso-erythritol (Konishi et al. 2007b). Even across the strains, variations are observed, especially in the case of MEL-C producer. Based on the abovementioned facts, the genus Pseudozyma can be classified into the following four groups (Morita et al. 2007): 1. 2. 3. 4.

Large amount of MEL-A producers Both MELs and other BSs producers Mainly MEL-B and MEL-C producers Do not produce MEL

MEL production can also be considered as an important taxonomic index for the characterization of the genus, Pseudozyma yeast, together with the other biochemical characterization techniques including coenzyme Q pattern, analysis of polysaccharides present on the cell wall, and cellular fatty acids analysis (Morita et al. 2007).

6 Bioprocesses Used for MEL Production The production of MEL in shake flask is reported while very few attempts have been made to produce it in a fermentor. Seed cultures are prepared by inoculating cells grown on slants into test tubes containing the growth medium [4% glucose, 0.3% NaNO3, 0.03% MgSO4, 0.03% KH2PO4, 0.1% yeast extract (pH 6.0)] at 25 C

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on a reciprocal shaker (150 rpm) for 2 days (Fukuoka et al. 2007c; Kitamoto et al. 1990a, b; Rau et al. 2005a, b, c). Exactly 1 mL of the seed culture is transferred to a 300 mL Erlenmeyer flask containing 30 mL of the basal medium [4% olive oil, 0.3% NaNO3, 0.03% MgSO4, 0.03% KH2PO4, 0.1% yeast extract (pH 6.0)], which is then incubated on a rotary shaker (220 rpm) at 28 C for 7 days. Isolation of glycolipid is carried out by ethyl acetate. Then, the solvent layer is collected and evaporated (Fukuoka et al. 2007c; Kitamoto et al. 1990a, b; Rau et al. 2005a, b, c). The concentrated glycolipids are dissolved in chloroform and then purified by silica gel (Wako-gel C-200) column chromatography using a gradient elution technique with chloroform acetone (10:0 0:10, vol/vol) solvent mixtures (Fukuoka et al. 2007c; Rau et al. 2005a, b, c). Table 3 summarizes the different conditions reported for the production of MEL by Pseudozyma sp. Rau et al. (2005a, b, c) reported the production of MEL in a bioreactor. Solvent extraction, adsorption on different types of Amberlite XAD followed by solvent

Table 3 Effect of culture conditions on the composition of MEL by Pseudozyma sp. Type of MEL Yield (g/L) Conditions References MEL B7 30 n alkane or vegetable oils; batch Kawashima et al. culture (1983) Mixture of MEL 47 Soybean oil and resting cells; batch Kitamoto et al. A, B and C culture (1992) Mixture of MEL 40 Vegetable oils, yeast extract and Kitamoto et al. A, B, C and D batch culture (1990b) 30 Sunflower oil; batch culture Kitamoto et al. (1990b) Mixture of MEL 140 Octadecane ; batch culture Kitamoto et al. A, B and C (2001a) Mixture of MEL 165 Soybean oil and fed batch reactor Kitamoto et al. A, B, and C (2002) Rau et al. (2005b) Mixture of MEL 95 Glucose:soybean oil (1:1) and homologs intermittent addition of soybean fed batch culture MEL B 25 Soybean oil batch culture Kim et al. (2006) MEL C 15 Soybean oil batch culture Konishi et al. (2007b), Fukuoka et al. (2008) MEL C 76.3 Soybean oil and fed batch reactor Konishi et al. (2008) MEL A with 1.61, 1.97 Glucose, sucrose, batch culture Morita et al. (2009c) relatively high hydrophilicity Glucose, sucrose, batch culture Morita et al. (2009c) 6.2, 6.4 (12.8 MEL B with under relatively high hydrophilicity optimal conditions) 1.08, 1.94 Glucose, sucrose, batch culture Morita et al. (2009c) MEL C with relatively high hydrophilicity MML 18.2 Soybean oil and mannitol Morita et al. (2009b)

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extraction, and heating the culture suspension to 110 C for 10 min were the three downstream methods reported. The last method gave the highest yield (of 93 wt%) with a purity of 87 wt%, whereas the solvent extraction method gave only 79 wt% yield with 100 wt% purity.

6.1

Factors Affecting MEL Production

In this section, the various aspects of MEL production, including the influence of the type and quantity of the nutrients in the medium, environmental parameters, and the optimum time period for the better yield of MEL, will be presented.

6.1.1

Carbon Source

Almost all vegetable oils (except palm oil and coconut oil) serve as a good carbon source for the production of MEL by various Pseudozyma sp. Soybean oil, olive oil, and safflower oil are the best carbon sources. P. rugulosa NRBC 10877 and P. parantarctica JCM 11752 (Morita et al. 2008c) produced the highest amount of MEL with soybean oil when compared to other vegetable oils tested (safflower oil, soybean oil, palm oil, corn oil, olive oil, rapeseed oil, and coconut oil) (Morita et al. 2006b). Water-soluble carbon sources like glucose and sucrose simplify the separation steps when compared to the vegetable oils.

6.1.2

Nitrogen Source

The type of nitrogen source considerably affected MEL formation. Sodium nitrate (0.3%, wt/wt) was clearly the best nitrogen source while ammonium nitrate and ammonium sulfate were not. As previously reported (Kitamoto et al. 1990b; Rau et al. 2005a, b), the decrease of pH by the consumption of ammonium salts should lead to a decrease in MEL yield. Strain KM-59 and U. scitaminae showed increase in MEL production when concentration of yeast extract was increased up to 2.0 g/L and 5.0 g/L, respectively (Konishi et al. 2008; Morita et al. 2009a). Further increase led to slight decrease in the production of MEL.

6.1.3

Effects of Hydrophilic Precursor

In all the cases, the amount of MEL was significantly increased by the addition of the hydrophilic carbon source. The amount of MEL increased with an increase in the concentration of the carbohydrate added. The amount of MEL produced by P. rugulosa increased (70 90%) when erythritol was added (Morita et al. 2006b). The yield was 31.5 g/L on addition of 8% erythritol. Addition of glucose and

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mannose gave approximately 50% increase in the MEL yield by P. rugulosa (Morita et al. 2006b). Strain KM-59 showed increase in the production of MEL with increase in the concentration of glucose up to 60 g/L, and further increase showed no effect (Konishi et al. 2008). Similarly, addition of mannose and erythritol did not improve the production of MEL by KM-59. Smut fungus, U. scitaminea, demonstrated increase in MEL production with increase in the concentration of sucrose up to 150 g/L (Morita et al. 2009a).

6.1.4

Effect of Temperature

Almost all Pseudozyma sp., including high MEL producers, exhibited low MEL yield with increase in temperature, except P. parantarctica JCM11752. This microorganism showed the best growth at 36 C whereas the best yield was observed at 34 C (Morita et al. 2008c). Other organisms produced maximum MEL in the temperature range of 25 30 C (Morita et al. 2006b, 2009a).

6.1.5

Time Course of MEL Production

P. parantarctica JCM 11752 showed maximum production of MEL on the 28th day and no cell growth was observed after 7 days. This shows that the activity of the resting cells was maintained efficiently for 4 weeks. Similarly, U. scitaminea exhibited high production of MEL on the 21st day (Morita et al. 2009a). Fed batch fermentation with intermittent feeding of carbon source showed improvement in the production of MEL with P. rugulosa NRBC 10877 (Morita et al. 2006b) and strain KM-59 (Konishi et al. 2008). Addition of soybean oil, yeast extract, and glucose in the fed batch fermentation was helpful in maintaining the activity of resting cells for 16 days in the case of strain KM-59 and for 3 weeks in the case of P. rugulosa.

7 Phase Behavior of MEL Self-assembly (SA) can be defined as the spontaneous and reversible organization of molecular units into ordered structures by noncovalent interactions without the application of external force. Any amphiphilic molecule possesses this ability. Ionic and nonionic surfactants at high concentrations can self-assemble into 3D-ordered lyotropic liquid crystals including sponge, cubic, lamella, and hexagonal phases. Natural sugar-based surfactants exhibit the above mentioned behavior. They can self-assemble into specific lyotropic liquid crystalline phases, which are stabilized by hydrogen bonds between the sugar moieties. Chirality of the sugar also affects their lyotropic and thermotropic phase behaviors (Imura et al. 2007a; Kitamoto

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et al. 2000). The four classes of MEL with variation in their hydrophilicity show different self-assembling properties including liposomes, self-assembled monolayer (SAM), lamella phase, sponge phase, liquid lyotropic crystals, and bicontinuous cubic phase (Imura et al. 2004, 2005, 2006, 2007b; Kitamoto 2008; Worakitkanchanakul et al. 2008).

7.1

Formation of Thermodynamically Stable Vesicles and Coacervates by MEL

Self-assemblies formed by MEL-A and MEL-B are showed in Fig. 12. MEL-A spontaneously forms sponge phase at concentrations above 1 mM whereas MEL-B and MEL-C form giant unilamellar vesicles of diameter larger than 10 mm (Imura et al. 2004; Kitamoto et al. 2009). This is due to the slight decrease in the spontaneous curvature resulting from the absence of the 4-O-acetyl group. This induces a drastic morphological change in the self-assembled structure, from coacervates to vesicles. Giant vesicles can be used as cell models for studying many cellular processes including endocytosis, exocytosis, cell fusion, and

Fig. 12 Different self assemblies formed by Mannosylerythritol lipid A and B (“Reprinted from Kitamoto et al. (2009). Copyright with permission from Elsevier”)

Mannosylerythritol Lipids: Microbial Production and Their Applications Fig. 13 Mechanism of formation of thermodynamically stable vesicles by MEL A/DLPC mixture. L3, MEL sponge phase; La1, thermodynamically stable vesicle; La, Large multilamellar vesicle ( filled circle MEL A negatively curved lipid; open circle DPLC zero curved lipid)

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L3

La1

La

transport phenomena (Dobereiner 2000). The sponge phases can also be considered as coacervates, L3 phase, “plumber’s nightmare,” or “blue I phase” (Imura et al. 2004). MELs form thermodynamically stable vesicle (La1) (Fig. 13) from the sponge phase (L3) when they are mixed with L-a-dilauroylphosphatidyl choline (DLPC) (Imura et al. 2005). Since they are negatively curved lipids, mixing DLPC (phospholipids) with this sponge phase induces the fusion of both the lipids and favors their distribution in the inner monolayer of the vesicle. This causes the asymmetric distribution of the two lipids leading to the formation of thermodynamically stable vesicle (Fig. 13). Smallest sized vesicles (633.3 nm) are obtained at a DLPC mole fraction of 0.3 and are stable at 25 C for more than 3 months. Such vesicles formed from the natural glycolipids are preferred for drug delivery and gene transfer due to their biocompatibility when compared to their synthetic counterparts (Imura et al. 2005).

7.2

Multilamellar Vesicles and Large Unilamellar Vesicles

Self-assembling properties of MEL-A and -B can be studied with fluorescenceprobe spectroscopy, dynamic light scattering (DLS) spectroscopy, freeze-fracture transmission electron microscopy (FF-TEM), and synchrotron small/wide-angle X-ray scattering (SAXS/WAXS) spectroscopy. MELs self-assemble into large unilamellar vesicles (LUV) of diameter larger than 160 nm just above their critical-aggregation concentration (CAC), which are 4.0  10 6 and 6.0  10 6 M for MEL-A and MEL-B, respectively (Imura et al. 2006). Above a CAC (II) value of

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2.0  10 5 M, MEL-A is found to drastically change into a sponge-like structure (L3) which is composed of a network of randomly connected bilayers with a waterchannel diameter of 100 nm. This diameter is relatively large when compared to those obtained with synthetic surfactant multicomponent systems (24 nm), namely cetylpyridinium/hexanol/dextrose/brine systems. MEL-B, which has a hydroxyl group at the C-4 position instead of an acetyl group on mannose, gives only one CAC. The self-assembled structure of MEL-B seems to gradually move from LUV to multilamellar vesicles (MLV) with a lattice constant of 4.4 nm (Imura et al. 2006). The bilayer thickness is 3.2 nm for both MEL-A and MEL-B.

7.3

Lyotropic-Liquid-Crystalline Phases of MEL

Water penetration scans are used to study the various liquid lyotropic crystalline phases formed by different MELs. At high concentrations, the formation of an inverted hexagonal phase (H2) for MEL-A and a lamella phase (La) for MEL-B were observed under a polarized optical microscope. These results indicate that the difference in the spontaneous curvature between MEL-A and MEL-B molecules is due to an extra acetyl group present in the former. This difference decides the direction of their self-assembly. The microorganisms are able to engineer distinct self-assembled structures by varying the substitutions on the polar head group (Imura et al. 2006). MEL-A is able to self-assemble into a variety of distinctive lyotropic liquid crystals including L3, bicontinuous cubic (V2), and lamellar (La) phases (Imura et al. 2006). The lattice constants are estimated to be 11.39 and 3.58 nm for V2 and La, respectively. Differential scanning calorimetry (DSC) measurements revealed that the phase transition enthalpies from these Lyotropic liquid crystals (LCs) to the fluid isotropic (FI) phase are in the range of 0.22 0.44 kJ/mol. L3 region of MEL-A is spread considerably over a wide temperature range (20 65 C) when compared to that of other surfactants (Imura et al. 2007a). This is probably due to its unique structure, which is molecularly engineered by the microorganisms. MEL-B from P. tsukubaensis (yield 20 g/L) has 1-O-b-(20 ,30 -di-O-alka(e)noyl-60 -O-acetyl-D-mannopyranosyl)-Derythritol (Fukuoka et al. 2008), which is a diastereomer of the conventional MEL-B. This selfassembles into a lamellar (La) phase over a wide range of concentrations and temperatures, whereas MEL-A (diacetyl) forms L3, V2, and La phases. It is observed that the difference in the number of acetyl groups and the configuration of erythritol moiety play an important role in the aqueous-phase behavior of MEL/ water mixture. The interplanar distance (d) of La phase of MEL-B is estimated to be 4.7 nm at low concentrations (60 wt%), and this d-spacing decreases with increase in concentration (~3.1 nm at >60%). This phase is found to be stable up to 95 C and below a concentration of 85% (wt). The melting temperature of the liquid crystalline phase dramatically decreases with increase in MEL-B concentration (above 85 wt%). The MEL-B is able to form vesicle of dimension 1 5 mm (Worakitkanchanakula et al. 2008). MEL-C from P. siamensis CBS 9960 forms

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liquid crystal phases such as hexagonal (H2) and La phases at a wide range of concentrations (Morita et al. 2008b). The phase behavior of ternary MEL-A/water/n-decane (Worakitkanchanakul et al. 2009; Kitamoto et al. 2009) (Fig. 14a) and MEL-B/water/n-decane (Fig. 14b) systems were reported by Worakikanchanakul et al. When n-decane is used as an oil phase, di-acetylated MEL-A formed single-phase, namely water-inoil (W/O) microemulsion in a remarkably large region. This includes sponge (L3), reverse bicontinuous cubic (V2), and lamellar (La) phases. Whereas monoacetylated MEL-B obtained from P. tsukubaensis (Imura et al. 2007a) gave a singlephase, bicontinuous microemulsion and showed a triangular phase diagram dominated by the La phase. This difference can be attributed to the opposite configuration of the erythritol moiety in MEL-B and it suggests that MEL-A has negative spontaneous curvature whereas MEL-B has zero spontaneous curvature. Oil-in-liquid crystal (O/LC) emulsion is easily prepared in the biphasic Laþoil region of the MEL-B/water/n-decane system. The obtained gel-like emulsion is stable for a month. These information highlight the importance of further study on structure function relationship of MELs and other BSs.

7.4

Self-Assembled Monolayer Structures of MEL

MEL-A has been used as a model to understand the role of molecular interaction between glycolipids and proteins in biological recognition events including

Fig. 14 Phase diagrams of ternary complexes of MEL A and B. (a). Phase diagram of the MEL A/water/n decane system at 25 C: L3, sponge phase; V2, reverse bicontinuous cubic phase; La, lamellar phase; O/W, O/W typemicroemulsion; W/O, W/O type microemulsion; D, bicontinuous microemulsion; W, excess water; O, excess oil. (b) Phase diagram of the MEL B/water/n decane system at 25 C: La, lamellar phase; O/W, O/W type microemulsion; W/O, W/O type microemul sion; D, bicontinuous microemulsion; W, excess water; O, excess oil. (“Reprinted from Kitamoto et al. (2009). Copyright with permission from Elsevier”)

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cell adhesion, signal transduction, and immune function. MEL exhibits highly ordered self-assemblies and affinity towards glycoproteins. Self-assembled monolayers (SAMs) of MEL on the alkanethiol SAMs using MEL-A bilayer membrane structures are called supported “Hybrid Bilayer Membranes” (HBMs) (Fig. 15). The interaction between the HBMs and two different immunoglobulins, namely HIgG and HIgM, was studied by using surface plasmon resonance (SPR) and atomic force microscopy (AFM). The affinity constants (Ka) between SAM and HIgG and HIgM are 9.4  106 and 5.4  106/M, respectively. The binding affinity between the SAM of MEL A and HIgG is 25 times higher than that observed with Staphylococus aureus protein A, which is very commonly used for the purification of HIgG. Also, it is six times higher than that observed with MEL-A and poly (2-hydroxyethyl methacrylate) (PHEMA) (Imura et al. 2007b). MEL-A SAMs on alkanethiolates are well aligned along the surface when compared to those on polymer surfaces prepared by the solvent evaporation method. Although the individual carbohydrate and protein interaction is relatively weak (Ka ~ 103 to 104/M), the high binding affinity observed here can be attributed to the multivalent effect of the SAM of MEL (Imura et al. 2006, 2007b; Kitamoto 2008). Table 4 summarizes the different classes of MELs and their corresponding self-assembling properties.

Lectins and immunoglobulins

Multivalent interactions

MEL- A self assembled monolayers

Octadecanethiol C18 SH

SH

SH

SH

SH

SH

Gold coated solid support

Fig. 15 Schematic representation of hybrid bilayer membrane and molecular interactions

Mannosylerythritol Lipids: Microbial Production and Their Applications Table 4 Self assembling properties of various MEL homologs Self assembling Type of MEL Size of Critical structures aggregation the vesicles concentration (M) (mm) MEL A 1 20 4.0  106 Spherical droplet, large unilamellar vesicles (LUV), sponge (L3), bicontinuous cubic (V2), coacervate and lamellar Conventional MEL B 1 20 6.0  106 Giant vesicles, multi lamellar vesicles (MLV) MEL B (new 1 5 Lamellar (La) phase diastereomer over a wide P. tsukubaensis) concentration and temperature ranges, Vesicle MEL C Giant vesicles, myelins and lamellar (La) MEL C from P. siamensis CBS 9960 MML

Hexagonal (H) and lamellar (L3) Myelins and lamellar (La)

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References

Imura et al. (2004, 2005, 2006, 2007a), Kitamoto et al. (2009)

Imura et al. (2004, 2006), Kitamoto et al. (2009) Vertesy et al. (2002), Kitamoto et al. (2009)

Konishi et al. (2008), Kitamoto et al. (2009) Rodrigues et al. (2006), Kitamoto et al. (2009) Morita et al. (2009b)

8 Applications MELs have antimicrobial activities and other biological activities, which are described below (Hamme et al. 2006; Rosenberg and Ron 1999).

8.1

Antimicrobial Activity of MEL

Both MEL-A and MEL-B show strong activity against gram-positive bacteria, weak activity against gram-negative bacteria, and no activity against fungi. The minimum inhibitory concentrations against gram-positive bacteria are significantly lower than those exhibited by sucrose and sorbital monoesters of fatty acids. Other MELs accumulated in the cell of S. melanogramma also show antibacterial and antifungal activities (Deml et al. 1980). Matsumura et al. reported that mannopyranosyl groups are the most effective amongst the n-alkyl glycosides tested against microorganisms. It may be possible to enhance the antimicrobial activity of the present MELs further by the chemical modification of the sugar moiety (Kitamoto et al. 1993b).

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MEL Induces Cell Differentiation and Apoptosis

MEL-A and -B have excellent growth inhibition and differentiation-inducing activities against human leukemia cells including myelogenous leukemia cell K562 [26], promyelocytic leukemia cell HL60 [25, 26], and the human basophilic leukemia cell line KU812 (Isoda et al. 1997a; Kitamoto et al. 2002). Both inhibit the growth of HL60 cells and induce changes to their morphology at concentrations between 5 and 10 mM. They (at 10 mg/L) also induce granulocytic differentiation (Isoda and Nakahara 1997; Kitamoto et al. 2002) and inhibit the activity of phospholipidand Ca2+-dependent protein kinase C in HL 60 cells. MEL-A inhibits the serine/ threonine phosphorylation of a 30 kDa protein in HL-60 cells and the tyrosine phosphorylation of 55-, 65-, 95-, 135-kDa proteins in K562 cells (Kitamoto et al. 2002). Therefore, MEL appears to directly affect the intracellular signal transduction through phosphate cascade systems. Both induce significantly the neurite outgrowth of rat pheochromocytoma PC-12 cells and partial cellular differentiation (Wakamatsu et al. 2001; Kitamoto et al. 2002). MEL-A has been recently demonstrated to inhibit the growth of mouse melanoma B 16 cells in a dose-dependent manner. At 10 mM concentration, it causes the condensation of chromatin, fragmentation of DNA, and the arrest of sub-G1. This indicates that the cells are undergoing apoptosis. MEL also induces tyrosinase activity and enhances the production of melanin (Zhao et al. 1999). This trigger of mouse melanoma B 16 cells may be through a signaling pathway that involves protein kinase Ca (PKCa) (Zhao et al. 2001).

8.3

Purification of Glycoproteins

MEL-A, -B, and -C exhibit high binding affinity to human immunoglobulins (Im et al. 2001, 2003; Kitamoto et al. 2000; Kitamoto 2008). They also show binding affinity towards Ig M and other glycoproteins. SAM of MEL-A shows six times higher affinity than the immobilized MEL-A in PHEMA (Imura et al. 2007b). These results indicate their potential application as a ligand for the purification of lectins.

8.4

Vehicles for Gene Delivery

MELs can be used as a vehicle for gene (Fig. 14) and in drug delivery due to their ability to form thermodynamically stable vesicles with the ability to fuse with the membrane (Inoh et al. 2001, 2004; Kitamoto 2008; Ueno et al. 2007a, b). Among various BSs, MEL-A provides the highest sufficiency (Inoh et al. 2004). It remarkably accelerates the adhesion of positively charged liposome DNA complex to the

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cell membrane and incorporates this complex into the cell (Inoh et al. 2001; Kitamoto 2008).

8.5

Inhibition of Ice Agglomeration

Ice-slurry systems are finding wide applications as environmental friendly cold thermal storage units, especially as air conditioners (Inaba 2000; Kitamoto et al. 2002). In these units, there is a possibility for the ice particles to agglomerate or grow together and block the pipeline, causing superfluous power loads thereby leading to loss of efficiency. MEL at low concentration (10 mg/L) gets adsorbed on the ice surface and suppresses this agglomeration process. Their effective performance has also been tested successfully in a large-scale model (300 L) (Inaba 2000; Kitamoto et al. 2000, 2001b).

8.6

Cosmetic Applications of MEL

Skin care property of MEL-A was estimated by applying it at different doses to the SDS-treated cultured human skin model, and the cell viability was determined by the MTT method (Morita et al. 2009d). The recovery effect of MEL-A on the cells from the SDS-induced damage was observed in a dose-dependent manner. Compared to the control, the viability of the cells treated with MEL-A solutions of 0.1, 1, 5, 10 wt% was 18.4, 36.2, 73.2, 91.3%, respectively. Olive oil alone gave little effect on the cell viability after the SDS treatment. In the case of ceramides, the cells treated with 0.1 wt% solutions showed little effect on the viability, while the cells with 1 wt% solution completely recovered from the SDS-induced damage. These results suggested that MEL-A has ceramide-like skin care property. MEL A has structural similarity to ceramides and also forms liquid crystals (Morita et al. 2009d; Kitamoto et al. 2009). These properties facilitate them to penetrate into the intercellular spaces easily. Hence, MEL-A has a great potential for a novel skin care material with potential moisturizing activity.

9 Perspectives The past 60 years of research on MEL has proved that it is the most promising microbial extracellular glycolipid ever known because of its high yield, excellent surface activity, diverse biochemical functions, biocompatibility, and its wide range of applications. However, the commercial viability of MEL depends upon its production cost, which has to be significantly lowered with the help of recent

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developments in bioprocess technology together with the advanced separation techniques. Hence, the following research topics need to be explored: l

l

l l

l

l

l

l

l

l l l

l

10

Reduce the production cost by using cheaper raw material, optimize the production conditions, and downstream processes Use of water-soluble substrates like sucrose and glucose for their production to make the downstream process easier Engineer the overproducing mutant and recombinant strains Determine the role of cell wall components (of Candida sp. Torulopsis sp. and Pseudozyma sp.) and understand how they tolerate high levels of MEL. This may provide new approaches to increase the yield as well as this knowledge could be extrapolated for other classes of BSs Relate the effect of hydrophilic precursors to the change in the physicochemical properties of the corresponding MEL and understand the specificity of the enzyme that is involved in the biosynthesis of MEL Identify the enzymes involved in their biosynthesis and regulate or augment the production of a single type of MEL with higher yield Develop structure-activity relationship to tailor the compounds with the required features. This can be done by making derivatives (by chemical or enzymatic synthesis and modifications) and analyzing their corresponding activity performance Investigate the various factors that determine the self-assembling properties to enhance their scope of applications in the biomedical field MEL can interact with glycoproteins (lectins); hence use them as model systems to study the mechanism of different biochemical functions Application of MEL as cellular models, vehicle for drug, and gene delivery Establish mechanisms of antimicrobial and antitumor activities of MEL Similarity studies on the enzymes involved in the synthesis of different classes of glycolipid (rhamnolipids, sophorolipids, cellobiose lipids, MEL, etc.) may give insight into the specificity of the active site for the synthesis of particular kind of glycolipid Understand the role of MEL production in microbial physiology might unravel the reason behind their predominant association with plants

Conclusions

Mannosylerythritol lipid, 2,3-di-O-alka(e)noyl-b-D-mannopyranosyl-(1!4)-Omeso-erythritol, partially acetylated at C-4 and/or C-6 position, is produced by Ustilago sp., Kurtzmanomycis sp. I-11, Schizonella sp., and Pseudozyma sp. The type of MEL produced varies with the strains and the species of Pseudozyma. Hence, its production can be used as an important taxonomic index to identify Pseudozyma yeast. This is considered as a most promising BS because of its high yield, excellent surface activity, diverse biochemical functions, and

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biocompatibility. MELs exhibit interesting self-assembling properties, which enhance their efficiency in gene transfection. Recent research on the ability of MEL-A to form water-in-oil microemulsion without the use of a cosurfactant seems to add an impetus to the development of the microemulsion technology, which is limited due to the fact that they require cosurfactants, salts, or alcohols for stabilization. MEL induces differentiation of several carcinoma cell lines. They exhibit antimicrobial and antitumor activities. The high binding affinity of the monolayer of MEL-A to the immunoglobulins helps in the purification of lectins. However, their large-scale production and recovery processes have to be optimized in order to compete economically with the chemical surfactants. Even though enormous information is available on the growing number of microorganisms that produce MEL with unique physicochemical properties, very less knowledge is available on the enzymes involved in each step of its synthesis. Its regulation, structure function relationship, and mechanism of its biochemical functions such as differentiation on carcinoma cell lines are poorly understood and needs major research thrust.

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Haskins RH, Thorn JA, Boothroyd B (1955) Biochemistry of the ustilagenales. XI. Metabolic products of Ustilago zeae in submerged culture. Can J Microbiol 1:749 756 Hewald S, Josephs K, Bolker M (2005) Genetic analysis of Biosurfactant production in Ustilago maydis. Appl Environ Microbiol 71:3033 3040 Hewald S, Linne U, Schere M, Marahiel MA, Kamper J, Bolker M (2006) Identification of a gene cluster for biosynthesis of mannosylerythritol lipids in the basidiomycetous fungus Ustilago maydis. Appl Environ Microbiol 72:5469 5477 Im JH, Nakane T, Yanagishita H, Ikegami T, Kitamoto D (2001) Mannosylerythritol lipid, a yeast extracellular glycolipid, shows high binding affinity towards human immunoglobulin G. BMC Biotechnol 1:5 Im JH, Yanagishita H, Ikegami T, Takeyama Y, Idemoto Y, Koura N, Kitamoto D (2003) Mannosylerythritol lipids, yeast glycolipid biosurfactants, are potential affinity ligand materi als for human immunoglobulin G. J Biomed Mater Res 65A:379 385 Imura T, Yanagishita H, Kitamoto D (2004) Coacervate formation from natural glycolipid: one acetyl group on the headgroup triggers coacervate to vesicle transition. J Am Chem Soc 126:10804 10805 Imura T, Yanagishita H, Ohira J, Sakai H, Abeb M, Kitamoto D (2005) Thermodynamically stable vesicle formation from glycolipid biosurfactant sponge phase. Colloids Surf B Biointerfaces 43:115 121 Imura T, Ohta N, Inoue K, Yagi N, Negishi H, Yanagishita H, Kitamoto D (2006) Naturally engineered glycolipid biosurfactants leading to distinctive self assembled structures. Chem Eur J 12:2434 2440 Imura T, Hikosaka Y, Worakitkanchanakul W, Sakai H, Abe M, Konishi M, Minamikawa H, Kitamoto D (2007a) Aqueous phase behavior of natural glycolipid biosurfactant mannosyler ythritol lipid A: sponge, cubic, and lamellar phases. Langmuir 23:1659 1663 Imura T, Ito S, Azumi R, Yanagishita H, Sakai H, Abe M, Kitamoto D (2007b) Monolayers assembled from a glycolipid biosurfactant from Pseudozyma (Candida) antarctica serve as a high affinity ligand system for immunoglobulin G and M. Biotechnol Lett 29:865 870 Inaba H (2000) New challenge in advanced thermal energy transportation using functionally thermal fluids. Int J Therm Sci 39:991 1003 Inoh Y, Kitamoto D, Hirashima N, Nakanishi M (2001) Biosurfactants of MEL A increase gene transfection mediated by cationic liposomes. Biochem Biophys Res Commun 289:57 61 Inoh Y, Kitamoto D, Hirashima N, Nakanishi M (2004) Biosurfactant MEL A dramatically increases gene transfection via membrane fusion. J Control Release 94:423 431 Isoda H, Nakahara T (1997) Mannosylerythritol lipid induces granulocytic differentiation and inhibits the tyrosine phosphorylation of human myelogenous leukemia cell line K562. Cyto technology 25:191 195 Isoda H, Kitamoto D, Shinmoto H, Matsumura M, Nakahara T (1997a) Microbial extracellular glycolipid induction of differentiation and inhibition of the protein kinase C activity of human promyelocytic leukemia cell line HL60. Biosci Biotechnol Biochem 61:609 614 Isoda H, Shinmoto H, Kitamoto D, Matsumura M, Nakahara T (1997b) Differentiation of human promyelocytic leukemia cell line HL60 by microbial extracellular glycolipids. Lipids 32:263 271 Kakugawa K, Tamai M, Imamura K, Miyamoto K, Miyoshi S, Morinaga Y (2002) Isolation of yeast Kurtzmanomyces sp. I 11, novel producer of mannosylerythritol lipid. Biosci Biotechnol Biochem 66:188 191 Kawashima H, Nakahara T, Oogaki M, Tabuchi T (1983) Extracellular production of a manno sylerythritol lipid by a mutant of Candida sp. from n alkanes and triacylglycerols. J Ferment Technol 61:143 149 Kearns DB, Losick R (2003) Swarming motility in undomesticated Bacillus subtilis. Mol Micro biol 49:581 590

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Kim H S, Yoon BD, Choung DH, Oh HM, Katsuragi T, Tani Y (1999) Characterization of a biosurfactant, mannosylerythritol lipid produced from Candida sp SY16. Appl Microbiol Biotechnol 52:713 721 Kim H S, Jeon J W, Kim B H, Ahn C Y, Oh H M, Yoon B D (2006) Extracellular production of a glycolipid biosurfactant, mannosylerythritol lipid, by Candida sp. SY16 using fed batch fermentation. Appl Microbiol Biotechnol 70:391 396 Kitamoto D, Akiba S, Hioki C, Tabuchi T (1990a) Extracellular accumulation of mannosylery thritol lipids by a strain of Candida antarctica. Agric Biol Chem 54:31 36 Kitamoto D, Haneishi K, Nakahara T, Tabuchi T (1990b) Production of mannosylerythritol lipids by Candida antarctica from vegetable oils. Agric Biol Chem 54:37 40 Kitamoto D, Fuzishiro T, Yanagishita H, Nakane T, Nakahara T (1992) Production of mannosy lerythritol lipids as biosurfactants by resting cells of Candida antarctica. Biotechnol Lett 14:305 310 Kitamoto D, Nemoto T, Yanagishita H, Nakane T, Kitamoto H, Nakahara T (1993a) Fatty acid metabolism of mannosylerythritol lipids as biosurfactants produced by Candida antarctica. J Jpn Oil Chem Soc 42:346 358 Kitamoto D, Yanagishita H, Shinbo T, Nakane T, Kamisawa C, Nakahara T (1993b) Surface active properties and antimicrobial activities of mannosylerythritol lipids as biosurfactants produced by Candida antarctica. J Biotechnol 29:91 96 Kitamoto D, Yanagishita H, Hayara K, Kitamoto HK (1998) Contribution of a chain shortening pathway to the biosynthesis of the fatty acids of mannosyierythritol lipid (biosurfactant) in the yeast Candida antarctica: effect of b oxidation inhibitors on biosurfactant synthesis. Biotech nol Lett 20:813 818 Kitamoto D, Ghosh SGO, Nakatani Y (2000) Formation of giant vesicle from diacylmannosyler ythritols and their binding to concanavalin A. Chem Commun 2000:861 862 Kitamoto D, Ikegami T, Suzuki GT, Sasaki A, Takeyama Y, Idemoto Y, Koura N, Yanagishita H (2001a) Microbial conversion of n alkanes into glycolipid biosurfactants, mannosylerythritol lipids, by Pseudozyma (Candida antarctica). Biotechnol Lett 23:1709 1714 Kitamoto D, Yanagishita H, Endo A, Nakaiwa M, Nakane M, Akiya T (2001b) Remarkable antiagglomeration effect of a yeast biosurfactant, diacylmannosylerythritol, on ice water slurry for cold thermal storage. Biotechnol Prog 17:362 365 Kitamoto D, Isoda H, Nakahara T (2002) Functions and potential applications of glycolipid biosurfactants from energy saving materials to gene delivery carriers. J Biosci Bioeng 94:187 201 Kitamoto D (2008) Naturally engineered glycolipid biosurfactants leading to distinctive self assembling properties. Yakugaku Zasshi 128:695 706 Kitamoto D, Morita T, Fukuoka T, Konishi M, Imura T (2009) Self assembling properties of glycolipid biosurfactants and their potential applications. Curr Opin Colloid Interface Sci 14:315 328 Konishi M, Imura T, Fukuoka T, Morita T, Kitamoto D (2007a) A yeast glycolipid biosurfactant, mannosylerythritol lipid, shows high binding affinity towards lectins on a self assembled monolayer system. Biotechnol Lett 29:473 480 Konishi M, Morita T, Fukuoka T, Imura T, Kakugawa K, Kitamoto D (2007b) Production of different types of mannosylerythritol lipids as biosurfactants by the newly isolated yeast strains belonging to the genus Pseudozyma. Appl Microbiol Biotechnol 75:521 531 Konishi M, Morita T, Fukuoka T, Imura T, Kakugawa K, Kitamoto D (2008) Efficient production of mannosylerythritol lipids with high hydrophilicity by Pseudozyma hubeiensis KM 59. Appl Microbiol Biotechnol 78:37 46 Kurz M, Eder C, Isert D, Li Z, Paulus EF, Schiell M, Toti L, Vertesy L, Wink J, Seibert G (2003) Ustilipids, acylated b D mannopyranosyl D erythritols from Ustilago maydis and Geotrichum candidum. J Antibiot (Tokyo) 56:91 101 Lin SC (1996) Biosurfactant: recent advances. J Chem Technol Biotechnol 63:109 120

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Morita T, Konishi M, Fukuoka T, Imura T, Kitamoto D (2006a) Analysis of expressed sequence tags from the anamorphic basidiomycetous yeast, Pseudozyma antarctica, which produces glycolipid biosurfactants, mannosylerythritol lipids. Yeast 23:661 671 Morita T, Konishi M, Fukuoka T, Imura T, Kitamoto D (2006b) Discovery of Pseudozyma rugulosa NBRC 10877 as a novel producer of the glycolipid biosurfactants, mannosylerythritol lipids, based on rDNA sequence. Appl Microbiol Biotechnol 73:305 313 Morita T, Konishi M, Fukuoka T, Imura T, Kitamoto HK, Kitamoto D (2007) Characterization of the genus Pseudozyma by the formation of glycolipid biosurfactants, mannosylerythritol lipids. FEMS Yeast Res 7:286 292 Morita T, Konishi M, Fukuoka T, Imura T, Yamamoto S, Kitagawa M, Sogabe A, Kitamoto D (2008a) Identification of Pseudozyma graminicola CBS 10092 as a producer of glycolipid biosurfactants, mannosylerythritol lipids. J Oleo Sci 57:123 131 Morita T, Konishi M, Fukuoka T, Imura T, Kitamoto D (2008b) Production of glycolipid biosurfactants, mannosylerythritol lipids, by Pseudozyma siamensis CBS 9960 and their interfacial properties. J Biosci Bioeng 105:493 502 Morita T, Konishi M, Fukuoka T, Imura T, Kitamoto D (2008c) Identification of Ustilago cynodontis as a new producers of glycolipid biosurfactants, mannosylerythritol lipids, based on ribosomal DNA sequence. J Oleo Sci 57:549 556 Morita T, Ishibashi Y, Fukuoka T, Imura T, Sakai H, Abe M, Kitamoto D (2009a) Production of glycolipid biosurfactants, mannosylerythritol lipids, by smut fungus, Ustilago scitaminea NBRC 32730. Biosci Biotechnol Biochem 73:788 792 Morita T, FukuokaT KM, Imura T, Yamamoto S, Kitagawa M, Sogabe A, Kitamoto D (2009b) Production of a novel glycolipid biosurfactant, mannosylmannitol lipid, by Pseudozyma parantarctica and its interfacial properties. Appl Microbiol Biotechnol 83:1017 1025 Morita T, Ishibashi Y, Fukuoka T, Imura T, Sakai H, Abe M, Kitamoto D (2009c) Production of glycolipid biosurfactants, mannosylerythritol lipids, using sucrose by fungal and yeast strains, and their interfacial properties. Biosci Biotechnol Biochem 73:2352 2355 Morita T, Kitagawa M, Suzuki M, Yamamoto S, Sogabe A, Yanagidani S, Imura T, Fukuoka T, Kitamoto D (2009d) A yeast glycolipid biosurfactant, mannosylerythritol lipid, shows poten tial moisturizing activity toward cultured human skin cells: the recovery effect of MEL A on the SDS damaged human skin cells. J Oleo Sci 58:639 642 Mulligan CN (2004) Environmental applications for biosurfactants. Environ Pollut 133:183 198 Neu TR (1996) Significance of bacterial surface active compounds in interaction of bacteria with interfaces. Microbiol Rev 60:151 166 Rau U, Nguyen LA, Schulz S, Wray V, Nimtz M, Roeper H, Koch H, Lang S (2005a) Formation and analysis of mannosylerythritol lipids secreted by Pseudozyma aphidis. Appl Microbiol Biotechnol 66:551 5593 Rau U, Nguyen LA, Roeper H, Koch H, Lang S (2005b) Fed batch bioreactor production of mannosylerythritol lipids secreted by Pseudozyma aphidis. Appl Microbiol Biotechnol 68:607 613 Rau U, Nguyen LA, Roeper H, Koch H, Lang S (2005c) Downstream processing of mannosyler ythritol lipids produced by Pseudozyma aphidis. Eur J Lipid Sci Technol 107:373 380 Rodrigues L, Banat IM, Teixeira J, Oliveira R (2006) Biosurfactants: potential applications in medicine. J Antimicrob Chemother 57:609 618 Ron EZ, Rosenberg E (2001) Natural roles of biosurfactants. Environ Microbiol 3:229 236 Rosenberg E, Ron EZ (1999) High and low molecular mass microbial surfactants. Appl Micro biol Biotechnol 52:154 162 Shibahara M, Zhao X, Wakamatsu Y, Nomura N, Nakahara T, Jin C, Nagaso H, Murata T, Yokoyama KK (2000) Mannosylerythritol lipid increases levels of galactoceramide in and neurite outgrowth from PC12 pheochromocytoma cells. Cytotechnology 33:247 251 Spoeckner S, Wray V, Nimtz M, Lang S (1999) Glycolipids of the smut fungus Ustilago maydis from cultivation on renewable resources. Appl Microbiol Biotechnol 51:33 39

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Sugita T, Takashima M, Poonwan N, Mekha N, Malaithao K, Thungmuthawawat B, Pransarn S, Luangsook P, Kudo T (2003) The first isolation of ustilaginomycetous anamorphic yeasts, Pseudozyma species, from patients’ blood and a description of two new species: P. parantarctica and P. thailandica. Microbiol Immunol 47:183 190 Tanaka A, Fukui S (1989) Metabolism of n alkane. In: Rose AH, Harrison JS (ed) The yeasts, metabolism and physiology of yeasts, Vol 3. Academic Press, London, New York, pp 261 287 Ueno Y, Hirashima N, Inoh Y, Furuno T, Nakanishi M (2007a) Characterization of biosurfactant containing liposomes and their efficiency for gene transfection. Biol Pharm Bull 30:169 1723 Ueno Y, Inoh Y, Furuno T, Hirashima N, Kitamoto D, Nakanishi M (2007b) NBD conjugated biosurfactant (MEL A) shows a new pathway for transfection. J Control Release 123:247 253 Vejux A, Kahn E, Dumas D, Besse´de G, Me´ne´trier F, Athias A, Riedinger JM, Frouin F, Stoltz JF, Ogier Denis E, Todd Pokropek A, Lizard G (2005) 7 Ketocholesterol favors lipid accumula tion and colocalizes with Nile red positive cytoplasmic structures formed during 7 ketocho lesterol induced apoptosis: analysis by flow cytometry, FRET biphoton spectral imaging microscopy, and subcellular fractionation. Cytom A 64A:87 100 Vertesy L, Kurz M, Wink J, Noelken G (2002) Ustilipides, method for the production and the use thereof. US Patent 6,472,158 Wakamatsu Y, Zhao X, Jin C, Day N, Shibahara M, Nomura N, Nakahara T, Murata T, Yokoyama KK (2001) Mannosylerythritol lipid induces characteristics of neuronal differentiation in PC12 cells through an ERK related signal cascade. Eur J Biochem 268:374 383 Wander RJA, Vreken P, Ferdiandusse S, Jansen GA, Waterham HR, van Roermunde CWT, Grunsven EGV (2001) Peroxisomal fatty acid a and b oxidation in humans: enzymology, peroxisomal metabolite transporters and peroxisomal diseases. Biochem Soc Trans 29:250 267 Worakitkanchanakul W, Imura T, Fukuoka T, Morita T, Sakai H, Abe M, Rujiravanit R, Chavadej S, Minamikawa H, Kitamoto D (2008) Aqueous phase behavior and vesicle formation of natural glycolipid biosurfactant, mannosylerythritol lipid B. Colloids Surf B Biointerfaces 65:106 112 Worakitkanchanakul W, Imura T, Fukuoka T, Morita T, Sakai H, Abe M, Rujiravanit R, Chavadej S, Minamikawa H, Kitamoto D (2009) Phase behavior of ternary mannosylerythritol lipid/water/oil systems. Colloids Surf B Biointerfaces 68:207 212 Zhao X, Wakamatsu Y, Shibahara M, Nomura N, Geltinger C, Nakahara T, Murata T, Yokoyama KK (1999) Mannosylerythritol lipid is a potent inducer of apoptosis and differentiation of mouse melanoma cells in culture. Cancer Res 59:482 486 Zhao X, Geltinger X, Kishikawa S, Ohshima S, Murata S, Nomura N, Nakahara T, Yokoyama KK (2000) Treatment of mouse melanoma cells with phorbol 12 myristate 13 acetate counteracts mannosylerythritollipid induced growth arrest and apoptosis. Cytotechnology 33:123 130 Zhao X, Murata T, Ohno S, Day N, Song J, Nomura N, Nakahara T, Yokoyama KK (2001) Protein kinase Ca plays a critical role in mannosylerythritol lipid induced differentiation of melanoma B16 Cells. J Biol Chem 276:39903 39910

Sophorolipids Inge N.A. Van Bogaert and Wim Soetaert

Contents 1 2 3

Introduction: A Brief History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 180 Structure and Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 Producing Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 3.1 Rhodotorula bogoriensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 3.2 Candida apicola . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184 3.3 Candida bombicola . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184 3.4 Wickerhamiella domercqiae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 3.5 Candida batistae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 4 Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 4.1 Feedstock . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 4.2 Sophorolipid Specific Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 5 The Fermentation Process . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196 5.1 Culture Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196 5.2 Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 5.3 Downstream Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 6 Genetic Engineering of C. bombicola . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 6.1 Developing the Molecular Tool Box . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198 6.2 Genetic Engineering of C. bombicola for the Production of Medium Chain Sophorolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199 7 Applications of Native Sophorolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199 8 Modified Sophorolipids and Their Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 202 9 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205

Abstract Sophorolipids are surface-active compounds synthesized by a selected number of yeast species. Their production by nonpathogenic yeasts at very high yields (over 400 g/L) makes them attractive targets for industrial production and I.N.A. Van Bogaert (*) and W. Soetaert Laboratory of Industrial Biotechnology and Biocatalysis, Department of Biochemical and Microbial Technology, Faculty of Bioscience Engineering, Ghent University, Coupure Links 653, 9000 Ghent, Belgium e mail: [email protected]

G. Sobero´n‐Cha´vez (ed.), Biosurfactants, Microbiology Monographs 20, DOI 10.1007/978 3 642 14490 5 7, # Springer Verlag Berlin Heidelberg 2011

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commercialization and offers an environmental friendly alternative for the petrochemical-based surfactants. Sophorolipids not only display surface-lowering properties but also find application in the cosmetics, pharmaceuticals, and medical field, thanks to their biological activities. Besides the various applications of native and modified sophorolipids, this chapter also discusses the suggested biochemical pathway and its consequences for fermentative sophorolipid production. Furthermore, an overview is given about the various described fermentation processes and the genetic modification of Candida bombicola.

1 Introduction: A Brief History Sophorolipid history starts in the early 1960s when Gorin and colleagues describe the hydroxy fatty acid glycosides of sophorose from Torulopsis magnolia (1961). This osmophilic yeast produces an extracellular oil, heavier than water. The main components of this mysterious oil were determined quite in detail as glycolipids consisting of 2-O-b-D-glucopyranosyl-D-glucopyranose units linked b-glycosidically to 17-Lhydroxyoctadecanoic or 17-L-hydroxy-9-octadecenoic acids, the sugar moieties being partly acetylated (Fig. 1). Later on, in the same decade, several other sophorolipid-producing organisms were described; some of them include Torulopsis gropengiesseri, Torulopsis bombicola, and Candida bogoriensis (Jones 1967; Spencer et al. 1970; Tulloch et al. 1968). It took until the 1980s when new reports on sophorolipids produced by Candida apicola (former Torulopsis magnolia) and Candida bombicola (former Torulopsis bombicola) appeared. At first instance, sophorolipid-producing yeasts were studied because of their alkane assimilating properties, which could be enhanced by sophorolipids (Inoue and Ito 1982); but later on, the focus shifted to the sophorolipids itself. In a climate of growing environmental awareness, biosurfactants regained attention due to their

OR 6⬘

4⬘ HO OR 4⬘⬘

6⬘⬘ 5⬘⬘

O O

3⬘⬘

O O

HO O

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OH

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CH3

1⬘ O

2⬘⬘

HO

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Fig. 1 Structure of a classic sophorolipid (lactonic form) from C. bombicola. R

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biodegradability, low ecotoxicity, and production via fermentation processes based on renewable resources. Sophorolipids turned out to be interesting microbial surfactants from an economic point of view when yields of 70 g/L were reported (Cooper and Paddock 1984). Yields kept increasing due to extensive optimalization of the culture conditions and nowadays over 400 g/L can be achieved (Pekin et al. 2005), rendering sophorolipid an attractive alternative for petroleum-based surfactants. Although several sophorolipid-producing organisms will be discussed in the following pages, this chapter will mainly focus on the sophorolipids synthesized by C. bombicola ATCC 22214. This strain is preferred by most research groups active in the sophorolipid field; it can produce over 400 g/L sophorolipids and now is used for commercial production and applications.

2 Structure and Properties The hydrophilic part of the biosurfactant discussed in this chapter is the disaccharide sophorose. Sophorose is a diglucose with an unusual b-1,2 bond and may contain in the case of sophorolipids acetyl groups at the 60 -and/or 600 positions (Fig. 1). The hydrophobic part of the amphiphilic molecule is made up by a terminal or subterminal hydroxylated fatty acid, b-glycosidically linked to the sophorose molecule. The carboxylic end of the fatty acid is either free (acidic or open form) or internally esterified at the 400 or, in some rare cases, at the 60 -or 600 -position (lactonic form). The hydroxy fatty acid itself counts in general 16 or 18 carbon atoms and can have one or more unsaturated bonds (Asmer et al. 1988; Davila et al. 1993). As such, the sophorolipids synthesized by C. bombicola are in fact a mixture of related molecules with differences in the fatty acid part (chain length, saturation, and position of hydroxylation) and the lactonization and acetylation pattern. These different structural classes cause wide variation in physicochemical properties. Lactonized sophorolipids have different biological and physicochemical properties as compared to acidic forms. In general, lactonic sophorolipids have better surface tension lowering and antimicrobial activity, whereas the acidic ones display better foam production and solubility. Furthermore, acetyl groups render the molecules less water soluble but enhance their antiviral and cytokine stimulating effects (Shah et al. 2005). Diacetylated lactonic sophorolipids have quite a rigid structure; dominance of one such specific type of lactonic sophorolipid results very often in the formation of crystals instead of the more common viscous oil, rendering them relatively easy to isolate (Fig. 2). What exactly determines the degree of lactonization remains unclear, but the lactonic/acidic balance is to a certain point influenced by fermentation conditions such as the level of yeast extract, oxygen supply, and the provided lipidic substrate (Casas and Garcia-Ochoa 1999). Especially when the alkanes hexadecane, heptadecane, or octodecane are used, 85% or more of the sophorolipids are diacetylated lactones (Davila et al. 1994; Cavalero and Cooper 2003). Crystallization is more difficult to achieve with oils or fatty acids but can be

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Fig. 2 (a) sophorolipid crystals as seen in the production medium (light microscope, 100 magnitude); (b) sedimentation of sophorolipids as a brown viscous oil after cultivation

obtained under limited feeding of the lipidic carbon source (Davila et al. 1994; Rau et al. 1996). Sophorolipids lower the surface tension in water from 72.80 mN/m down to 40 30 mN/m, with a critical micelle concentration (CMC) of 40 100 mg/L. These latter values are about two orders of magnitudes lower compared to chemicalderived surfactants, adding to their environmental friendly profile. Recently, Hirata et al. (2009a) investigated the properties of sophorolipid blends with different lactone/acid ratios. Interestingly, the natural blend with 72% lactonic sophorolipids displayed the lowest surface tension and CMC, suggesting that natural synergism between sophorolipids creates a better balance for interfacial activities. Sophorolipids preserve their surface-lowering properties despite high salt concentrations and are, in addition, active across a wide temperature range. A feature one must keep in mind is their instability at pH values higher than 7.0 7.5: beyond this point, irreversible hydrolysis of the acetyl groups and ester bonds is observed. Sophorolipids are readily biodegradable surfactants as determined by standard manometric respirometry and stable metabolite studies (Renkin 2003; Hirata et al. 2009b). There is no evidence for the accumulation of stable metabolites in both the accumulation phase and the running out phase of simulated waste water systems; sophorolipids are totally degraded till 1 carbon metabolites. Furthermore, they are far less toxic for aquatic organisms than the conventional detergents; the inhibitory effect on the crustacea Daphnia magna, the ciliate Tetrahymena terhmophila, and the microalgae Pseudokirchneriella subcapitata is tenfold less as compared to conventional surfactants (Renkin 2003). Tests with crude and acidic sophorolipids pointed out that they are not irritating to the skin, do not trigger allergic reactions, and have an oral safety level, which is greater than or equal to 5 mL/kg weight (Hillion et al. 1998). Cytotoxicity was evaluated by the dimethylthiazoldiphenyltetrazoliumbromide (MTT) method with human epidermal keratinocytes and was proven to be low (Hirata et al. 2009b).

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3 Producing Microorganisms Here, we present several yeast models in which sophorolipids production has been studied.

3.1

Rhodotorula bogoriensis

This yeast species was initially called Candida bogoriensis and is sort of an outlier when compared to the other sophorolipid-producing strains: where all other species are Ascomycota, R. bogoriensis belongs to the Basidiomycota phylum. Also from a structural point of view, the sophorolipids are different. The predominant form contains a C22 hydroxy fatty acid, and even small amounts of C24 sophorolipids are retrieved (Fig. 3). Furthermore, these fatty acids are not terminal or subterminal hydroxylated and linked to the sophorose unit, but they show internal hydroxylation and subsequent binding. This later feature disfavors internal esterification and, consequently, no lactonic sophorolipids are observed (Nunez et al. 2004). On the hand, the sophorose unit can be acetylated identical to the C. bombicola sophorolipids. Light’s research group partially purified and characterized the involved acetyltransferase and some

Fig. 3 Structure of a typical sophorolipid from R. bogoriensis. R

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other sophorolipid-related enzymes, but this will be discussed in the biosynthesis section. Although this organism has been known for several decades, not much attention has been given to the optimalization of the production, as a yield of only a few grams per liter is quite small compared to the ones obtained with C. bombicola fermentations.

3.2

Candida apicola

The first described sophorolipid-producing yeast, T. magnolia, was incorrectly identified and was in 1968 reclassified as Torulopsis apicola (Hajsig), currently known as Candida apicola (Tulloch and Spencer 1968). This species is phylogenetically quite related to C. bombicola and its glycolipids and production characteristics are nearly identical. Some German research groups prefer(red) working with this species (e.g., Hommel et al. 1987).

3.3

Candida bombicola

Candida bombicola was initially named Torulopsis bombicola and will be referred to under this name in older articles. The large majority of the scientific literature on sophorolipids reports on research with this strain. In 1998, Rosa and Lachance (1998) described the novel yeast species Starmerella bombicola and introduced it as the teleomorph of Candida bombicola based on the high 18 S rDNA identity between both strains (more than 98%) and their ability to mate with each other to form ascospores. Just as its anamorph, S. bombicola is able to produce sophorolipids. Although several C. bombicola isolates are available in culture collections, the ATCC22214 isolate is the strain of choice as this one is the most efficient sophorolipid producer (unpublished results). As its name already suggests, the first Candida bombicola isolate was obtained from the honey of a bumblebee or Bombus species (Spencer et al. 1970). Just like Candida bombicola, many yeasts of the Starmerella clade are associated with bees or substrates visited by bees and it is suggested that a mutually beneficial interaction exists (Rosa et al. 2003). As a consequence of its habitat, C. bombicola is able to grow at high sugar concentrations and standard fermentations are usually started with a glucose level of 100 g/L or more. Even though this yeast carries the genus name “Candida,” this does not mean that it is closely related to the commonly known human pathogen Candida albicans. C. bombicola is a nonpathogenic yeast and does not use the alternative translation of the CUG codon (serine instead of leucine) as is described among others for C. albicans, C. cylindracea, C. parapsilosis, C. melibiosica, C. zeylanoides, C. rugosa, C. maltosa, C. tropicalis, C. lusitaniae, C. guillermondii, and C. viswanathii, (Sugita and Nakase 1999).

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Wickerhamiella domercqiae

Chen et al. (2006a) recently proved sophorolipid synthesis in W. domercqiae Y2A, isolated from an oil-containing wastewater sample. They observed more than six glycolipids and identified one of the three main products as 17-L-(-oxy)-octadecanoic acid 1,400 -lactone 60 ,600 -diacetate, which is identical to the major component of the sophorolipids of C. apicola and C. bombicola. Furthermore, the authors demonstrated the potential use of this compound as an antitumor agent (Chen et al. 2006b). Quite surprisingly, a yield of 320 g/L sophorolipids could be obtained, only slightly below the value of 350 g/L for C. bombicola cultivated under the same conditions (Song 2006).

3.5

Candida batistae

Regarding the fact that the production of sophorolipids is not restricted to a single yeast species, but to a number of related microorganisms, it is not unlikely to presume that other species belonging to or related to the Wickerhamiella, Starmerella, and Rhodotorula clades are also capable to synthesize some sort of sophorolipid. This phylogenetic approach was applied on species closely related to C. bombicola and led to the discovery of another sophorolipid-producing yeast: C. batistae. In general, the sophorolipids are similar to the ones of C. bombicola, but there are differences in the relative values of certain components in the mixture. Under the same production conditions for instance, C. batistae produces clearly more acidic sophorolipids than lacontic ones, whereas this is the other way around for C. bombicola. Another interesting feature is the preference of terminal hydroxylation of oleic acid over subterminal hydroxylation. Again, this is different compared to the C. bombicola hydroxylation pattern (see Sect. 4.2.1). 6 g/L of sophorolipids were obtained after 3 days of cultivation, but it is expected that the productivity can be improved (Konishi et al. 2008).

4 Biosynthesis Several aspects of sophorolipids biosynthetic pathway are presented in this part of the review.

4.1

Feedstock

The suggested sophorolipid biosynthesis pathway is outlined in Fig. 4. One can observe two main inputs: a hydrophilic substrate such as glucose and an lipophilic substrate. Ideally, both are provided in the production medium.

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Since sophorolipid-producing yeast strains such as C. bombicola and C. apicola are capable of growing on alkanes, they possess the enzymes required for the terminal oxidation of alkanes, in this particular case cytochrome P450 monooxygenases belonging to the CYP52 family. In the subsequent steps, the derived alcohol will be converted via an aldehyde to its corresponding fatty acid, which can then be

O

(3)

HO

CH3 (4) Fatty acid

(2)

Aldehyde

(1)

Alcohol

n-alkane

Triglyceride

O2, NADPH (5)

H2O, NADP+

O

OH

HO

CH3 Hydroxy fatty acid UDP-Glucose (6)

UDP

OH O

HO HO

O

CH3

OH O OH

Glucolipid UDP-Glucose (7)

UDP

OH

HO OHHO

O

OH O

HO OH HO O

O

CH3

O

(8)

O O

OH

Sophorolipid (acidic form)

Lactonized sophorolipid

Acetyl-CoA

Acetyl-CoA (9)

CoA-SH

H2C

O

HO O HO O

(9)

CoA-SH

O O

HO OHHO

CH3

O O

HO

OH

H3C

O

HO OH HO O

O H3C

O

O

O O

H3C O

CH3

O

O O

OH

Acetylated acidic sophorolipid

HO O HO O

O O

CH3

O

HO OH

Acetylated lactonized sophorolipid

Fig. 4 Proposed sophorolipid biosynthetic pathway. (1) cytochrome P450 monooxygenase, (2) alcohol dehydrogenase, (3) aldehyde dehydrogenase, (4) lipase, (5) cytochrome P450 monooxygen ase, (6) glucosyltransferase I, (7) glucosyltransferase II, (8) lactonesterase, (9) acetyltransferase

Sophorolipids

187

metabolized in b-oxidation or act as precursor for specific biosynthetic processes such as sophorolipid synthesis. The level of sophorolipid formation during fermentations based on alkanes depends on the chain length of the used substrate. Hexadecane, heptadecane, and octadecane give the best production yields. They appear to be directly converted into hydroxy fatty acids (Table 1) and incorporated into the sophorolipid molecules, in this way strongly influencing the fatty acid composition of the sophorolipid mixture (Davila et al. 1994; Cavalero and Cooper 2003). Shorter alkanes are only to a minor extent incorporated, whereas the vast majority is either elongated to C16 or C18 fatty acids or metabolized via b-oxidation. More or less the same is true for eicosane (n-C20) or longer alkanes. A few percentages of eicosane could be detected in the sophorolipid fatty acid moiety, whereas none were observed for the longer alkanes. These alkanes are metabolized by b-oxidation, either completely or partially, to give rise to shorter fatty acids which can be incorporated into the sophorolipid molecules. The yields are however higher when compared to the shorter alkanes and even comparable to those obtained for n-C16 and n-C18 (Tulloch et al. 1962; Jones and Howe 1968). In the same way, also fatty acids directly supplemented to the medium or derived from lipids act as feedstock for sophorolipid synthesis. If no hydrophobic substrate is present in the medium, fatty acids will be formed de novo starting from acetylcoenzyme A (CoA) derived from the glycolysis pathway (Van Bogaert et al. 2008a). Quite obviously, glucose is the hydrophilic substrate of choice. Sucrose can also act as substrate, but the obtained sophorolipid level is lower (Klekner et al. 1991), just as well as for the low-cost alternative soy molasses, which by the way can act as a nitrogen source on top (Solaiman et al. 2007). As C. bombicola was originally isolated from honey, some researchers have tested it as a substrate. It was however only supplemented at the end of the fermentation process when the initially added glucose was consumed. So, despite the good yields, no real conclusions can be drawn on the effect of honey (Pekin et al. 2005). In another attempt to reduce substrate costs, cheese whey was proposed as hydrophilic carbon source. Zhou and Kosaric (1993, 1995) first investigated the fermentation with galactose and lactose, the main sugar components of whey. C. bombicola was not able to grow when only lactose was present, but when vegetable oil was also supplemented, growth and sophorolipid formation were observed. Yet, lactose was not consumed nor could b-galactosidase activity be detected (Daniel et al. 1998a). An overview on the production of sophorolipids on different hydrophilic substrates is given in the first part of Table 1. Experiments with 13C-labeled glucose pointed out that the bulk of the added glucose first passed through glycolysis, in this way supplementing trioses for the gluconeogenesis of glucose for sophorolipid synthesis (Hommel et al. 1994). This explains why fermentations with sugars different to glucose or even without glucose or degradable sugars still yield the conventional sophorolipids.

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Table 1 Overview of sophorolipid production methods on various substrates Substrate Production C yield Remarks References (g/L) (g/g) Among others on the hydrophilic substrate 2% Glucose 0.325 Gobbert et al. (1984) 2% Fructose 0.205 2% Mannose 0.245 2% Saccharose 0.16 2% Maltose 0.1 2% Raffinose 0.205 2% Octadecane 0.3 (0.25@21 C) 2% Paraffin (C14 15) 0.275 (0.36@21 C) 2% Oleic acid 0.35 (0.605@21 C) 2% Soy bean oil 0.34 10% Glucose 38 0.25% YE Klekner et al. (1991) 10% Sucrose 33 10% Glucose 31 Batch Zhou and Kosaric (1993) 10% Glucose 137 10.5% Safflower oil 10% Galactose 9 10% Galactose 24 10.5% Olive oil 10% Lactose 0 10% Lactose 46 10.5% Olive oil 10% Sucrose 19 10% Sucrose 58 10.5% Safflower oil 10% Cheese whey 0 10% Cheese whey 6 10.5% Olive oil 10% Glucose 160 0.4% YE Zhou and 10.5% Canola oil 0.1% Urea Kosaric (1995) 10% Lactose 110 Batch 10.5% Canola oil 10% Cheese whey 12 10.5% Canola oil 10.5% Canola oil 70 10% Glucose 135 10.5% Safflower oil 10.5% Safflower oil 55 10% Glucose 91 10.5% Sunflower oil 10.5% Sunflower oil 47 10% Glucose 83 10.5% Olive oil 10.5% Olive oil 30 (continued)

Sophorolipids Table 1 (continued) Substrate 30% Rapeseed oil, 10% deproteinized whey concentrate Single cell oil from Cryptococcus curvatus grown on deproteinized whey concentrate 40% Rapeseed oil Soy molasses Oleic acid Glucose Oleic acid Glucose Soybean oil Glucose Tallow oil Glucose Linseed oil Soy molasses Oleic acid

189

Production (g/L) 280

Remarks

References

Lactose not consumed

Daniel et al. (1998a)

422

Rapeseed oil added after single cell oil consumption

Daniel et al. (1998b)

21

1% YE 0.1% Urea Fed batch

Solaiman et al. (2004)

79

C yield (g/g)

41 17 54 53

W/o YE and urea; Solaiman et al. soy molasses (2007) as N source With YE and urea fed batch

Soy molasses Oleic acid Glucose Oleic acid Glucose Soybean oil Glucose Edible beef tallow Glucose Linseed oil 10% Glucose 10% Soybean oil

75

29

1% YE 0.1% Urea

10% Sugarcane molasses 10% Soybean oil 10% Sugarcane molasses 10% Soybean oil 10% Sugarcane molasses 10% Soybean oil Sugarcane molasses Soybean oil

23

W/o YE and urea

64

1% YE 0.1% Urea

Turkish corn oil Glucose Honey

>400

Honey added when glucose depleted

100 41 32 54 Daverey and Pakshirajan (2009a)

17 19 Daverey and Pakshirajan (2009b) Pekin et al. (2005) (continued)

190 Table 1 (continued) Substrate

I.N.A. Van Bogaert and W. Soetaert

Production (g/L)

C yield (g/g)

Among others on alkanes 1% Glucose 0.5% Alkane as mentioned

Remarks

SL% with same FA C #

C12 C14 C15 C16 C17 C18 C20 Glucose Rapeseed FAEE Glucose

8 10 8 10 8 10 8 10 8 10 8 10 8 10 320

0.21 0.38 0.35 0.81 0.68 0.24 0.05 0.65

10 18 40 85 100 71 0 Fed batch

20

0.06

Fed batch

Glucose Dodecane Glucose Tetradecane Glucose Hexadecane Glucose Octadecane Glucose Rapeseed FAEE Glucose Rapeseed oil Glucose Sunflower FAME Glucose Sunflower oil Glucose Palm FAME Glucose Palm oil Glucose Linseed FAME Glucose Fish oil Glucose Rapeseed FAEE

17

0.07

0.5% Dried corn steep liquor

20

0.8

95

0.32

175

0.33

340

0.65

255

0.53

235

0.52

172

0.43

240

0.67

82

0.39

122

0.25

51

0.21

250 300 >300

0.61 0.65 Fed batch 6 Feeding strategies

Among others on vegetable oils and fatty acids Glucose only 1

Glucose/safflower oil (2:1) Glucose/safflower oil (1:1) Glucose/safflower oil (1:2)

0.1% YE

References

Cavalero and Cooper (2003)

Davila et al. (1992) Davila et al. (1994)

Davila et al. (1997)

Cooper and Paddock (1984)

3 5 10 (continued)

Sophorolipids Table 1 (continued) Substrate Safflower oil Glucose only Glucose/safflower oil (2:1) Glucose/safflower oil (1:1) Glucose/safflower oil (1:2) Safflower oil 10% Glucose 10% Glucose 10% Safflower oil 10% Glucose 10% Corn oil 10% Glucose 10% Soya bean oil 10% Glucose 10% Sunflower 10% Glucose 10%Soya bean oil 10% Glucose Glucose 10% Glucose Soybean oil

191

Production (g/L) 1 0 10 30 18 0 0 18

C yield (g/g)

Remarks

References

0.5% YE

0.5% YE

20 18 17 67

0.35

15 33

10% Glucose Stearic acid 10% Glucose Stearic FAME 10% Glucose Oleic acid 10% Oleic acid Oleic acid 11% Glucose 10% Soybean oil 11% Glucose 10% Soybean oil Glucose Soybean oil 10% Glucose 10% Palmitic acid

20

10% Glucose 10% Coconut oil 10% Glucose 10% Grapeseed oil 10% Glucose 10% Corn oil 10% Glucose 10% Olive oil 10% Glucose 10% Sunflower oil 10% Glucose 10% Sunflower oil

3

7 L Fermentor 1% YE 0.1% Urea Stepwise feeding of second compound

Asmer et al. (1988)

Lee and Kim (1993)

32 38 77 80

0.37

Batch

120

0.6

Fed batch

40 80

Continuous culture 0.5% YE Batch

4.5

Kim et al. (1997) Casas and Garcia Ochoa (1999)

5 5.5 9.5 11 120 (8d)

0.6

0.1% YE resting cells (continued)

192 Table 1 (continued) Substrate Glucose Oleic acid

Glucose Rapeseed oil Oleic acid Glucose

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Production (g/L) 180

C yield (g/g)

300 (125 h)

0.68

350 1.5 g/(Lh)

Remarks Fed batch SL crystals if limited oleic acid feeding Fed batch

60 46

Glycerol Soy oil FAEE Glycerol Soy oil FAPE Glucose Soybean dark oil Glucose Soybean oil Glucose Corn oil 10% Glucose

42

10% Soybean dark oil 10% Glucose 10% Soybean oil 10% Glucose 10% Corn oil 10% Glucose 10% Rice germ oil 10% Glucose 10% Rapeseed oil Glucose: 30 40 g/L Rapeseed oil 10% Glucose 3.75%Waste frying oil 10% Glucose 4% Restaurant waste oil 10% Glucose 4% Oleic acid

Rau et al. (1996)

Rau et al. (2001)

Fed batch Guilmanov et al. Focus on aeration (2002) [50 80 mM O2/(Lh)]

Among others on waste streams Glycerol 9 Biodiesel co product stream Glycerol Soy oil FAME

References

Ashby et al. (2005) Fed batch SL methyl esters detected No SL esters detected No SL esters detected Fed batch

Ashby et al. (2006)

Kim et al. (2009)

65

Batch Oleic acid contents (%) 16 22

98

24

102

40

120

52

365 (8d)

Fed batch

18 90

Kim et al. (2005)

100 24 65

50 34 42

1% YE 0.1% Urea Batch

Fleurackers (2006) Shah et al. (2007)

(continued)

Sophorolipids Table 1 (continued) Substrate 10% Glucose 4% Lipidic substrate: C14:0 C16:0 C18:0 C20:0 C18:1 C18:2 C18:3 Stearic fatty acid residue Stearic fatty acid residue Tallow fatty acid residue Tallow fatty acid residue Coconut fatty acid residue Coconut fatty acid residue Hydroxylated substrates 15% Glucose 1.5% 2 Dodecanol

193

Production (g/L)

20 33 52 23 60 30 23 43 61 88 120 19 40

C yield (g/g)

0.1 0.15 0.23 0.13 0.26 0.14 0.11 0.18 0.21 0.38 0.41 0.08 0.14

22

Remarks

References

1% YE 0.1% Urea

Felse et al. (2007)

Batch Fed batch Batch Fed batch Batch Fed batch 0.4% YE Only S enantiomere retrieved in SL 0.4% YE

Brakemeier et al. (1998a)

15% Glucose 12 Brakemeier et al. 1.5% 1 Dodecanol (1998b) 15% Glucose 15 1.5% 2 Dodecanone 15% Glucose 17 1.5% 3 Dodecanone 15% Glucose 3 1.5% 4 Dodecanone YE yeast extract, FAME fatty acid methyl ester, FAEE fatty acid ethyl ester, SL sophorolipid

4.2

Sophorolipid-Specific Enzymes

The enzymes that participate in sophorolipid biosynthetic pathway are described.

4.2.1

Hydroxylation of the Fatty Acid and Its Consequences

The incorporation of an appropriate fatty acid into a sophorolipid molecule is preceded by a terminal (o) or subterminal (o-1) hydroxylation step. In yeasts, such reactions are typically performed by cytochrome P450 monooxygenase from the CYP52 family; these enzymes mediate the o or o-1 hydroxylation of alkanes and/or fatty acids. Indeed, Lottermoser et al. (1996) identified two CYP52 genes from C. apicola (CYP52E1 and CYP52E2), but neither of them was linked to a specific biochemical reaction. Yet, evolutionary history of cytochrome P450 genes

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is characterized by several events of gene duplication and conversion, resulting in a broad diversity among those genes also within the genome of a single organism, and it is not always clear what induces them or if they are expressed at all (Nebert and Gonzalez 1987; Nelson 1999). For C. bombicola ATCC 22214, we have identified nine different cytochrome P450 monooxygenase genes belonging to the CYP52 family. Some of them expose very high similarity (91% AA identity) to the CYP52E2 gene of C. apicola, whereas others belong to the new CYP52 subfamilies CYP52M and CYP52N. Whereas CYP52N1 and -E3 are probably involved in alkane assimilation, CYP52M1 takes no part in this process and was clearly induced upon sophorolipid synthesis, indicating a possible role in sophorolipid synthesis (Van Bogaert et al. 2009a). As mentioned before, the fatty acid tail of a sophorolipid preferentially contains 16 or 18 carbon atoms. This strict length distribution is driven by the specificity of the hydroxylation step; divergent fatty acids will either be metabolized via b-oxidation or in the case of shorter fatty acids elongated to C16 or C18 fatty acids. The fact that also arachidonic acid (C20:4) is readily hydroxylated while arachidic acid (C20:0) is not illustrates the enzyme’s specificity towards absolute length and not carbon number. This length also determines the position of the hydroxylation; palmitic acid is predominantly hydroxylated at the terminal position, but the longer the chain, the more the terminal/subterminal oxidation ratio decline; for stearic acid, for example, no terminal oxidation is observed (Tulloch et al. 1962; Davila et al. 1994). Prior to incorporation, fatty acids can undergo desaturation (Brett et al. 1971; Davila et al. 1994). Besides the specificity of the cytochrome P450 monooxygenases, also other factors such as substrate uptake and toxicity, specificity of the subsequent glucosyltransferases, and properties of the glycolipid will influence incorporation of a certain substrate. Nevertheless, it is observed that oils and there derived fatty acids or fatty acid esters with chain lengths of 16 or 18 carbon atoms, such as most common vegetable oils, are readily incorporated and consequently yield high sophorolipid production. In most cases, the fatty acid composition of the vegetable oil used is reflected in the fatty acid pattern of the sophorolipid mixture, illustrating the direct incorporation of the substrates. The use of different oily substrates will be discussed in Sect. 6.2. Yet, the length-controlling action can be circumvented by supplying the yeast with suitable hydroxylated substrates. Brakemeier et al. (1995, 1998a) used secondary alcohols (C12 C16) as the lipophilic carbon source and observed direct incorporation for the majority of the substrate. The resulting compounds display better surface-active properties as compared to native sophorolipids. Furthermore, formation of small amounts of hydroxy fatty acids and alkandiols were detected. These later molecules allow the formation of unusual glycolipids with two sophorose units separated by a hydrocarbonic spacer (Fig. 5), and although these are interesting structures, the amphiphilic configuration of the molecule disappears, rendering it only slightly soluble in water and a less effective surfactant. An issue one must keep in mind when using prehydroxylated substrates is the configuration of the hydroxy fatty acids retrieved in the native sophorolipids; all

Sophorolipids

195 OR

HO HO

HO OR HO O

OR

O O O

O O O

OH

OR

OH O OH OH

OH OH

Fig. 5 Glycolipid with two sophorose units as described by Brakemeier et al. (1998a). R COCH3

H or

o-1 hydroxy fatty acids have the S-configuration, and when racemic mixtures are offered, only the S-enantiomere will be integrated; it looks like the subsequent glucosyltransferase, which couples the hydroxy fatty acids to (uridine diphosphate (UDP)-glucose), requires this specific configuration. Experiments with ricinoleic acid (12-hydroxy-9-cis-octadecenoic acid), having an R-hydroxylgroup at the o-6 position, indeed confirm this hypothesis; this acid is not incorporated into sophorolipid molecules. Alternatively, the position of the hydroxylgroup away from the distal end could sterically hinder the glucosyltransferase activity. The problem regarding the right configuration can be bypassed by using ketones. 2-, 3-, or 4-dodecanones are reduced into their corresponding alcohols by C. bombicola and subsequently incorporated into sophorolipid molecules (Brakemeier et al. 1998b). Although the additional step of reducing the ketone seems less efficient, the yields were actually higher when compared to the corresponding racemic alcohol mixture as only half of the alcohols have the required S-configuration. When the ketone-group moves away from the distal end, less sophorolipids are produced (Table 1); it seems like the in-chain position of the hydroxyl group sterically hinders subsequent enzymatic reactions.

4.2.2

Coupling the Glucose Molecules

Glucose is glycosidically coupled (position C10 ) to the hydroxyl group of the fatty acid through the action of a specific glycosyltransferase I. The transferase reaction requires nucleotide-activated glucose (uridine diphosphate (UDP)-glucose) as glucosyldonor (Breithaupt and Light 1982). In a subsequent step, a second glucose is glycosidically coupled to the C20 position of the first glucose moiety by glycosyltranferase II. Both glycosyltransferases involved in sophorolipid synthesis of R. bogoriensis were partially purified (Esders and Light 1972; Breithaupt and Light 1982). The two enzyme activities could however not be separated and it remains therefore open for discussion whether the consecutive glucose transfers are carried out by two different (but copurified) enzymes or by one and the same (multi)enzyme. It is supposed that sophorolipid synthesis in C. bombicola involves analogous enzymes.

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Final Modifications

Further variation in glycolipid structure occurs through lactonization and acetylation. It is believed that in C. bombicola, a specific lactone esterase gives rise to a closed-chain sophorolipid specific for position C400 . In addition, an acetyltransferase acetylates sophorolipids at positions C60 and C600 (Esders and Light 1972). The reaction is carried out by an acetyl-CoA-dependent acetyl transferase. The transferase from R. bogoriensis has been partially purified, but the corresponding enzyme has not yet been identified in C. bombicola.

5 The Fermentation Process Several aspects of the fermentation process used to produce sophorolipids are discussed.

5.1

Culture Conditions

Sophorolipid synthesis starts when the yeast cells enter stationary phase and is probably triggered by a high carbon-to-nitrogen ratio (Davila et al. 1992). The cells can be kept viable, producing in stationary phase quite well, and a typical production process takes about 10 days. Sophorolipid production medium typically contains over 100 g/L glucose, a source of nitrogen such as yeast extract or corn steep liquor and favorable, but not essential, small amounts of minerals. Higher yields of sophorolipids will be obtained when a hydrophobic carbon source is added, either in batch or fed-batch, and additional glucose feedings can help increase the yield as well. Gobbert et al. (1984) pointed out that the optimal temperature for sophorolipid formation is 21 C, although the optimal growth temperature of C. bombicola ATCC 22214 is 28.8 C. Nevertheless, most fermentations are run at 25 or 30 C due to practical reasons. The amount of obtained sophorolipid is nearly identical for both temperatures, whereas for fermentations at 25 C, biomass growth is lower and the glucose consumption rate is higher as compared to the fermentation at 30 C (Casas and Garcia-Ochoa 1999). During the exponential growth phase, pH drops till 3.5 or below and should ideally be kept at 3.5 for optimal sophorolipid production (Gobbert et al. 1984). Throughout the whole fermentation process, the culture broth should be supplied with enough oxygen; the yeast cells are very sensitive to oxygen limitation during their exponential growth, and good aeration conditions are important for sophorolipid production as the cytochrome P450 monooxygenase uses molecular oxygen. Guilmanov et al. (2002) investigated the effect of aeration by means of shake-flask experiments. The optimal aeration for high sophorolipid yield, expressed in terms of oxygen transfer rate, lies between 50 and 80 mM O2/(Lh). Low aeration levels resulted in the enrichment in saturated fatty acid sophorolipids at the expense of unsaturated fatty acid ones.

Sophorolipids

5.2

197

Substrates

Throughout this chapter, it became quite clear that a lot of substrates can act as hydrophobic carbon source: oils, fatty acids, and their corresponding esters, alkanes, etc. In most cases, relatively pure and well-characterized substrates are used, and although the use of renewable resources such as glucose and vegetable oil already contributes to the environmental friendly character of this surfactant, one can go further and exploit waste streams such as biodiesel by-product streams, soybean dark oil, and waste frying oil (Ashby et al. 2005; Kim et al. 2005; Fleurackers 2006). Despite the abundance of reports on sophorolipid fermentations with various substrates, it is hard to compare them as often different culture conditions or medium compositions are used. Nevertheless, some researchers compared various hydrophilic sources. An overview of various experiments concerning different substrates or feeding strategies is given in Table 1. One can see that vegetable oils rich in oleic acid promote sophorolipid production and, consequently, rapeseed oil is one of the substrates of choice (Davila et al. 1994; Kim et al. 2009). Furthermore, a wellcontrolled fed-batch feeding strategy of both glucose and the hydrophobic substrate will enhance sophorolipid production as well. Nonincorporated substrates, such as alkanes, fatty acids, or esters, are mainly oxidized to CO2, and for good substrates, typically a carbon conversion yield between 60 and 70% is obtained.

5.3

Downstream Processing

Sophorolipids can be extracted from the culture broth with organic solvents such as ethyl acetate. Residual lipidic carbon source can however be coextracted and cause difficulties during further applications. For this reason, additional extraction with hexane is most frequently used, but other solvents such as pentane (Cavalero and Cooper 2003) or t-butyl methyl ether (Rau et al. 2001) can also be applied. Since sophorolipids are heavier than water, it is possible to centrifuge them down or to just decant the sophorolipids from the fermentation medium after heating. This method is very convenient when working with large volumes and high yields. Further elimination of water and impurities can be achieved by addition of polyhydric alcohols and subsequent distillation (Inoue et al. 1980). Quite obviously, the formation of sophorolipid crystals (discussed in Sect. 3) will simplify the downstream procedure. Chromatographic purification with silica gel or preparative reversed phase columns is required if one is interested in a specific sophorolipid structure, e.g., for pharmaceutical applications (Lin 1996).

6 Genetic Engineering of C. bombicola The genetic engineering of yeasts to improve sophorolipid production has not been studied in depth, but there are some approaches that have been undertaken in this respect that are discussed in this section.

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6.1

I.N.A. Van Bogaert and W. Soetaert

Developing the Molecular Tool Box

As been demonstrated in the previous sections, a lot of effort has been spent to optimize the C. bombicola fermentation process in order to improve the yield or reduce production costs, but the genetics of the producing yeast strain itself has been mainly neglected. Nevertheless, this can be a useful tool in the study of the sophorolipid synthesis pathway and open up perspectives for improved production. A first step in the genetic engineering of this yeast species is the development of a suitable transformation and selection method. C. bombicola can be transformed by the slightly modified Lithium-Acetate method used for Saccharomyces cerevisiae (Van Bogaert et al. 2008b) or by electroporation. In search of a selection system, the sophorolipid-producing yeast turned out to be quite resistant to several antibiotics, and the only antibiotic that could be used successfully was hygromycin (Van Bogaert et al. 2008c). In addition, selection systems based on dominant drug resistance genes are expensive and not very suitable for industrial fermentations and/or applications. Therefore, a transformation system based on the auxotrophic marker orotidine-50 -phosphate decarboxylase was developed as well (URA3; Van Bogaert et al. 2008b). For the moment, strains with multiple auxotrophies are constructed and a system for marker recycling is developed, allowing multiple genetic manipulation steps. One of the logical next steps in the genetic engineering of C. bombicola is the construction of an expression system. It was demonstrated that the TEF1a promoter of Eremothecium gossypii and the TK promoter of the Herpes simplex virus also show activity in C. bombicola (Van Bogaert 2008). On the other hand, it is believed that homologous promoters give rise to better and higher expression levels. Glyceraldehyde-3-phosphate dehydrogenase (GPD) is one of the key enzymes taking part in the glycolysis, an essential pathway present in every living cell. The constitutively and highly active promoter of native GPD genes has been successfully applied for the expression of heterologous genes in several yeasts and filamentous fungi. It was demonstrated that even short GPD promoter fragments of 190 bp could still be used for the expression of heterologous genes in C. bombicola. However, promoter fragments bigger than 488 bp tend to recombine with the genomic GPD promoter, in this way knocking out the GPD gene, which is essential for cell viability. This event can be avoided in future expression experiments by flanking the heterologous gene with a functional URA3 gene and transforming the ura3-negative C. bombicola strain. This would lead to recombination at the dysfunctional URA3 gene instead of the GPD promoter. The efficacy of a short GPD promoter can be a convenient characteristic for the construction of compact expression cassettes or vectors for C. bombicola. However, the exact expression level of the different GPD promoter fragments should first be investigated by, e.g., GFP expressing and quantification. When testing ura3 complementation, it became clear that directed integration by homologous recombination happens quite frequently in C. bombicola (Van

Sophorolipids

199

Bogaert et al. 2008b). Therefore, a gene-disruption strategy based on homologous recombination was evaluated. This turned out to be most efficient when large fragments of 1,000 bp of the target gene flank the selection marker. This strategy was successfully applied to create a MFE-2-negative C. bombicola strain. The creation and evaluation of this mutant is outlined in the following section.

6.2

Genetic Engineering of C. bombicola for the Production of Medium-Chain Sophorolipids

As seen forward from Sect. 5.2.1, medium-chain sophorolipid can be obtained when supplying C. bombicola with suitable hydroxylated substrate. Yet, primary alcohols are to a large degree metabolized in the b-oxidation pathway, while secondary alcohols and ketones are rather expensive substrates for industrial use. In order to redirect the primary alcohols towards sophorolipid synthesis, the competing b-oxidation pathway was blocked at the genome level of the yeast by targeting the multifunctional enzyme 2 gene. In contrast to other enzymes contributing to the metabolization of fatty acids, MFE2 is believed to occur as a single copy in the genome, making the construction of a C. bombicola strain with a blocked b-oxidation quite feasible. Several deletion strains were tested in a fermentation run with 1-dodecanol, and all of them showed a 1.7 2.9 times higher production of sophorolipids, indicating that in strains with a blocked b-oxidation pathway, the substrate is redirected towards sophorolipid synthesis. Analog results were obtained for 1-tetradecanol and 1,12-dodecanediol. These experiments illustrate that the mutants can be used to improve the production and yield on medium-chain substrates, in this way lowering the production costs (Van Bogaert et al. 2009b). Somewhat unexpected results were obtained when running fermentations with the mutants on rapeseed oil; instead of equal or slightly better sophorolipid yields, inferior results were obtained. It is possible that the cells are in some way inhibited by dehydrogenated acyl-CoA derivates, which accumulate due to the blocked b-oxidation. This learns that for production of sophorolipid on conventional substrates, the wild type strain should be preferred above the mutant strains.

7 Applications of Native Sophorolipids The most recognized feature of sophorolipids is their ability to act as a surfactant. Surfactants are widely used in the food, pharmaceutical, cosmetic, and cleaning industries and are traditionally produced by chemical means based on petrochemical raw materials. These compounds are often toxic to the environment, and their

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use may lead to significant ecological problems, particularly in washing applications as these surfactants inevitably end up in the environment after use (Mann and Bidwell 2001; Mann and Boddy 2000). The ecotoxicity, bioaccumulation, and biodegradability of surfactants are therefore issues of increasing concern. In this respect, biosurfactants such as sophorolipids offer the advantages of biodegradability, low ecotoxicity, and the production on renewable-resource substrates. Those characteristics draw the attention of the Belgian company Ecover NV (http://www.ecover.com), a manufacturer of ecological detergents and cleansing agents and also active in the natural cosmetics and professional cleansing sector. They saw potential in the use of sophorolipids in hard surface cleaners such as multisurface cleaner, floor soap, and window cleaner (Develter and Fleurackers 2007). These products were redesigned based on a sophorolipid input and released as the so-called eco-surfactants. The company is now evaluating the use of sophorolipids in other products such as detergents. Furthermore, the Japanese company Saraya Co, LTD (http://www.saraya.com) has commercialized sophoron, a dish washer containing sophorolipids as cleaning agent (Futura et al. 2002). Sophorolipids can also be applied in laundry detergents (Hall et al. 1996). The emulsifying properties of sophorolipids can as well be exploited in the petroleum industry; they are useful in secondary oil recovery, in removing hydrocarbons from drill material, and in the regeneration of hydrocarbons from dregs and muds (Baviere et al. 1994; Marchal et al. 1999; Pesce 2002). Sophorolipids can also be applied for in situ bioremediation and degradation of hydrocarbons present in porous media such as soils and groundwater tables (Ducreux et al. 1997; Kang et al. 2010) and in the removal of heavy metals from sediments (Mulligan et al. 2001). Furthermore, the emulsifying property of sophorolipids can be used in the food industry to improve the quality of wheat flour products (Akari and Akari 1987) and in the cold storage transportation in air conditioning systems for the prevention of ice particle formation (Masaru et al. 2001). Furthermore, sophorolipids can be used both as a passive and active ingredient in cosmetics; in addition to its role as emulsifier, they also exhibit several beneficial biological characteristics. First of all, sophorolipids have antibacterial properties and are particularly active against Gram-positive bacteria such as Propionibacterium acnes and Corynebacterium xerosis, the causal agents of acne and dandruff, respectively (Mager et al. 1987). Moreover, they are claimed to trigger several beneficial events regarding the protection of hair and skin, making them attractive components in cosmetic, hygienic, and pharmacodermatological products. They stimulate the dermal fibroblast metabolism and collagen neosynthesis, inhibit free radical and elastase activity, possess macrophage-activating and fibrinolytic properties, and act as desquamating (i.e., eliminating the surface portion of the protective layer of the epidermis as part of the wound healing process) and depigmenting agents (Hillion et al. 1998; Borzeix 1999; Maingault 1999). Sophorolipids are also believed to stimulate the leptin synthesis through adipocytes, in this way reducing the subcutaneous fat overload (Pellecier and Andre´ 2004). The French company Soliance (http://www.groupesoliance.com) produces sophorolipid-based cosmetics

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for body and skin and the Korean MG Intobio Co. Ltd commercializes Sopholine cosmetics. Because of their antimicrobial properties, sophorolipids can be applied in germicidal mixtures for cleaning fruits and vegetables (Pierce and Heilman 1998). The antimicrobial action is not merely restricted towards bacteria; sophorolipids also act as antifungal agents against plant pathogenic fungi such as Phytophthora sp. and Pythium sp. (Yoo et al. 2005) and inhibit algal bloom (Gi 2004). During the last years, it became clear that sophorolipids could have potential as therapeutic agents in different fields. Isoda et al. (1997) were the first to report on their ability to trigger cell differentiation instead of cell proliferation and the inhibition of protein kinase C activity of the human promyelocytic leukemia cell line HL60. The anticancer action is not caused by a simple detergent-like effect, but is attributed to a specific interaction with the plasma membrane. Chen et al. (2006a) conducted anticancer tests with the purified diacetylated lacton. This component exhibits cytotoxic effects on several human cancer cell lines. The cytotoxic effect on the human liver cancer cells H7402 was further investigated and turned out to be attributed to the molecule’s ability to induce apoptosis (Chen et al. 2006b). Furthermore, sophorolipids can be used in the treatment of herpes-related viral infections (Gross and Shah 2007a) and Shah et al. (2005) demonstrated their antihuman immunodeficiency virus and sperm-immobilizing activities. Interestingly, sophorolipids tend to act as a septic shock antagonist in animal models (Bluth et al. 2006). Yet, some questions arose about the mode of action; decrease in mortality could be caused by direct modulation of immune and inflammatory responses or by the antibacterial properties of the sophorolipid molecules (Napolitano 2006). To answer this question, sophorolipids and their derivates were tested against a selection of standard bacterial isolates; they did not show any significant antibacterial activity at clinically relevant concentrations, confirming their potential use as antiinflammatory or immunomodulating agents (Sleiman et al. 2009). Recently, sophorolipids also enter nanotechnology. Metal nanoparticles have been explored in various fields such as catalysis, mechano- and electrical applications and biomedical use. Yet, a prerequisite for his later application is their dispersibility in aqueous solutions, and therefore, one needs a suitable capping agent. Sophorolipids turned out to be good reducing and capping agents for cobalt and silver particles (Kasture et al. 2007, 2008). Singh et al. (2009) demonstrated the antibacterial activity of sophorolipid-coated silver nanoparticles against both Gram-positive and -negative bacteria. They also evaluated the cytotoxic and genotoxic effect of sophorolipid-coated silver and gold nanoparticles and came to the conclusion that gold particles were more cyto- and genocompatible (Singh et al. 2010). In a next step, bioactive molecules will be linked to these nanoparticles and these systems will be evaluated for medicinal and diagnostic applications. Sophorolipids dispersed in water form nm-size micelles of various geometries depending on their concentration. This property allows using them as a structuredirecting agent in the synthesis of nanostructured silica thin films. These materials aroused a growing interest because of their tunable pore size distribution and extremely high specific surface area. These peculiarities make them ideal for

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many applications such as sensing, photovoltaic electrodes, filtration, and catalysis. So far, these materials are almost exclusively made with synthetic surfactants employed as structure-directing agents, which raise important questions about sustainability and toxicity (Baccile et al. 2010). Finally, sophorolipids are a source of difficult to synthesize sophorose and o and o-1 hydroxy fatty acids. The fatty acids can be released after aggressive acid hydrolysis or can be obtained enzymatically (Saerens et al. 2009). They can be directly or indirectly used in polymerization reactions or can be lactonized into macrocyclic esters, which find application in the perfume and fragrance industry (Inoue and Miyamoto 1980). Sophorose is a potent but expensive inducer for commercial cellulase production by the fungi Hypocrea jecorina and Penicillium purpurogenum. Yet, when sophorolipids are supplemented to the H. jecorina production medium in the stationary phase, enhanced cellulase production is observed as well. The fungus is able to degrade the sophorolipids and sets free the inducing sophorose. As a next step, C. bombicola was cocultured with H. jecorina and cellulase production was much higher compared to the earlier experiment (Lo and Ju 2009). Interestingly, sophorolipids are also capable of inducing amylase production in Bacillus subtilis and laccase and manganese peroxidase production in Pleurotus ostreatus. Yet, it remains to be revealed whether the sophorolipids themselves or the sophorose trigger enzyme secretion (Gross and Shah 2007b).

8 Modified Sophorolipids and Their Applications For a number of detergency applications, a native sophorolipid mixture containing a lot of diacetylated lactonic sophorolipids is less suitable as these molecules are less soluble compared to the acid forms. The mixture can however be modified by simply exposing it to alkaline pH; the internal ester bound will be cleaved, as well as the ones binding the acetyl groups, resulting in a more homogeneous mixture of nonacetylated acidic sophorolipids. One can further increase the hydrophilic character of this mixture by cleaving the double bond of oleic acid by ozonolysis. By either applying a reductive or oxidative workup, a terminal aldehyde or carboxyl group is provided. This latter product is an excellent wetting agent and has the potential to act as a calcium sequestering agent (Fig. 6, Develter and Fleurackers 2007). Also, the majority of the other described modifications are carried out at the carboxylic end of the fatty acid. The one first reported in 1971 is the synthesis of sophorolipid alkyl esters to enhance the characteristics of prepared food products such as bakery and oily emulsions (Fig. 7). The authors found that the beneficial effects of the molecules increased with the chain length of the ester (Allingham 1971). Zhang et al. (2004) observed the same trend. They synthesized and compared the properties of sophorolipid methyl, ethyl, propyl, and butyl esters and found that the CMC decreases to about one half per additional carbon group to the

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Fig. 6 Sophorolipid obtained after ozonolysis and oxidative workup of a molecule with an unsaturated fatty acid tail OH

HO

HO OH HO O

O (

O

)6

(

O

(

)n

CH3

O H3C

HO

)7 O

OH

Fig. 7 Sophorolipid alkyl esters, n

0, 1, 2, 3 or 5 (Zhang et al. 2004)

ester moiety. There are a number of patents on the use of these sophorolipid esters in cosmetics (Abe et al. 1981). The sugar moiety hydroxyl groups of the sophorolipid ester can be substituted with hydroxyalkyl groups, giving rise to hydroxypropyl-etherified glycolipids ester. These esters have been used in pencil-shaped lip rouge, lip cream, and eye shadow, in powdered compressed cosmetic material, as well as in aqueous solutions (Kawano et al. 1981a, b). Furthermore, the ester itself can be subjected to further alterations such as transesterification with 1-butanol or 2-methylpropanol (Carr and Bisht 2003) or used for the creation of galactopyranose-sophorolipids (Nunez et al. 2003, Fig. 8). Other researchers synthesized amide derivates with Novozym 435 starting from sophorolipid esters and stated that those derivates may have potential as tunable immunoregulators. The introduction of methacryl or tyrosine groups on the other hand allows the molecules to be functional in polymerization processes (Singh et al. 2003). Yet, sophorolipids can also be subjected to direct enzymatic polymerization. Hu and Ju (2003) optimized the reaction conditions for lipase-mediated conversion of diacetylated lactonic sophorolipids to monoacetylated lactonic sophorolipids,

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HO

HO OH HO O

O O

(

)6

(

O

OH O

CH3

HO

)7

O O HO

OH

OH OH

Fig. 8 Galactopyranose sophorolipid ester (Nunez et al. 2003)

which were in the same reaction polymerized to oligomers and polymers, probably through ring-opening polymerization. Another type of derivative is the amino acid sophorolipid conjugates. The amino acids are coupled to the carboxylic end of acidic sophorolipids by using (di) carbodiimide. In this way, the nonionic sophorolipid can be converted to a cationic, zwitterionic, or anionic surfactant with increased water solubility and polar head groups that allow further chemical derivatization (Zerkowski et al. 2006). Azim et al. (2006) evaluated the antibacterial, anti-HIV, and spermicidal activity of such conjugates. All molecules exhibited the desired action, but leucinesophorolipid was the most effective one. Finally, several enzymatic procedures are described to alter the molecule’s acetylation or lactonization pattern, but as no clear functional relevance or link to a specific application is given, these will not be discussed (Asmer et al. 1988; De Koster et al. 1995; Bisht et al. 1999).

9 Conclusion Sophorolipids belong to the most promising biosurfactants, thanks to the high production yields and synthesis by nonpathogenic species. As sophorolipids find applications in the cleaning, environmental, and food industry as well as in the personal care, cosmetic, and pharmaceutical sectors, it is clear that their economical competitiveness depends on their final utilization. If sophorolipids are, for example, used in the cleaning industry, they have to vie with other environmentally friendly surfactants, such as the alkyl-polyglucosides (APGs), which have a market price of 2 €/kg. However, in the cosmetic or pharmaceutical sectors, higher price dimensions are standard, and therefore sophorolipids should be able to compete, especially when they are used as an active ingredient due to their biological properties. The current production price of native sophorolipids amounts to 2 5 €/kg, depending on substrate cost and production scale, but when additional modifications or the use of speciality substrates are required, the cost will be higher. The production process of native sophorolipids starting from glucose and vegetable oil is already quite mature; yields much higher than 400 g/L are from a biological and practical point of view (e.g., aeration) out of the limit, unless some sort of continuous

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separations system could be developed. The use of low-cost substrates on the other hand has, in most cases, a negative effect on the yield and efficiency; yet, further study on this field could probably overcome some of the problems encountered. Extensive research has been performed on the optimization of the fermentation process and the use of various hydrophobic substrates, while reports on the genetics of the producing yeast strain are scare. However, genetic engineering of these yeast species could open up perspectives for higher yields and modification of the glycolipid mixture produced. In this context, molecular tools were developed for C. bombicola and successful mutant strains can be created.

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Index

A A and C domains, 70 Acetylation, 181, 196, 204 Acetyltransferase Mac1, 156, 157 Mac1 and Mac2, 156, 157 Acyl carrier protein (ACP), 23, 111, 112 Acyl homoserine lactones (AHLs), 26, 30, 31 Acyltransferases, 156, 157 Adenylation (A), 67 Adenylation domain(s), 63, 67 69, 71 A domain(s), 67, 70, 79 AlgC, 24, 28 Alkanes, 180, 181, 186, 187, 193, 194, 197 Alkanethiol SAMs, 168 Amberlite XAD, 161 Amino acid sophorolipid conjugates, 204 Ammonium sulfate, 162 Antiagglomeration agent, 155 Antibacterial activities, 169 Antifungal activities, 169 Antimicrobial activity, 169 Antitumor, 155, 172, 173 Antitumor activity, 110 Atomic force microscopy (AFM), 168 B Bacillomycin, 59, 61, 63 65, 70, 72 Bacillus subtilis, 106 Batch cultivations, 34 36 b hydroxydecanoic acid, 14, 22 b hydroxyfatty acid, 16, 20, 26, 28 Biocompatibility, 165, 171, 173 Biological activity, 135 137 Biomedical field, 172 Bioreactor, 161 Biosurfactants

critical micelle concentration (cmc), 2 fatty acid, 146, 147, 151 154, 156, 158, 160, 169 glycolipids, 146, 151, 155, 158, 161, 165, 167 limitations, 4 5 lipopeptides, 146 origin microorganism, 3 particulate, 146 polymeric, 146 representative structures, 3 Biosynthesis genes, 137 intermediates, 137 pathway, 137 Blue I phase, 165 b oxidation, 158 2 Bromooctanoic acid, 158 Burkholderia, 15, 17, 29, 42, 43 C Caenorhabditis elegans, 109 Candida apicola, 180, 184 Candida batistae, 185 Candida bombicola, 180, 184 C. antarctica KCTC 7804 6 O acetyl 2,3 di O alkanoyl b D mannopyranosyl (1!4) O meso erythritol, 151 Carbon sources, 22, 23, 26 28, 33, 35 39 Carbon to nitrogen ratio, 196 Carcinoma cell lines, 173 C domain, 67, 70 Cellobiose lipid (CL), 147, 155, 156, 159, 172 Cellular differentiation, 109 Cellular processes cell fusion, 164

211

212 Cellular processes (cont.) endocytosis, 164 exocytosis, 164 transport phenomena, 165 Ceramides, 171 Chain shortening, 158 Cheese whey, 187 Chirality, 163 Coacervates, 164 165 Coenzyme A (CoA), 23 Coenzyme Q pattern, 160 Communication mediating (COM) domains, 68, 69, 79 Condensation domain, 68, 69, 71 Configuration, 151, 166, 167 Congeners, 14, 17, 19 21, 40 43 Continuous cultivations, 35 37 Cord factor antigenicity, 128 dimycolate (TMD), 127, 128 potential virulence, 128 toxicity, 128 trehalose, 127 129 two mycolic acids, 128 Corn steep liquor, 35, 39 Cosurfactant, 173 Critical aggregation concentration (CAC), 165 Critical micellar concentration (CMC), 2, 75 77 CYP52, 186, 193, 194 Cytotoxicity, 182

Index Escherichia coli, 16, 95 Exolipids, 94, 100, 103, 110, 115 Expressed sequence tags (ESTs), 158 Extracellular vesicles, 99 101 F FAS II, 23, 25 27, 29 Fatty acids, 181, 183, 187, 193 195, 197, 199, 202 Fatty acid synthetases, 23 Fed batch cultivation, 35, 36 Fluid isotropic (FI) phase, 166 Fluorescence microscopy, 150 Fluorescence probe spectroscopy, 165 Foam fractionation, 40, 41 Foaming, 36 38, 41 Fractal analysis, 105, 106 Freeze fracture, 165

D Diastereomer, 151, 166 Differential scanning calorimetry (DSC), 166 Direct colony TLC, 104 3D ordered lyotropic liquid crystal, 163 Downstream processing, 39 41 Drop collapsing test, 18 Drug delivery, 155, 165, 170 dTDP L rhamnose, 28, 32, 41 Dynamic light scattering (DLS) spectroscopy, 165

G Gas chromatography (GC), 20 Gene cluster, 156 Gene of Serratia hexS, 94, 115, 116 pswP, 110, 111, 113, 115, 116 swrA, 111, 114 swrI, 114 swrW, 111 113, 115, 116 Gene transfection, 155 Giant colony branching fractal, 106 multicellular behavior, 113 Giant vesicles, 164 Glucose, 14, 24, 27, 28, 32, 36, 39 Glyceraldehyde 3 phosphate dehydrogenase (GPD), 198 Glycolipids, 14, 22 Glycoproteins, 168, 170, 172 Glycosides, 16 Glycosyltranferase, 195 Emt1, 155 157 Gompertz equation, 34

E Ecover, 200 Efficient MEL A producers, 159 Electrospray ionization (ESI), 20 Environmental, 29, 30, 33 34, 43 Epimerase domain (E), 69 Epimerisation (E), 67, 68 Epimerisation domains, 63, 70, 71 Erythritol, 145 173

H HAA, 12 Hemolysin, 15 Heterologous production, 41 42 High level MEL producers, 159 High performance liquid chromatography (HPLC), 20 HPLC. See High performance liquid chromatography

Index Hybrid bilayer membranes, 168 4 Hydroxy 2 alkylquinolines (HAQs), 26, 27, 30, 31 Hydroxy fatty acids, 180, 181, 183, 187, 194, 195, 202 I Ice slurry systems, 155, 171 Immobilized cells, 35, 36 Immunoglobulins, 168, 170, 173 In situ product removal (ISPR), 38 Internal transcribed spacer, 159 Internal transcribed spacer 1 (ITS1), 159 Ion exchange chromatography, 40 Iron (Fe), 33, 37 39 K Ketone, 195, 199 Kurtzmanomyces, 147, 151, 159 Kurtzmanomyces sp. I 11 6 O b D mannopyranosyl (1!4) O mesoerythritol, 151 L Lactonic form, 180 183, 202, 203 L a dilauroylphosphatidyl choline (DLPC), 165 LasR, 30 32 Lectins, 155, 170, 172, 173 Lichenysin, 59, 61, 63, 65, 68 70, 74, 79, 80 Lipopolysaccharide (LPS), 27, 28 Liposome DNA complex, 170 L3 phase, 165 Lyotropic liquid crystals cubic, 163, 164, 166 hexagonal phase, 166 lamella, 166 sponge, 167 LysR, 94, 115, 116 M Major facilitator family mmf1, 157 Mannosylerythritol lipids (MELs) MEL A, 146, 147, 151, 154, 155, 159, 160, 164 171, 173 MEL B, 146, 147, 151, 154, 160, 164 167, 169 MEL C, 146, 147, 151 154, 160, 164, 166 MEL D, 146, 158 Mannosylmannitollipid (MML) 6 O [(40 ,60 di O acetyl 20 ,30 di O alka(e) noyl) b D mannopyranosyl] D mannitol, 154

213 Manometric respirometry, 182 Mass spectrometry (MS), 20 21 MEL A/water/n decane, 167 MEL B/water/n decane, 167 MEL C 4 O [(20 ,40 di O acetyl 30 O alka(e)noyl) b D mannopyranosyl] D erythritol, 152 4 O [40 O acetyl 20 ,30 di O alka(e)noyl b D mannopyranosyl] D erythritol, 153 4 O [(40 O acetyl 30 O alka(e)noyl 20 O buta noyl) b D mannopyranosyl] D erythritol, 152 Metal nanoparticles, 201 MFE2, 199 Micelles formation, 2 6 Microemulsion, 167, 173 Modifications, 180, 196, 202, 204, 205 Moisturizing activity, 171 Mono acylated MEL 4 O (30 O alka(e)noyl b D mannopyranosyl) D erythritol, 153 MTT method, 171 Multivalent effect, 168 MvfR, 30 32 Mycosamine transferases, 156 Mycosubtilin, 59, 63 65, 70, 73 75, 80, 81, 83 N NBRC 10877, 150, 151, 154, 159 Nile Red, 150 Nitrate, 33, 35, 38, 39 Non pathogenic, 41 Non ribosomal code, 67, 79 Nonribosomal peptide synthetase (NRPS) surfactin synthetase SfrA, 114 unimodular NRPS, 113 W1 synthetase SwrW, 111 Non ribosomal peptide synthetases (NRPS), 58, 61 63, 65, 67 72, 79 NRPS communication mediating (COM) domains, 68 Nutritional, 29, 33, 38 40 O 4 O b D mannopyranosyl meso erythritol, 146, 160 4 O (20 ,60 di O acyl b D mannopyranosyl) D erythritol, 150 Oil in liquid crystal (O/LC) emulsion, 167 Oils, 180 182, 185, 187, 194, 197, 199, 200, 204

214 Opportunistic pathogen, 15, 43 Optimal temperature, 196 Orcinol, 18, 19 Ozonolysis, 202, 203 P P. antarctica JCM 10317, 154 P. antarctica T 34, 153, 158 PCP. See Peptidyl carrier protein PCP domain(s), 69, 70 Peptidyl carrier protein (PCP), 67, 69, 70 Phosphate cascade systems., 170 Physicochemical property CMC values, 135 emulsion stabilizing, 135 stabile, 135 strong surface activity, 135 surface tension, 135 Physiology, 172 Pili, 106, 107 PKS/NRPS, 68, 70 71 Plants, 159, 172 Plumber0 s nightmare, 165 Poly (2 hydroethyl methacrylate) (PHEMA), 168 Poly(3 hydroxyalkanoates), 24, 26 Polyhydroxyalkanoates, 24, 25 Polyketide synthases (PKS), 68, 70, 71 PPTase, 110, 111, 113 Precipitation, 40 Prodigiosin, 96 98, 100, 101, 110, 113 116 Production biosurfactant, 6 7 cell bound, 139 extracellular, 139 growth associated, 139 140 growth limiting, 137, 139 growth uncoupled, 139 by resting cells, yield, 137 wall associated, 139 Productivity, 36, 37, 39, 43 Protein kinase C, 170 Protein kinase Ca, 170 P. rugulosa NBRC 10877, 151, 154, 159 Pseudomolecular ion, 21 Pseudomonas, 25 27, 36, 41 43 Pseudomonas aeruginosa, 14, 15, 17, 23, 25 43, 103 Pseudozyma flocculosa, 160 Pseudozyma fusiformata, 160 Pseudozyma graminicola CBS 10092, 153 Pseudozyma hubeiensis, 152

Index Pseudozyma parantarctica JCM11752, 154, 162, 163 Pseudozyma prolifica, 160 Pseudozyma rugulosa, 151, 152, 154, 159, 162, 163 Pseudozyma shanxiensis, 152 Pseudozyma siamensis CBS 9960, 152 Pseudozyma sp, 147, 160 163, 172 Pseudozyma thailandica, 160 Pseudozyma tsukubaensis 1 O b (20 ,30 di O alka(e)noyl 60 O acetyl D mannopyranosyl) D erythritol, 151 Pumilacidin, 59, 63 Q Quorum sensing, 24 27, 30 34, 43 N acyl homoserine lactones (AHLs), 114 R Real time reverse transcriptase PCR, 158 Reciprocal shaker, 161 Repellent, 109 Resting cells, 35, 36, 163 Rhamnolipids, 13 43, 103, 104, 106, 109, 116 Rhamnose, 14 17, 19 22, 24, 25, 27 28, 32, 41, 42 Rhamnosylation, 22, 26 Rhamnosyltransferase, 22, 24, 28 32, 41, 42 RhlA, 24, 25, 28 34, 41, 42 RhlB, 24, 26, 28 32, 42 RhlC, 24, 26, 28 32, 42 RhlR, 30 34 Rhodotorula bogoriensis, 183 184 rmlBDAC, 32, 41 RpoN, 33 Rubiwettin, 97, 99 104 S S adenosylmethionine, 24, 27 Salmonella, 106, 109 Saraya, 200 Schizonella melanogramma, 147, 169 Schizonellin, 147 Secondary alcohols, 194, 199 Self assembled monolayers (SAMs), 164, 167 169 Self assembling properties, 155, 164, 165, 168, 169, 172, 173 bicontinuous cubic phase (V2), 164, 166, 167 hexagonal (H2), 163, 166, 167

Index lamella phase, 163, 164, 166 lamellar (La), 166, 167 large unilamellar vesicles (LUV), 165 166 liposomes, 155, 164 liquid lyotropic crystals, 163, 164, 166, 167 L3 phase, 165 multilamellar vesicles (MLV), 165 166 self assembled mono layer (SAM), 164, 167 169 sponge phase, 164, 167, 1675 Semi continuous cultivations, 36 37 Serratamolide, 102, 104, 110 Serratia liquefaciens, 101 Serratia marcescens, 95 102, 104 111, 113 116 Serratia rubidaea, 97 100, 103 Serrawettin W1, 96 98, 101 105, 107 116 W2, 96 98, 101 105, 108, 109, 111, 113, 114, 116 W3, 96 98, 101 103, 105, 108, 109, 116 Sfp, 67, 80 Signal transduction, 168, 170 Soap stock, 39, 40 Sodium nitrate, 162 Solid medium hard agar medium, 104, 105, 109 low agar medium, 106 nutrient poor, 107, 109, 117 Solid state fermentation, 35, 37 Solvent extraction, 161, 162 Sopholine, 201 Sophorolipids, 147, 172 Sophorose, 180, 181, 183, 194, 195, 202 Soy molasses, 187 Specificity, 172 Sponge phases, 164, 165, 167 Spreading growth. See also Swarming flagellum dependent, 107, 109 flagellum independent, 104 107 surfactant dependent, 107, 116, 117 Staphylococus aureus protein A, 168 Structure function relationship, 167, 173 Substrates C14 alkane, 138 C16 alkane, 138 nonalkanes, 139 Succinoyl trehalose lipid extracellular, 131 powerful surfactants, 131 succinic acid containing, 131 versatile biochemical actions, 131

215 Surface activity, 15 containment force, 106, 107 surface tension, 93, 107, 116 wetting activity, 95 100, 113, 116 Surface plasmon resonance (SPR), 168 Surface tension, 181, 182 Surfactants, 146, 163, 166, 173 Surfactin, 106 109, 111, 114 Swarming hyperflagellate, 109 multicellular behavior, 113 unipolar flagella, 109 Synchrotron small angle X ray scattering (SAXS) spectroscopy, 165 T TE domain, 69 Temperature dependent phenotype exolipid production in E. coli transformant, 114 116 Tensiometer, 18 Thin layer chromatography, 19 Thioesterase, 69 71 Thioesterase (TE), 67 Thioesterase domain (TE), 67, 69, 70 Thiolation (T), condensation (C), 67 Torulopsis bombicola, 180, 184 Torulopsis gropengiesseri, 180 Torulopsis magnolia, 180 Transcriptional regulator, 30, 31 Transformation, 198 Transmission electron microscopy (FF TEM), 165 Trehalolipid identified, 122, 127, 132, 136 interest, 122 production, 137 140 purified, 129, 130, 133, 136 trehalose, 121 140 trehalose lipid, 121 140 Trehalolipid producing bacteria Actinomycetales, 122 Arthrobacter, 122 Brevibacteria, 122 Corynebacterium, 122, 126 Gordonia, 122 Micrococcus, 122 Mycobacterium, 122, 126 Mycobacterium avium, 126 Mycobacterium bovis, 126 Mycobacterium flavescens, 126 Mycobacterium paraffinicum, 122, 126 Mycobacterium phlei, 126

216 Trehalolipid producing bacteria (cont.) Mycobacterium tuberculosis, 122, 126 Mycobacteriurn fortuitum, 126 Nocardia, 122 Pseudomonas, 123 Rhodococcus, 122, 126 Rhodococcus erythropolis, 122, 126 Rhodococcus fascians, 126 Trehalose containing glycolipid 2,3 di O acyltrehalose (DAT), 132 mycoside F, 132 trehalose diesters, 129 133 triesters, 126, 132 133 Trehalose lipids corynomycolic, 127, 137 di acyltrehalose, 132 dicorynomycolates, 129, 135, 137, 139 hexa acyltrehalose, 134 ketomycolates, 129 monocorynomycolates, 129 mycolic acids, 126 130, 134 nocardiomycolic acids, 130 octaacyltrehalose, 134 octaacyl trehalose, 129 succinoyl containing trehalose lipid, 136 succinoyl trehalose, 29 succinoyl trehalose lipid (STL), 131 132, 135, 136 surface active, 129 139 TDCMs, 136 TDM (cord factor), 128, 136 tetraester, 133 trehalose, 121 140 trehalose diester, 127 133 trehalose succinic acid, 134 Trehalose tetraesters anionic, 133 decanoic, 133 decanoic acid, 133

Index octanoic, 133 octanoic acids, 133 succinate, 133 succinic acid, 133 Triacylated MEL 1 O alka(e)noyl 4 O [(40 ,60 di O acetyl 20 ,30 di O alka(e)noyl) b D mannopyranosyl] D erythritol, 152 U Ultrafiltration, 40 Ustilago maydis NBRC 5346, 154 Unilamellar vesicles, 164 166 Ustilagic acid, 155, 160 Ustilaginales, 159 Ustilago, 146, 147, 159, 172 Ustilago cyanodontis NBRC 7530, 153 Ustilago maydis NBRC 5346, 154 Ustilago scitaminea, 151, 154, 163 Ustilago scitaminea NBRC 32730 4 O b ( 20 ,30 di O alka(e)noyl 60 O acetyl D mannopyranosyl) erythritol, 151 V Vegetable oils, 28, 38, 154, 162 Vesicles, 146, 164 166. 170 171 Viral infections, 201 Virulence factor, 14, 30 W Water channel, 166 Water in oil microemulsion, 167, 173 Wickerhamiella domercqiae, 185 Wide angle X ray scattering (WAXS) spectroscopy, 165 Y Yield, 14, 22, 32, 34, 36, 37, 39, 40, 42, 43