Biosurfactants: Research and Development 032391697X, 9780323916974

Table of contents :
Biosurfactants
Contributors
Copyright
Preface
Microbial bio-based amphiphiles (biosurfactants): General aspects on critical micelle concentration, surface t ...
Introduction
Biosurfactants in solution
Hydrophilic-lipophilic balance (HLB)
Surface tension (ST) and critical micelle concentration (CMC)
Surface tension and CMC data dispersion
Self-assembly and phase diagrams
Micelles in solution
The fiber phase
Membranes
Less common structures
Comments
Unique macroscale properties of biomphiphiles
References
New insights in biosurfactants research
Introduction
Novel and traditional BS and their producers revisited
Biocatalysis, chemical, and genetic engineering strategies in BS research
Novel applications of BS
Concluding remarks
References
Bioinspired glycolipids: Metals interactions and aqueous-source metal recovery technologies
Introduction
Glycolipids
Microbially produced glycolipids
Bioinspired glycolipids
Complexation of metals by rhamnolipid
Metal selectivity
Mechanism of interaction
Glycolipid-based mining of metals from aqueous sources
Aqueous resources for critical materials
Exploitable aqueous resources
Potential to meet critical materials demand
Approaches and technologies
Ion flotation
Micellar-enhanced ultrafiltration
Precipitation-based technologies
Environmental benefits of glycolipid-based aqueous mining
Rhamnolipid treatment of metal-contaminated groundwater: A case study of uranium in Arizona
Conclusion
References
Rhamnolipids-Has the promise come true?
Introduction
Rhamnolipids prospects in retrospect view
Exploitation of antibiotic activity
Applications in environmental remediation, petroleum, and related areas
Cosmetics, personal care, and household products
Food application
Rhamnolipid bioproduction
Conclusions
References
Biosurfactants as food additives: New trends and applications
Biosurfactants in food formulation
BS as emulsifiers
Food preservatives: BS as antimicrobial agents
Antioxidants
Use of BS in food processing
BS as antiadhesive and antibiofilm agents
BS as cleaning agents
Nanotechnology, food, and BS
BS in food nanotechnology
Concluding remarks
References
Novel approaches in the use of biosurfactants in the oil industry and environmental remediation
Introduction
Types of biosurfactants
Marine biosurfactant-producing bacteria
Pseudomonas
Bacillus
Acinetobacter
Antarctobacter
Rhodococcus
Halomonas
Alcanivorax
Pseudoalteromonas
Marinobacter
Current exploitation of biosurfactants in the oil industry
Soil bioremediation
Microbial enhanced oil recovery (MEOR)
Marine oil spill response
Recent trends in the development of bio-based dispersants to combat marine oils spills
Conclusion and perspectives
References
Biosurfactants produced from corn steep liquor and other nonconventional sources: Their application in differe ...
Introduction
Use of naturally produced biosurfactants from CSL in different industries
Environmental applications
Nanotechnology applications
Cosmetic, pharmaceutical, and personal care applications
Agrochemical applications
Food applications
Use of other nonconventional sources to produce biosurfactants
Dairy industrial wastes
Fruit and vegetable wastes
Starch-rich wastes
Lignocellulosic wastes
Oily and glycerol-based wastes
Concluding remarks
Acknowledgment
References
Metabolic and process engineering on the edge-Rhamnolipids are a true challenge: A review
Introduction
Design of an optimal expression cassette
Biosynthetic genes
Promoter
Untranslated regions and translation initiation
Accessory genes
Integration concepts: Genome-based vs plasmid-based expression
Development of an enhanced chassis cell
Enhancing precursor-availability
Reduction of by-product formation and observed metabolic burden
Accessibility of uncommon substrates
Strain engineering for improved process control
Alternative genetic targets for improved RL synthesis
Fermentation of P. putida for production of RL
Cultivation strategies
Design considerations and hardware concepts
Dispersion and mixing
Preventing and disrupting foam formation
Design of foam-free production processes
Considerations for future bioeconomical-technical substrates
Bioreactor-coupled integrated downstream processing
Foam fractionation concepts
Purification based on micelle-forming properties
Concluding remarks
Acknowledgments
References
Improved production of novel (bola) glycolipid biosurfactants with the yeast Starmerella bombicola through an ...
Introduction
Diversifying and boosting glycolipid production with S. bombicola
Application of integrated -omics strategies for improved glycolipid biosynthesis with S. bombicola
A multiomics approach in industrial biotechnology: A work in progress
Case studies in omics integration in glycolipid production
Omics development in microbial fermentations and future perspectives
Acknowledgments and funding
References
Increasing the natural biodiversity of microbial lipopeptides using a synthetic biology approach
High natural biodiversity of lipopeptides
Bacterial lipopeptides
Lipopeptides produced by Bacillales
Bacillus-related lipopeptides
Surfactin
Iturin
Fengycin
Kurstakin
Other lipopeptides from Bacillus sp
Paenibacillus-related lipopeptides
Brevibacillus-related lipopeptides
Other lipopeptides produced by Bacillales
Actinobacteria-related lipopeptides
Pseudomonas-related lipopeptides
Pseudofactin
Viscosin
Orfamide
Amphisin
Putisolvin
Entolysin
Xantolysin
Tolaasin
Syringomycin
Syringopeptin 22
Syringopeptin 25
Syringafactin
Burkholderiales-related lipopeptides
Serratia-related lipopeptides
Serrawettin
Stephensiolide
Cyanobacteria-related lipopeptides
Other bacterial lipopeptides
Eukaryotic lipopeptides
Yeast and fungi-related lipopeptides
Animal-related lipopeptide
Uncharacterized biosurfactant lipopeptides
Production of novel lipopeptides
Change in the composition of amino acids
Precursor directed biosynthesis
Substrate recognition domain
Domain and module exchange
Altering the number of monomers
Altering monomer connectivity
Tailoring enzyme-Change conformation
Remodeling/assembling of biosynthetic gene clusters
Structure modification by modification of the FA moiety
Improving the homologous production of lipopeptides
Targeting gene regulation
Increase in transcription
Targeting the FA metabolism
Targeting the amino acid metabolism
Targeting the genome
Increase in transporters and toxicity resistance genes
Degradation
Heterologous production
Deciphering the complete biosynthesis mechanism
Choice of the host strain
Cloning strategy
Conclusion
Funding
References
Synthetic approaches to production of rhamnolipid and related glycolipids
Introduction
Rhamnolipids-Biosynthetic versus chemically synthesized
Congener distribution
Stereochemistry
Purity
Tailorability
Chemical synthesis of rhamnolipids
Commercialization of glycolipid synthesis
Performance of synthetic glycolipids
Conclusion
References
The use of biocatalysis for biosurfactant production
Introduction
Glycosyl hydrolases and/or glycosyl transferases
Reverse hydrolysis reaction (condensation)
Transglycosidation
Lipases
Glycolipids surfactants
N-acylation of amino acids
Proteases
Factors affecting the enzymatic production of biosurfactants
The nature of biocatalyst
The reactions media
Substrate ratio
Time of reaction
The immobilization of enzymes
Conclusions
Acknowledgments
References
Challenges and prospects for microbial biosurfactant research
Biosurfactants represent much more than environmental-friendly alternatives for chemical surfactants
Synthetic biology and omics approaches in biosurfactants research
Novel approaches for the sustainable production of biosurfactants
Bioinspired surfactants
Concluding remarks
References

Citation preview

Biosurfactants

FOUNDATIONS AND FRONTIERS IN ENZYMOLOGY SERIES Series Editor: Munishwar Nath Gupta

Series Website: https://www.elsevier.com/books-and-journals/book-series/books-serieslanding-page-request-foundations-and-frontiers-in-enzymology

Foundations and Frontiers in Enzymology

Biosurfactants Research and Development

Edited by

Gloria Sobero´n-Cha´vez

Departamento de Biologı´a Molecular y Biotecnologı´a, Instituto de Investigaciones Biom edicas, Universidad Nacional Auto´noma de M exico, Ciudad Universitaria, CDMX, Mexico

Contributors Rodrigo Arreola-Barroso Department of Cell Engineering and Biocatalysis, Institute of Biotechnology, National Autonomous University of Mexico, Cuernavaca, Morelos, Mexico Niki Baccile Sorbonne Universit
e, Laboratoire de Chimie de la Matie`re Condens
ee de Paris (LCMCP), UMR CNRS 7574, Paris, France Isabel Bator iAMB - Institute of Applied Microbiology, ABBt - Aachen Biology and Biotechnology, RWTH Aachen University, Aachen, Germany Lars Mathias Blank iAMB - Institute of Applied Microbiology, ABBt - Aachen Biology and Biotechnology, RWTH Aachen University, Aachen, Germany Stijn Bovijn Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent, Belgium Chett J. Boxley GlycoSurf, Inc., Salt Lake City, UT, United States Martijn Castelein Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent, Belgium B. Cid-P
erez Chemical Engineering Department, CINTECX; Analytical and Food Chemistry Department, Faculty of Chemistry, University of Vigo, Vigo, Spain J.M. Cruz Chemical Engineering Department, CINTECX, University of Vigo, Vigo, Spain Paula de Camargo Bertuso Interunits Graduate Program in Bioengineering, University of Sa˜o Paulo, Sa˜o Carlos, SP, Brazil Nicolas de Fooz Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent; Laboratory of Integrative Metabolomics (LIMET), Department of Translational Physiology, Infectiology and Public Health, Faculty of Veterinary Medicine Ghent University, Salisburylaan, Merelbeke, Belgium Eric D
eziel Centre Armand-Frappier Sant
e Biotechnologie, Institut National de la Recherche Scientifique (INRS), Universit
e du Qu
ebec, Laval, QC, Canada Sven Dierickx Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent; Laboratory of

xiii

xiv

Contributors

Integrative Metabolomics (LIMET), Department of Translational Physiology, Infectiology and Public Health, Faculty of Veterinary Medicine Ghent University, Salisburylaan, Merelbeke, Belgium Holger Dittmann Department of Bioprocess Engineering, University of Hohenheim, Institute of Food Science and Biotechnology, Stuttgart, Germany Melanie Filbig iAMB - Institute of Applied Microbiology, ABBt - Aachen Biology and Biotechnology, RWTH Aachen University, Aachen, Germany Sigrid G€ orgen Microbial Processes and Interactions Lab (MiPI), TERRA Teaching and Research Centre, Cross border Joint Research Unit (UMRt) BioEcoAgro, Gembloux Agro-Bio Tech/University of Lie`ge, Gembloux, Belgium Tony Gutierrez Institute of Mechanical, Process and Energy Engineering, School of Engineering and Physical Sciences, Heriot-Watt University, Edinburgh, United Kingdom Rudolf Hausmann Department of Bioprocess Engineering, University of Hohenheim, Institute of Food Science and Biotechnology, Stuttgart, Germany Marius Henkel Department of Bioprocess Engineering, University of Hohenheim, Institute of Food Science and Biotechnology, Stuttgart, Germany David E. Hogan Department of Environmental Science, University of Arizona, Tucson, AZ, United States Alexis C.R. Hoste Microbial Processes and Interactions Lab (MiPI), TERRA Teaching and Research Centre, Cross border Joint Research Unit (UMRt) BioEcoAgro, Gembloux Agro-Bio Tech/University of Lie`ge, Gembloux, Belgium Philippe Jacques Microbial Processes and Interactions Lab (MiPI), TERRA Teaching and Research Centre, Cross border Joint Research Unit (UMRt) BioEcoAgro, Gembloux Agro-Bio Tech/University of Lie`ge, Gembloux, Belgium Sonja Kubicki €sseldorf, Ju €lich, Institute of Molecular Enzyme Technology, Heinrich-Heine-Universit€at Du Germany Alexey Llopiz Department of Cell Engineering and Biocatalysis, Institute of Biotechnology, National Autonomous University of Mexico, Cuernavaca, Morelos, Mexico A. Lo´pez-Prieto Chemical Engineering Department, CINTECX, University of Vigo, Vigo, Spain Goedele Luyten Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent, Belgium

Contributors

xv

Raina M. Maier Department of Environmental Science, University of Arizona, Tucson, AZ, United States A. Martı´nez-Arcos Chemical Engineering Department, CINTECX, University of Vigo, Vigo, Spain A.B. Moldes Chemical Engineering Department, CINTECX, University of Vigo, Vigo, Spain Christina Nikolova Institute of Mechanical, Process and Energy Engineering, School of Engineering and Physical Sciences, Heriot-Watt University, Edinburgh, United Kingdom Marcia Nitschke Sa˜o Carlos Institute of Chemistry; Interunits Graduate Program in Bioengineering, University of Sa˜o Paulo, Sa˜o Carlos, SP, Brazil Tathiane Ferroni Passos Sa˜o Carlos Institute of Chemistry, University of Sa˜o Paulo, Sa˜o Carlos, SP, Brazil Alexandre Poirier Sorbonne Universit
e, Laboratoire de Chimie de la Matie`re Condens
ee de Paris (LCMCP), UMR CNRS 7574, Paris, France Sophie Roelants Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent, Belgium Gloria Saab-Rincon Department of Cell Engineering and Biocatalysis, Institute of Biotechnology, National Autonomous University of Mexico, Cuernavaca, Morelos, Mexico Luis Servı´n-Gonza´lez Departamento de Biologı´a Molecular y Biotecnologı´a, Instituto de Investigaciones Biom
edicas, Universidad Nacional Auto´noma de M
exico, Ciudad Universitaria, CDMX, Mexico Gloria Sobero´n-Cha´vez Departamento de Biologı´a Molecular y Biotecnologı´a, Instituto de Investigaciones Biom
edicas, Universidad Nacional Auto´noma de M
exico, Ciudad Universitaria, CDMX, Mexico Wim Soetaert Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent, Belgium Martı´n P. Soto-Aceves Departamento de Biologı´a Molecular y Biotecnologı´a, Instituto de Investigaciones Biom
edicas, Universidad Nacional Auto´noma de M
exico, Ciudad Universitaria, CDMX, Mexico; Department of Microbiology, University of Washington, Seattle, WA, United States Ryan M. Stolley GlycoSurf, Inc., Salt Lake City, UT, United States Stephan Thies €sseldorf, Ju €lich, Institute of Molecular Enzyme Technology, Heinrich-Heine-Universit€at Du Germany

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Contributors

Till Tiso iAMB - Institute of Applied Microbiology, ABBt - Aachen Biology and Biotechnology, RWTH Aachen University, Aachen, Germany Lisa Van Renterghem Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent, Belgium Lynn Vanhaecke Laboratory of Integrative Metabolomics (LIMET), Department of Translational Physiology, Infectiology and Public Health, Faculty of Veterinary Medicine Ghent University, Salisburylaan, Merelbeke, Belgium X. Vecino Chemical Engineering Department, CINTECX, University of Vigo, Vigo, Spain

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Publisher: Stacy Masucci Acquisitions Editor: Peter B. Linsley Editorial Project Manager: Barbara Makinster Production Project Manager: Punithavathy Govindaradjane Cover Designer: Mark Rogers Typeset by STRAIVE, India

Preface Biosurfactants: Research and Development presents different aspects of the fascinating molecules, with different chemical structures but with the common ability to act as surfactants, produced by different types of microorganisms like bacteria and fungi. This book highlights several unique characteristics and applications of biosurfactants that rely not only on their tension-active properties but also on their biological activities, presents some of the challenges of their industrial application, and discusses the main research areas in this emerging field. This book will be of interest not only to experts in the field, who will be able to review the most recent research results and the development of novel applications of biosurfactants, but also to students or young scientists interested in the areas of microbiology and biotechnology, who can learn about the fascinating properties of biosurfactants as well as the microorganisms that produce them and their genetic manipulation to increase their productivity. A unique characteristic of this book is that it contains contributions from authors working on different approaches and biological models; it includes a section that presents novel approaches for the synthesis of biosurfactants that are based on synthetic approaches or biocatalysis. Thus, Biosurfactants: Research and Development fulfills the need of having a book that describes the fascinating world of biosurfactants, presents an updated review of the field, and combines different approaches from the more fundamental aspects of these molecules to their different potential applications, which will be interesting to a wide audience. Surfactants, as tension-active compounds, may act as detergents, wetting agents, emulsifiers, foaming agents, and dispersants. These characteristics make them amenable for a wide range of industrial applications ranging from oil recovery, to agriculture, as part of soaps and detergents, and in the cosmetics and food industries, among others. Most of the surfactants currently in the market are chemically synthetized and are produced at a very low cost, but generally they are toxic and recalcitrant compounds. Biosurfactants that are produced by microorganisms like bacteria and yeasts represent an eco-friendly alternative, but currently they occupy only a small share of the market, mainly because of challenges in their large-scale synthesis and high costs of their production. In addition to being nontoxic and biodegradable, biosurfactants have the advantage over chemically synthesized surfactants of being produced by microorganisms and thus possess biological activities such as signaling molecules and antibiotics. Thus, these chemically diverse compounds have unique potential applications based on their physicochemical properties as surfactants that are combined with different bioactivities. These unique potential applications include their use in biomedical and food industries, cosmetics, and agriculture, for example. As mentioned previously, the aim of this book is to include new insights and approaches that address different aspects of biosurfactant research and development. It is not intended to review the literature on different types of biosurfactants, but to highlight novel strategies and potential applications. We are currently in an exciting period for the field of biosurfactant research, since glycolipids such as sophorolipids produced by yeasts, rhamnolipids produced by Pseudomonas, and alkylglycosides that are semisynthetic surfactants are all available in the market, and there are many challenges to make these and other surfactants more competitive and able to fulfil specific needs. Some of the research approaches that have recently been developed to achieve this purpose are reviewed in this book, which is divided into five sections and thirteen chapters. Section I,

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Preface

“Introduction,” includes two chapters that present the general characteristics of biosurfactants, their physicochemical characteristics, and their comparison with chemically synthesized surfactants, highlighting the novel properties of these tension-active compounds produced by microorganisms, and also provide an introduction to the different research areas that are currently being pursued. Section II, “Novel Approaches for the Production and Use of Biosurfactants,” includes five chapters that focus on the presentation of different research approaches for the study of well-known biosurfactants and some of their applications, covering both traditional and novel uses. Section III, “Genetic Manipulation and the Production of Novel Biosurfactants,” includes three chapters that describe the genetic manipulation of different microorganisms to increase biosurfactant productivity or to produce molecules with improved characteristics. Section IV, “Use of Alternative Strategies for Biosurfactant Production,” includes two chapters that focus on novel strategies for biosurfactant production that are not based on the use of microorganisms. The last section, Section V “Concluding Remarks,” contains a chapter that presents the perspectives and challenges for biosurfactant research and innovation. In summary, this book represents an eclectic approach to bring to your attention the fascinating world of biosurfactant research. I hope you will enjoy traversing the sections and chapters in the company of the authors who have so enthusiastically contributed to the book. Gloria Sobero´n-Cha´vez

CHAPTER

Microbial bio-based amphiphiles (biosurfactants): General aspects on critical micelle concentration, surface tension, and phase behavior

1

Niki Baccile and Alexandre Poirier Sorbonne Universit
e, Laboratoire de Chimie de la Matie`re Condens
ee de Paris (LCMCP), UMR CNRS 7574, Paris, France

1. Introduction Surfactants are a class of chemicals applied in a vast array of applications and markets, reaching production volumes of about 20 million tons per year [1,2], with and economic weight of 43.7 billion dollars in 2017, projected to reach 66.4 billion dollars by 2025 [3]. The word “surfactant” is the contraction of “SURFace ACTive AgeNT”, indicating their ability to adsorb at interfaces, with the property of lowering the surface tension (ST) of water. This behavior is attributed to their ‘amphiphilic’ nature defined as molecules with a hydrophilic (“water-loving”) and a hydrophobic (“water-hating”) part. Due to the widespread use and applications of surfactants, research on surfactants constitutes a scientific domain of its own. Surfactants have played a decisive role in shaping the concepts of sustainability and green chemistry. Fatty acid soaps guarantee cleanliness and hygiene since time immemorial. Surfactants are involved in the environmentally friendly production of rubber, plastics, paints, and adhesives in the aqueous phase. In the field of polymer synthesis, surfactants make this possible in water, thus lowering, or even eliminating, the risks of these processes, such as fire hazards. Toxic emissions are reduced towards zero and occupational safety is increased. However, their ubiquitous use in our everyday lives also has some drawbacks. Surfactants have been associated with pollution problems, but also with dermatological issues such as skin irritation and even allergic reactions. Moreover, many of the produced surfactants are derived from petrochemical resources and associated with harsh and/or polluting production processes. Many products have already been banned for reasons of toxicity and/or pollution in the past 30 years and more are expected to follow. For these reasons, investigations aiming at finding nontoxic, benign, products, and more specifically natural bio-based alternatives to petrochemical surfactants started as a subfield in surfactant’s science since the 1960s, and developed as a field per se since the 1970s, motivated by the oil crisis and raising of oil costs [4–6]. Employment of linear alkylbenzene sulphonates and methyl ester sulphonates instead of Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00001-6 Copyright # 2023 Elsevier Inc. All rights reserved.

3

4

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

their branched counterparts, use natural fatty alcohol alternatives to synthetic alcohol polyglycolethers or sulphates; green fatty alcohol (or guerbet) alcohol polyglycol ethers, -ethersulphate, -phosphates, and -sulphosuccinate surfactants replaced alkylphenol polyglycolethers. These are some of the most common strategies employed by industry to develop more benign molecules [7,8]. The quest of more ecofriendly surfactants is then just a natural consequence of this long-date trend. Bio-based surfactants, or biosurfactants, are defined as molecules that are fully based on biomass such as sugars, plant oils, amino acids, etc. This field is characterized by two different production approaches. In the first, chemical, approach, bio-based hydrophilic and hydrophobic molecules are covalently linked through organic [9,10] chemistry. In the second, biological, approach, biosurfactants are either extracted from plants or produced through biocatalytical (use of enzymes) or microbial processes. Although the frontiers between and within these approaches are sometimes blurry, a broad community agrees on employing the word “biosurfactants” in relationship to amphiphilic surface active agents produced by a microbial fermentation process. One then speaks of “microbial biosurfactants” [4,11,12]. Research on microbial biosurfactants is known since the 1960s [13,14], but it is becoming a trendy topic since two decades, during which a large number of review papers and books have been published [11,12,15–27]. They commonly address the topic of microbial biosurfactants’ classification, the synthesis’ strategy, derivatization and genetic modification towards development of new chemistry [26,28,29], their aqueous and antimicrobial properties and their application potential in various fields [4,12,23,25–27,30–33]. The number of existing biosurfactants from microbial origin is quite impressive, as well as the number of microorganisms producing them [34]. However, only few can be produced in sufficient amount, with acceptable purity and homogeneity to be satisfactorily studied from a physicochemical point of view. Rhamnolipids (RLs), sophorolipids (SLs), cellobioselipids (CLs), mannosylerythritol lipids (MELs), and surfactin are broadly recognized as the most classical ones. Trehalolipids (TLs) are an interesting case. We are not aware of any specific study on the solution and interface properties of TLs, despite some nonnegligent work that has been done on this family of compounds since the mid-50s [35]. In the meanwhile, chemical derivatizations of existing biosurfactants [26], and more recent trends in the production of new biosurfactants from engineered strains [23,25,26,28,29,34,36–41], constitute promising alternatives to expand the biosurfactant portfolio in the future. The availability of these new compounds since less than a decade and ready collaboration between researchers across disciplines has made their advanced characterization possible. Fig. 1 summarizes the most important biosurfactants found in the literature. It also includes some derivatives, like glucolipids (GLs) or stearic acid SL. The list is far from being exhaustive, as a number of new derivatives, may them be of chemical of biotechnological origin, are produced regularly. We anticipate that, considering the latest research on the solution properties of the molecules given in Fig. 1, the word “biosurfactant” is reductive and one should rather speak of bioamphiphiles, whereas a surfactant is an amphiphile with surface active properties. In fact, most of the molecules in Fig. 1 only show surface active properties under specific conditions of pH and temperature and in some cases they do not show them at all. Addressing to them as biosurfactants only could then be erroneous in some cases. In the field of colloids science, the surfactant and lipid communities are generally distinct, although connected by many bridges. Property- and application wise, the same distinction should occurs in this field. However, one must acknowledge that the word biosurfactant is nowadays largely employed and it would be quite tedious to introduce a newer terminology. We then try to identify when the molecules in Fig. 1 behave as surfactants and when they behave as lipids.

HO OH HO O

HO HO

Bolaform amphiphile

OH

Deacetylated acidic C22:013 sophorolipid

O

OH

O

O

O OH

R Headgroup 1

Deacetylated acidic C16:0 sophorolipid

Spacer

Headgroup 2

HO HO

Headgroup 1 = or =/ Headgroup 2

HO HO

OH

OH O

HO OH HO O

O

O

O OH

OH

Deacetylated acidic C18:0 sophorolipid

O

HO OH HO O

OH

O O

CH3

O

OH OH

Sophorolipids (SL) Glucolipids (GL)

As-produced sophorolipid (SL mixture from wild type S. bombicola OAc

HO O

Acetylated lactone C18:-cis SL

O

HO OAc HO O

(Un)saturation(s) O

Acetylated acidic C18:1-cis SL

O

O

OH

HO HO

O

O

O

R1 HO HO O

(- glucose)

CH3

OH

GL

OH

R1 = OAc, H R2 = OAC, H

SL

HO HO

Deacetylated acidic C18:0- glucolipid OH

O OH

De(acetylation)

HO OH HO O

OH CH3

O

HO HO

OH

O

OH

OH CH3

Deacetylated acidic C18:1-cis glucolipid O

OH

O O

O

HO HO

General acidic sophorolipid (SL) structure

R2

O O

CH3

OH

O

CH3

OH

HO OAc HO O

OH

O

O O

Deacetylated acidic C18:1-trans SL

O O

SL

OAc

HO HO

HO HO

HO OH HO O

CH3

OH

O

Deacetylated acidic C18:1-cis SL O

O O OH

O

CH3

OH

di-rhamnolipid (di-RL)

Cellobioselipids (CL) hydrolyzed HO HO OH

OH O

O HO

OH O OH

OH

OH

O R1 = H, OH R2 = H, OH

mannosylerythritol lipids (MELs)

mono-rhamnolipid (mono-RL)

CH3

OH

R2

O R1

O O HO

O HO O

O

O

O

O O

HO HO

OH O

HO HO

FIG. 1 Most important biosurfactants found in the literature.

OH

O

O

H3C

( )n

O OR

2

( )n H2C-OH O H OH OR1 OO H OH O

O MEL-A: R1 = R2 = Ac MEL-B: R1 = Ac, R2 = H MEL-C: R1 = H, R2 = Ac MEL-D: R1 = H, R2 = H (n=6-10)

surfactin O

HO

O O

N H

NH

HN O

H

O

O

O NH O

NH H N

H N

O O O

OH

6

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

This chapter focuses on the physicochemical properties and phase behavior of microbial biosurfactants in aqueous solution within the broader context of surfactants in solution. Please note that this chapter has been adapted from a recent review article written by the authors, to which the reader can address for a more tutorial presentation of the field [42]. The chapter will end with some perspectives in colloids and materials science that are peculiar to this class of compounds.

2. Biosurfactants in solution This section recalls few major concepts of surfactants in solution, connecting properties with molecular structure. In parallel, the same ideas will be outlined for the major biosurfactants.

2.1 Hydrophilic-lipophilic balance (HLB) The hydrophilic-lipophilic balance (HLB) and hydrophilic-lipophilic difference (HLD) are two widespread approaches to forecast the emulsification ability of surfactants [43–45]. The HLB was conceived to create an empirical relationship between the surfactants’ properties [e.g., oil-in-water (o/ w) or water-in-oil (w/o) emulsifier, wetting agent or detergent] and their molecular composition, whereas the latter is generally expressed in terms of the balance of the hydrophilic and hydrophobic portions of the molecule. Initially developed for polyoxyethylene-type surfactants, HLB has been widened to a much broader class of molecules by including the contribution of specific chemical groups, which have a strong influence on the properties. Despite its astonishing simplicity, the HLB method has been employed in the surfactant industry for years and it works nicely on well-established molecules, like nonionic surfactants. Nonetheless, this method fails for a number of systems because it does not take into consideration the effect of temperature, electrolytes and ionic strength, impurities, and additives in general. Another drawback is certainly the pletora of existing methods to calculate the HLB method, which was actually by-passed by the HLD method, developed in the late 70s. HLD, developed by Salager [46–48], is much less known but it constitutes an evolution of HLB because it includes external parameters such as temperature, salinity, and the nature of the oil. In the end, both HLB and HLD revealed to be useful for few standard ionic and nonionic surfactants, but they cannot easily be generalized to complex amphiphiles, like divalent, gemini, branched or bolaform (Fig. 1) surfactants. The HLB of major surfactants is well-known. For instance Tween derivatives have HLBs between 10 and 20 while Brij from 4 to 16, depending on the length of the PEO headgroup [49]. On the contrary, few scientific publications discuss the HLB of biosurfactants and one of the latest was published on surfactin in 2005 [50]. Using the Griffin formula (HLB ¼ 20  (MWH/MWS), with MWH and MWS, respectively, being the molecular weight of the hydrophilic part and of the whole surfactant), one finds values between 6 and 13 for MELs [51–53], 21 for surfactin [50] and, on the basis of the chemical formulas, one can estimate valued contained between 5 and 15 for SLs and RLs [54]. For many biosurfactants, the properties expected according to the calculation of the HLB are in agreement with the broad range of properties experimentally observed, like o/w emulsification and detergency [55,56]. However, calculated HLB for biosurfactants can be very broad, as in the case of TLs, or the expected properties may not correspond to the value of HLB, thus generating confusion and bad expectations.

2 Biosurfactants in solution

7

Marqu
es et al. estimate an HLB of 11 for a TLs mixture so that o/w emulsion is expected, although w/o emulsion is obtained [57]. Acidic SLs are expected to be o/w emulsifiers, but in fact their bolaform nature make them poor emulsifying agents. To improve their emulsifying character, the hydrophobic character of the tail must be improved by chemical modification [58]. HLB also fails to predict the behavior in mixture of compounds with different HLBs. Some studies provide the HLB for a given biosurfactant, as reported for individual MELs, but calculated HLB fails to predict and understand the interfacial behavior of a mixture of MELs, which constitute the actual raw compound [51]. Finally, HLB becomes unsuitable to predict the behavior of polymeric and proteic biosurfactants like surfactin, because the HLB range expected by surface efficiency of surfactin [59,60] is far from the HLB calculated by emulsification method [50]. HLB of surfactin is varying with environmental conditions like pH, specific ions condensation, and temperature and it is a source of debate [61–63]. These specificities render HLB useless and require more refined understanding of the biosurfactant behavior in solution and in oil and water mixtures.

2.2 Surface tension (ST) and critical micelle concentration (CMC) The ST is a parameter of paramount importance in a number of physical phenomena like adsorption, wetting, catalysis, distillation, and much more, with direct involvement in the conception of industrial products in coating, food, detergents, cosmetics, and so on. ST is defined as the energy required to create a unit area of interphase [64] and surfactants play a crucial role in lowering the ST of water at the water-air interface from about 70 mN/m to about 25–40 mN/m. Upon mixing micromolar amounts of a surfactant in water, the water-air interface is occupied by surfactant monomers, pointing the hydrophilic headgroup towards water and the hydrophobic chain towards air. This phenomenon is at the origin of the reduction in ST and to the increase in surfactant packing at the interface [65]. When the surfactant reaches the conditions of maximum packing, it will start aggregating into spheroidal aggregates, micelles, in the bulk solution. The concentration at which aggregation occurs is called critical micelle concentration, widely known as CMC [2], and also referred to as CMC1, in opposition to CMC2, the concentration value above which micellar growth is rapidly implemented [66]. CMC is classically determined by the inflection point in ST versus concentration experiments, although many other techniques, such as turbidity, self-diffusion NMR, solubilization, pyrene fluorescence, and many others can be equally used. The typical CMC1 values for a broad set of surfactants settles in the order of the mM range, although the dispersion is broad (between 105 and 101 M) and it strongly depends on the chemical structure of the surfactant, where type of headgroup and chain length are critical parameters [67]. There are four main families of classical head-tail surfactants and they are classified on the basis of their headgroup: cationic, anionic, nonionic, and zwitterionic. Whichever the chemical nature of the head group, the CMC decreases with increasing the length of the alkyl chain, where the decrease is more pronounced for nonionics than for ionics, respectively, a factor 3 and 2 upon addition a methylene group in the aliphatic chain. The CMC values of nonionic surfactants are about two orders of magnitude lower than the values of ionic surfactants. Interestingly, among ionic surfactants, the difference in CMC is milder, with cationics having higher CMC values than anionics, while among nonionics, CMC slightly increases with bulkiness of headgroup. Other parameters have an important influence on CMC such as the valency of counterions for ionic surfactants (the higher the valency, the lower the CMC), branching, unsaturation, cosolutes. Temperature is also an important parameter, which however

8

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

has a much stronger impact on the surfactant’s solubility itself through the Kraft phenomenon. The Kraft point is defined as the temperature below which the surfactant is insoluble and above which solubility experiences an exponential increase [67]. One last remark concerns the estimation of CMC for bolaamphiphiles (bolas). Bolas have attracted a lot of attention in the past years [68], but they have been studied in a less rational manner than single chain surfactants. For these reason, to the best of our knowledge, no general experimental trend in their CMC has been reported so far. Nonetheless, Nagarajan has calculated, and compared two types of experiments; the CMC values for bolaform surfactants and found that higher values than single-head amphiphiles are expected, the second headgroup in bolas improving the monomer solubility in water. Depending on the nature of the headgroup (ionic or nonionic), he gives values in the order of 102 M [69]. ST and CMC have been extensively reviewed in the past for many biosurfactants [12,24,27,31]. However, as shown hereafter, a rationalized comparison of ST and CMC across biosurfactants is not easily possible. For this reason, the purpose of the following paragraphs is a critical overview of ST and CMC in the context of biosurfactants, rather than list of hardly-to-compare values. ST experiments on biosurfactants started already in the 60s [70–72], although thorough ST measurements were only carried out from the 80s onward, when low-molecular weight (LMW) glycolipid biosurfactants, like RLs, SLs, or TLs, appeared to have a better market potential in view of replacing petroleum-based surfactants [73–75]. In the meanwhile, constant improvements in developing both structural variety and increasing production rates contributed to promote ST studies later on [76,77]. Interestingly enough, even if some biosurfactants were discovered in the 50s, like CLs, the study of their interfacial properties only started half a century later [78,79]. Biosurfactants have similar concentration-dependent ST profiles as reported for synthetic surfactants, but the mechanism of surface stabilization depends on their molecular weight. LMW biosurfactants, like RLs, SLs, or MELs ( 1

a)

b)

FIG. 2 (A) Scheme of an amphiphile molecule and its main geometrical parameters, Ae ¼ equilibrium surface area of the headgroup, L ¼ length of the tail, V ¼ volume occupied by the tail. (B) Typical values of the packing parameter, PP, and corresponding amphiphile morphology.

2 Biosurfactants in solution

13

If the PP model could predict the morphology and phase transition of a large number of amphiphiles, it has some drawbacks. For instance, the exact interpretation of the molecular parameters has been often limited to their simple geometry, such as considering Ae as equivalent to the steric hindrance of the surfactant’s headgroup [137], while the notion of Ae is thermodynamic. The value of Ae depends on the ionic strength in ionic surfactants (the higher the ionic strength, the smaller the surface area), but also on the binding affinity of counterions [138,139], the hydration of counterions (chao/kosmotropic effects) [138,139], the hydration of the headgroup, the hydrophobicity of some headgroups. The homogeneity of the tail’s density is another underestimated parameter in the PP model [137]. Entropy is not considered, although it has an important role [140], and so on. The PP model, for instance, can neither describe microemulsions and explain the existence of a second CMC (CMC2) [141], nor explain the formation of ribbons with specific counterions [142]. Since the late 80’s, several authors have developed complementary, and even alternative, models in the effort to build a generalized theory of self-assembly: Svenson has reviewed the major ones some time ago [143] and we address the more experienced reader to the works of Eriksson (1985 and onward) [144,145], Blankenschtein (1990 and onward) [146], Nagarajan (1991 and onward) [147–149] and, more recently, Bergstr€om (2000 and onward) [150,151], who tried to build a general micelle model [141,152]. A general, less technical, discussion on the self-assembly of surfactants can be found in several books [1,2] and a more general consideration on the limits of modelling the self-assembly of amphiphiles is presented in Ref. [146]. An important remark should be done. The PP model and the structures presented in Fig. 2 help understanding the difference between a surfactant- and a lipid-like behavior. Amphiphiles forming spherical and cylindrical micelles are generally considered to be surfactants (e.g., CTAB, SDS, TWEEN, …), while amphiphiles forming bilayer vesicle or flat membranes generally fall in the description of lipids (phospholipids, sphingolipids, …). As discussed below, biosurfactants present both properties according to the molecular structure and physicochemical conditions and, in this sense, one should rather speak of bioamphiphiles with surfactant-like or (phospho)lipid-like behavior. The models to understand and predict the self-assembly properties of amphiphiles have mainly been developed on the basis of head-tail amphiphiles, for which the number of existing phase diagrams is countless, but generally following the predictions of the PP model for classical ionic surfactants. Extrapolations could induce believing that the same models should be valid for more complex amphiphiles, such as gemini surfactants or bolaamphiphiles [66], although this is not the case [143]. In fact, considering the specific structure of the latter, which often requires a multistep organic synthesis scheme, thus limiting their commercial potential, the amount of theoretical work associated to complex structures is clearly less abundant. For bolaamphiphiles, for instance, Nagarajan has shown that the theory of self-assembly and the packing parameter approach are applicable [69]. If part of his predictions are interesting and verified for some specific examples available in the literature [153], the broader amount of work published on the self-assembly of bolaamphiphiles along the years has demonstrated that these molecules have a much richer, and more complex, phase behavior [68,154], depending on a broader number of parameters, which have never been rationalized so far. In particular, many bolas have a spontaneous tendency to form semicrystalline fibers or lipid nanotubes [68,154], which are morphologies that are not predicted by any of the theoretical models. This illustrates the limits of PP model to satisfactorily describe the self-assembly of bolas in particular, and of complex amphiphiles in general [143]. In a more recent work [149], Nagarajan evokes the use of refined models or complex DNA, peptide or polyoxometallate amphiphiles (but not bolaamphiles), and Bergstr€om has shown the validity of the general micelle model

14

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

to gemini surfactants [152]. However, the amount of available data where refined models were successfully applied to a broad range of new amphiphiles is still too small to generate a trustable and generalize picture of the structure-property relationship in more advanced systems. Failure of the packing parameter approach depends on many variables [143], of which a tentative nonexhaustive list can be additional simultaneous inter- and intra-molecular interactions due to the more complex structure of the amphiphile [155]; strong binding effects of the counterions as well as counterion-induced assembly and orientation [142]; presence of stimuli-responsive chemical groups which react with physicochemical parameters like pH, light, temperature, etc. [153] kinetic effects. To the best of our knowledge, the first report on the solution phase behavior of biosurfactants can be traced back to 1987, when the aggregation of RLs was explored as a function of pH [96]. Within the same context, the first study on surfactin is published in 1995, of MELs in 2000 [127], and SLs and CLs in 2004 [156] and 2012 [157], respectively. For comparison, the self-assembly properties of lipids and surfactants are studied since the 50’s and their rationalization has occurred in the 70s with the work of Tanford [135] and Israelachvili and coworkers [136,158]. This shows that a gap of at least 10 years, but more realistically 25–30 years, exists between the development of fundamental concepts in surfactants’ science and their employment in the field of biosurfactants. Several reasons explain such gap. In particular, the chemical structure of biosurfactants is more complex than common head-tail surfactants; biosurfactants are often bolaform and in many cases they have ionizable chemical groups like carboxylic acids for most glycolipids or lipopeptides. In this regard, external stimuli like pH, ionic strength, or temperature are particularly affecting their phase behavior. This multifunctionality strongly favors additional weak interactions in the self-assembly process, such as hydrogen bond or pi-pi stacking at the same time as ionic, steric, van der Waals, and entropic forces. For this reason, straightforward predictions of the morphology of biosurfactants’ aggregates and phase behavior in water often fails. The packing parameter theory based on molecular shape can sometimes be used to explain biosurfactants’ aggregation, but it is not adapted for this class of functional compounds, for which the notion of clearly distinct hydrophilic and hydrophobic regions is sometimes dim, as clearly stated for surfactin [119]. Most of the work dealing with the self-assembly and phase behavior of biosurfactants has been carried out at concentrations below about 10 wt%, while typical phase diagrams for surfactants and lipids are studied at least up to 70%–80% and for a large range of temperatures. One of the reasons to explain this gap is the lack of a recurrent and abundant sources of biosurfactants. So far, only specialized laboratories could produce these molecules and at a purity which was hardly reproducible. For this reason, most studies have been done under semidiluted conditions. At the same time, diluted conditions are compatible with variations in the physicochemical environment, like pH or ionic strength. Below, we discuss the main results obtained in the study of the self-assembly and phase behavior of selected biosurfactants. An overview is given in Table 3, while a more extensive description is given in Ref. [42].

2.5 Micelles in solution Micelles are certainly the most common self-assembled morphology observed for classical head-tail surfactants under dilute conditions. The corresponding packing parameter for spherical micelles is 0 < P < 0.33 and for elongated, rod- until worm-like micelles, is 0.33 < P < 0.5 (Fig. 2). From a structural point of view, micelles are very well characterized by SAXS/SANS (spherical micelles: lack of q-dependency

Table 3 Phase behavior of the most important biosurfactants. Concentrations above 10 wt% for MELs are underlined. Micellar (L1)

Vesicle

Name

Sphere

Cylinder

Fiber

Nonacetyl. acidic (C18:1-cis)

10 < pH < 4, C < 5 wt%

pH 4.5, 5 < C/wt% < 20

Mix with C18:0 SL pH < 7

Acetylated acidic (C18:1-cis)

7 < pH < 2

SUV

MLV

Lamellar Flat

Condensed

Coacervate (L3)

Cubic (V2)

Ref.

[129,156, 159–162] [36,159]

Neutral (1 < C/mM < 5)

Lactonic (C18:1-cis)

[159]

Symmetrical bola C16:0 sophorosides

T > 28°C

T < 28°C

[163]

Acidic (C18:0)

pH > 7.5

7.5 < pH < 3

[160,162, 164–166]

Acidic (C16:0)

pH > 7.5

7.5 < pH < 3

[167]

Acidic (C18:3-cis)

Neutral pH

Acidic (C22:013)

pH > 7.5

Glucolipid (C18:0)

pH > 7.5

Glucolipid (C18:1-cis)

pH > 7.5

[168] pH < 7

7 < pH < 4.5) T > Tm 6.5 < pH < 7.5

7 < pH < 4.5

(pH 3) pH 6 (Lam to MLV phase change)

pH < 4

[169]

Neutral/ Acidic (pH < 7.8)

pH < 4

[160,170–172]

7 < pH < 4.5, T < Tm

pH < 4

[160,170]

Continued

Table 3 Phase behavior of the most important biosurfactants. Concentrations above 10 wt% for MELs are underlined—cont’d Micellar (L1)

Vesicle

Lamellar

Coacervate (L3)

Cubic (V2)

Name

Sphere

Cellobioselipids hydrolyzed

pH > 7.5

di-RL

pH > 6.8

pH < 7 and pH 9 (C > 20–40 mM)

pH < 6

[86,96,114,133, 173–176]

Mono-RL

pH > 6.8

pH < 7 and pH 9 (C > 20–40 mM)

pH < 6

[86,96,114,133, 173–176]

Surfactin Cyclic

pH > 7.5

Ba2+ (pH 7.5)

pH < 5.5

Cylinder

Fiber

SUV

MLV

Flat

Condensed

pH < 7

6.5 < pH < 7.5

Ref. [160]

pH < 6.5

Lα Neutral, C > ∼ 65 wt %

Ba2+ (pH 7.5)

[119,177] Neutral, CAC2 (2  105 M) < C < ∼ 55 wt%

55 < C/wt% < 65

[126–128,178]

MEL-A

Neutral, < CAC2 (2  105 M)

MEL-B

Neutral, < CAC (6  106 M)

Neutral pH, CAC (6  106 M) < C < ∼60 wt%

Lα Neutral, C > ∼60 wt %

[126–128,179– 181]

MEL-C

Neutral

Neutral pH

Lα neutral (C not defined)

[92,127,182,183]

Neutral pH C < ∼ 60 wt%

Lα Neutral, C > ∼60 wt %

[179,181,184]

MEL-D

2 Biosurfactants in solution

17

followed by 4 q-dependency of the intensity; cylindrical micelles: 1/4 I(q) dependency, always in log-log scale) but important morphological information could be obtained by cryogenic transmission electron microscopy. Micelles are also characterized by lack of birefringency using optical microscopy under crossed polarizers. The micellization process can also be studied by many other concentrationdependent experiments, like light scattering, pyrene probe solubilization, ST. Most biosurfactants form micellar solutions in their ionized form at neutral-basic pH. This is the case for SLs, RLs, and GLs but also surfactin and CLs (Table 3). Whenever studied, micelles have a classical ellipsoid of revolution shape and their cross diameter roughly corresponds to the size of the molecule, as expected for bolaamphiphiles [69]. The presence of a negatively charged group introduces strong electrostatic repulsions between adjacent molecules and induce high curvatures, thus stabilizing spheroidal objects, which is in quite good agreement with the PP model. This is also verified for aminoderived biosurfactants, but at acidic pH values, when the amine is positively charged [185]. In the case of a commercial deacetylated, lactone-free, C18:1 SLs, micellar solutions are generally observed for the neutralized compound, at acidic pH, for the COOH derivative, and at basic pH, for the aminyl derivative. We believe that this is due to the presence of polyunsaturated impurities in typical commercial, and noncommercial, batches. A more detailed overview of this aspect is given in Ref. [42].

2.6 The fiber phase Fibrillation of amphiphiles and proteins is a well-known crystallization phenomenon occurring in soft matter and responsible for many living processes, like bacterial motility through actin [186] or neuronal degeneration through the protein Tau. Fibrillation is not uncommon for LMW bola [68,187,188] and peptides-based [189] amphiphiles, and several reports show similar processes in biosurfactants. The PP model does not decribe fibrillation, because the model is based on the hypothesis of a liquid hydrocarbon core, which is not the case for crystalline objects. In terms of characterization, flat and cylindrical fibers respectively provide a q2 and q1 (log-log) dependence of the scattered intensity, I(q), in SAXS/ SANS experiments. However, polydispersity, fiber aggregation and formation of spherulites generally provide a noninteger exponent, to be rather interpreted as a fractal dimension. In any case, optical microscopy and/or cryo-TEM should be employed for a clear-cut attribution of the shape and type of twist (ribbon, helical). In the case of crystalline flat fibers, a diffraction peak should be observed at distances in the order of the size of the molecule. Unexpectedly, the fiber phase is very common for microbial amphiphiles with large headgroups. Logically, when fibers form, these compounds should then not be considered as surfactants anymore. Table 3 shows that the fiber phase is observed for saturated, C18:0 and C16:0 SLs but also hydrolyzed CLs. Deacetylated C18:1 SLs could also form this phase [156], but some doubts still exist about its origin for this compound, whether it is a real phase or coming from C18:0 SLs impurities [129]. From a structural point of view, fibers are characterized by a flat semicrystalline morphology, of which the cross-section is in the order of 10–20 nm. They are generally infinitely long, although shorter fibers and spherulites can form under appropriate conditions of synthesis [190]. Fibers are generally observed in the neutral form of the bioamphiphiles, that is at acidic pH for the COOH and basic pH for the NH2 derivatives [185], but they have been reported as a possibly thermodynamic phase even for charged compounds, in the specific case of C18:1trans SLs [166]. Symmetrical bolaform SLs, and other more exotic SLs derivatives, also seem to form fibers [163,191]. Fiber-like

18

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

structures, but rather being described by a columnar phase, have also been reported for porphyrinic SLs [192,193]. At the moment, we must acknowledge that only bioamphiphiles with large di-glucose headgroups (sophorose or cellobiose) form fibers. The actual reason is not clear, but it could be a combination of melting temperature, Tm, above RT, directional hydrogen and crystallization. Most of the fibers being twisted (ribbons or helices), one could invoke the presence of chiral centers. However, this may not be the case, and steric hindrance between adjacent molecules packed at a nonzero angle one with respect to the other could explain the twist, as proposed by others for the formation of lipid nanotubes [194].

2.7 Membranes When speaking of membranes in the field of amphiphiles, one generally refers to bilayer membranes in the form of vesicles, multilamellar vesicles, or lamellar phases, although sponge and other more complex phase can occur. In the case of microbial amphiphiles, membranes have been often observed, thus confirming that the term biosurfactant is not appropriate in this case. One should then rather employ the term bioamphiphile, as for the case of fibers. Vesicles, generally a metastable phase, are unique objects having the capability to encapsulate, transport, and deliver a cargo (hydrophobic drugs, macromolecules, nanoparticles) with strong benefit in many domains from medicine to cosmetics. Lamellar phases are of major importance in living organisms. Phospholipids, not discussed in this review, are the major component of biological membranes. Lamellar phases have different notations according to their order, hydration or lipid tilt. These phases are an important topic of research in biophysics, for the better understanding of living organisms. The packing parameter for vesicles is 0.5 < PP < 1 while for lamellar phases PP ¼ 1. As a general rule, both vesicle and lamellar phases are characterized by a typical q2 dependence of the scattered intensity in SAXS/SANS experiments. In the case of multilamellar structures, at least two diffraction peaks should be observed, one being the harmonics of the other (1:2 ratio in position). Lamellar phases with poor long-range order, or swollen systems, make exception to these rules. If the q2 dependence is generally always observed, the diffraction peaks may either be broad or even not appear at all. Polarized light microscopy, or cryo electron microscopy on diluted samples, should be generally employed as complementary techniques for a more clear-cut attribution. A warning on the notation. Vesicles and lamellar phase have a number of notations, some only speaking to experts. One could refer to the encyclopedia of colloids science for more information [195], or to Ref. [42] for a short summary within the context of bioamphiphiles. Hereafter, we report a broad overview of membrane structures found for bioamphiphiles, but a more critical discussion is given in Ref. [42]. Membranes have been observed in the shape of vesicles and within the context of bioamphiphiles since 1987, in the case of RLs at acidic pH [96]. Ever since, a number of systems does show the formation of membranes, RLs (confirmed by further studies), surfactin, C22 derivative of SLs, GLs, MELs (Table 3). May them be multilamellar vesicles or lamellar phases, membranes have been observed both under diluted and concentrated conditions, whereas the only reliable work performed at concentrations above 10 wt% has been reported for MELs [178,181,184,196]. Differently than what it is found for phospholipids, the structure of the membranes prepared from microbial bioamphiphiles is generally interdigitated, with a total membrane thickness (between 3 and 4 nm) roughly corresponding to the size of a single molecule. This is generally explained by the

2 Biosurfactants in solution

19

bolaamphiphilic nature of most of these compounds. Most of the membranes are observed when the acidic group is neutralized, generally at acidic pH below 7, or sometimes below 6. In reality, in the pH range between 4 and 7, the membrane is composed by a difficult-to-evaluate mixture of charged (COO) and uncharged (COOH) derivatives, which guarantee the low membrane curvature and the colloidal stability at the same time. It is by the way not uncommon to observe, for RLs or GLs, for instance, a precipitation in a lamellar phase at pH below 4. More insight in the vesicle and lamellar phases are given in Ref. [42].

2.8 Less common structures A number of other structures have been reported. Some of them being minor secondary structures, and others being major but unique for a given compound under specific conditions: (1) wormlike micelles have been reported for SLs, GLs, and possibly surfactin [156,160,161,177]; (2) nanoplatelets were reported for SLs, CLs, and GLss [160,162,169]; (3) a columnar phase was reported for porphyrinic derivatives of SLs [192,197,198]; (4) a coacervate phase was reported for MEL-A at low concentrations [126,128]; (5) cubic structures (Ia3d or Pn3m) have been observed at high concentration for MELs and surfactin [126,128,178,196]; (6) ill-defined structures were reported by several authors on SLs and RLs [36,159,162,199,200].

2.9 Comments The packing parameter, PP, concept has been discussed by several authors in relationship to the selfassembly of biosurfactants. It was employed to discuss, understand and tentatively explain the self-assembly properties of RLs [86,114,133,175], SLs [159,160], GLs [160], and surfactin [119,175,196]. However, a deeper look at the existing data shows that this approach, classically used for standard head-tail surfactants, does not satisfactorily explain the experimental evidence and in this regard it cannot be employed in a straightforward manner to predict the self-assembly of biosurfactants. We have estimated that in about 50% and 25% of the studies on biosurfactants, the PP model either fails or it is only partially verified [42]. Predictions of the self-assembly and phase behavior through the PP model partially works, with exceptions, for many biosurfactants under alkaline conditions. It generally fails for saturated systems under acidic conditions, while the discussion is open for surfactin, of which the role of the peptidic part, may it be linear or cyclic, is an open question. The reason why the PP theory is unreliable to explain and predict the self-assembly of most biosurfactants has been addressed. Most biosurfactants fall in the category of bolaamphiphiles, a class of compounds which has been rationalized by Nagarajan in the late ‘80s [69]. According to his work, calculation of PP for bolaamphiphiles relies on twice the headgroup surface area and half the chain length per molecule. Despite such a discrepancy, the upper and lower end-values of PP defining the shape of the aggregate do not change between one-headed and bolaamphiphiles. This fact rules out possible errors between the calculated PP and experimental morphology observed for bolaamphiphiles structures. The poor agreement between the PP theory and biosurfactants self-assembly must then be found elsewhere. A list of possible problems is given hereafter, although a more detailed description is given in Ref. [42]. (a) The hypotheses behind the PP model theory may not be respected for BS, for instance the liquid-like nature of the hydrophobic tail, especially when fibrillation occurs; (b) calculation of PP for biosurfactants may be erroneous; (c) effect of

20

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

pH is important as pH controls the amount of ionic, neutral and mixture of ionic and neutral species; (d) hydrogen bonding and sugar headgroup configuration may have an underestimated role; (e) effect of impurities and mixture of congeners, due to the fact that biosurfactants are produced by microorganisms; (f) co-existence of multiple phases; (g) fast kinetics versus thermodynamic equilibrium, especially during variations of pH or temperature; (h) strong impact of physicochemical parameters; (i) possible effects of chirality and/or bulkyness of headgroup.

2.10 Unique macroscale properties of biomphiphiles Synthetic and natural amphiphiles may have a broad range of different chemical structures, each characterized by an associated phase behavior and, eventually, macroscale properties. The general trend is that a specific family of amphiphiles is characterized by well-defined properties. For instance, surfactant wormlike micelles can form micellar hydrogels, often found in liquid soaps and shampoos. Phospholipids stabilize vesicles, useful for encapsulation, while peptide amphiphiles generally form fibers and fibrillar hydrogels. Of course, some permeability across boundaries exist, especially if these classes of molecules are mixed together. The astonishing, and actually undervaluated aspect of biological amphiphiles, is their ability to mix nano- and macroscale properties within a relatively narrow range of molecular portfolio and physicochemical conditions. This is mainly due to the combined properties of the fatty acid, sugar and free carboxylic end-group. Although the combination of these aspects is not well-understood yet, the final effects are quite surprising, with a number of unexpected macroscale behaviors. Hereafter we illustrate just a few of them. In the following, we report the case of hydrogels, although mor examples can be found in Ref. [42]. Hydrogels can be obtained by colloids, polymers, and low molecular weight gelators (LMWG). Surfactants can also form hydrogels, but generally in their worm-like micelle phase. In this regard, bioamphiphiles behave more as peptide amphiphiles than as surfactants. LMWG are a well-known class of molecules, which, upon cooling, self-assemble into fibers and stabilize a solvent into a gel. If the solvent is organic, they are referred to organogelators, while in the case of water, they are referred to as hydrogelators [201,202]. Several reports demonstrate the hydrogelling effect of microbial bioamphiphiles, like CLs and various derivatives of SLs, in their fiber phase (Fig. 3A–C) [93,157,163,190]. More interestingly, new mechanisms of gelation have been described. The strength of C18:0 SL-based hydrogels was shown to be dependent on the rate of pH variation, from basic to acidic [190], while this correlation was never reported for LMWG before. More interestingly, the C16:0 SL does not show this correlation at all, but it rather shows a dependency of the gel strength on the final pH [167], as classically reported for other LMWG [204]. An even rarer hydrogelling event has been reported for C18:0 GLs, which form very special lamellar hydrogels at less than 10 wt% in water (Fig. 3C) [172,205], where lamellar hydrogels are extremely rare in the literature and only reported for specifically modified phospholipid membranes [206,207]. Processing of lamellar hydrogels from GLs into solid foams is possible via a controlled directional freezing method. Solid lamellar foams with 3D architecture show Young moduli in the order of 20–30 kPa in both the axial and equatorial directions, explained by the isotropic orientation of the macropores within the material, which can withstand up to 1000 times its own weight (Fig. 3D). This result is particularly unexpected considering the strong anisotropic growth of ice and never reported before for materials obtained with standard amphiphiles.

2 Biosurfactants in solution

a)

b)

OAc

OAc AcO

O

O

OH

OAc O

OH HO

O HO HO

ONa

11

O

OH OH

OH

OH O

HO HO

O

O O

O O

OH

HO

G'/Pa

103

G' G" (Pa)

OH OH OH

t=0 t = 15 h

104

10 1 0.1 G' G"

G'= ACn

102 101

n= 2.6 ± 0.4

0

10

10–1 0.001 20

OH OH

Ac: CH3CO-

105

0

O

O

100

0.01

21

40

60

n= 3.9 ± 1.1 0.1

80

1 C/wt%

10

Temperature (°C)

c)

HO

OH HO OH HO O

HO HO

O O

HO HO

O

O

OH

O

104

Interdigitated layers (IL)

OH

Crystalline twisted ribbon

G', G" GC18:0 Lamellar hydrogel G', G" SLC18:0 Fibrillar hydrogel

COOH

SLC18:0

CO

OH

103

C= 5 wt% pH 6.2 ± 0.3

G', G" / Pa

GC18:0

102

101

100 10–2

d)

C = 50 mg.mL–1

5g

10 g

10–1

20 g

100

w / rad.s–1

101

50 g

102

100 g

FIG. 3 Low-molecular weight biosurfactant-only gelators in water. (A) Temperature evolution of G0 , G00 for CLs [17] hydrogels. (B) Concentration evolution of G0 , G00 for C16:0 SL-hydrogels. (C) Comparison of frequency-dependent G0 , G00 between fibrillar and lamellar hydrogels, respectively, obtained from C18:0 SLs and GLs at acidic pH [172,190,203]. (D) Comparison of the axial compression applied to solid foams prepared from fibrillar and lamellar hydrogels using the ice-templating process. (A) Reproduced from Imura T, Yamamoto S, Yamashita C, Taira T, Minamikawa H, Morita T, et al. Aqueous gel formation from sodium salts of cellobiose lipids. J Oleo Sci 2014;63(10):1005–10, copyright Japan Oil Chemical Society. (B) Reproduced from Baccile N, Van Renterghem L, Le Griel P, Ducouret G, Brennich M, Cristiglio V, et al. Bio-based glyco-bolaamphiphile forms a temperatureresponsive hydrogel with tunable elastic properties. Soft Matter 2018;14:7859–72. Available from: c with permission from The Royal Society of Chemistry. (C and D) are reproduced from Baccile N, Ben Messaoud G, Zinn T, Fernandes FM. Soft lamellar solid foams from ice-templating of self-assembled lipid hydrogels: organization drives the mechanical properties. Mater Horizons 2019;6:2073–86 with permission from The Royal Society of Chemistry.

22

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

In conclusion, the purity, homogeneity, and mixture of congeners is a limit for the understanding of the properties of biosurfactants in solution. Similarly, their double hydrophilic nature and responsitivity to pH, temperature, or ionic strength have a strong impact on their properties. Surfactant-like or phospholipid-like behaviors can be observed for their ionic and neutral forms, respectively. Many aspects remain to be understand both for their properties in solution (CMC, for instance) but, and above all, for their self-assembly and phase behavior. When these aspects will be understood and controlled, the panorama of possible applications will certainly explode in fields other than the actual ones.

References [1] Kronberg B, Holmberg K, Lindman B. Surface chemistry of surfactants and polymers. Surface chemistry of surfactants and polymers. John Wiley & Sons, Inc.; 2014. 1–479 pp. [2] Holmberg K, J€onsson B, Kronberg B, Lindman B. Surfactants and polymers in aqueous solution. John Wiley & Sons, Ltd 2002. [3] MarketsandMarketsTM. Surfactants market by application & type-Global Forecast 2021, 2020. Available from: https://www.marketsandmarkets.com/Market-Reports/biosurfactants-market-493.html. [4] Kosaric N, Sukan F. In: Kosaric N, Sukan FV, editors. Biosurfactants—Production—Properties—Applications. Surfactant. Boca Raton: CRC Press; 1993. 504 pp. [5] Sobero´n-Cha´vez G, editor. Biosurfactants—from genes to applications. Berlin Heidelberg: Springer Verlag; 2011. [6] Inamuddin A, Imran M, Prasad R, editors. Microbial biosurfactants preparation, properties and applications. Singapore: Springer Singapore; 2021. [7] Fernandez AM, Held U, Willing A, Breuer WH. New green surfactants for emulsion polymerization. Prog Org Coat 2005;53:246–55. [8] Sajna KV, H€ofer R, Sukumaran RK, Gottumukkala LD, Pandey A. White biotechnology in biosurfactants. In: Pandey A, H€ofer R, Taherzadeh M, Nampoothiri KM, Larroche C, editors. Industrial biorefineries and white biotechnology. Amsterdam, Oxford, Waltham: Elsevier; 2015. p. 499–521. [9] Von RW, Hill K. Alkyl polyglycosides—properties and applications of a new class of surfactants. Angew Chem Int Ed Engl 1998;37:1328–45. [10] Hill K., Rybinski W von, Stoll G., editors. Alkyl polyglycosides: technology, properties and applications. Weinheim—New York—Basel—Cambridge—Tokyo: VCH Verlagsgesellschaft; 1996. [11] Lang S. Biological amphiphiles (microbial biosurfactants). Curr Opin Colloid Interface Sci 2002;7 (1–2):12–20. [12] Desai JD, Banat IM. Microbial production of surfactants and their commercial potential. Microbiol Mol Biol Rev 1997;61(1):47–64. [13] Tulloch AP, Spencer JFT, Deneima MH. A new hydroxy fatty acid sophoroside from Candida bogoriensis. Can J Chem 1968;46:345–8. [14] Tulloch AP, Hill A, Spencer JFT. Structure and reactions of lactonic and acidic sophorosides of 17-hydroxyoctadecanoic acid. Can J Chem 1968;46(21):3337–51. [15] Cameotra SS, Makkar RS, Kaur J, Mehta SK. Synthesis of biosurfactants and their advantages to microorganisms and mankind. Adv Exp Med Biol 2010;672:261–80. [16] Mukherjee AK, Das K. Microbial surfactants and their potential applications: an overview. Adv Exp Med Biol 2010;672:54–64. [17] Benincasa M, Marqu
es A, Pinazo A, Manresa A. Rhamnolipid surfactants: alternative substrates, new strategies. Adv Exp Med Biol 2010;672:170–84. [18] Makkar RS, Cameotra SS, Banat IM. Advances in utilization of renewable substrates for biosurfactant production. AMB Express 2011;1(1):5.

References

23

[19] De S, Malik S, Ghosh A, Saha R, Saha B. A review on natural surfactants. RSC Adv 2015;5(81):65757–67. [20] Rodrigues LR. Microbial surfactants: Fundamentals and applicability in the formulation of nano-sized drug delivery vectors. J Colloid Interface Sci 2015;449:304–16. [21] Coelho ALS, Feuser PE, Carciofi BAM, de Andrade CJ, de Oliveira D. Mannosylerythritol lipids: antimicrobial and biomedical properties. Appl Microbiol Biotechnol 2020;104(6):2297–318. [22] Van Bogaert INAA, Saerens K, De Muynck C, Develter D, Soetaert W, Vandamme EJ. Microbial production and application of sophorolipids. Appl Microbiol Biotechnol 2007;76(1):23–34. [23] Marchant R, Banat IM. Microbial biosurfactants: challenges and opportunities for future exploitation. Trends Biotechnol 2012;30(11):558–65. [24] Morita T, Fukuoka T, Imura T, Kitamoto D. Mannosylerythritol lipids: production and applications. J Oleo Sci 2015;64(2):133–41. [25] Paulino BN, Pess^oa MG, Mano MCR, Molina G, Neri-Numa IA, Pastore GM. Current status in biotechnological production and applications of glycolipid biosurfactants. Appl Microbiol Biotechnol 2016;100 (24):10265–93. [26] Delbeke EIP, Movsisyan M, Van Geem KM, Stevens CV. Chemical and enzymatic modification of sophorolipids. Green Chem 2016;18:76–104. [27] Mnif I, Ellouz-Chaabouni S, Ghribi D. Glycolipid biosurfactants, main classes, functional properties and related potential applications in environmental biotechnology. J Polym Environ 2018;26(5):2192–206. [28] Van Bogaert INA, Buyst D, Martins JC, Roelants SLKW, Soetaert WK. Synthesis of bolaform biosurfactants by an engineered Starmerella bombicola yeast. Biotechnol Bioeng 2016;113(12):2644–51. [29] Van Renterghem L, Roelants SLKW, Baccile N, Uyttersprot K, Taelman MC, Everaert B, et al. From lab to market: an integrated bioprocess design approach for new-to-nature biosurfactants produced by Starmerella bombicola. Biotechnol Bioeng 2018;115(5):1195–206. [30] Rosenberg E, Ron EZ. High- and low-molecular-mass microbial surfactants. Appl Microbiol Biotechnol 1999;52(2):154–62. [31] Kitamoto D, Morita T, Fukuoka T, Konishi M, Imura T. Self-assembling properties of glycolipid biosurfactants and their potential applications. Curr Op Coll Interf Sci 2009;14(5):315–28. [32] Mulligan CN. Environmental applications for biosurfactants. Environ Pollut 2005;133(2):183–98. [33] Nitschke M, Costa SGVAO. Biosurfactants in food industry. Trends Food Sci Technol 2007;18(5):252–9. [34] Abdel-Mawgoud AM, Stephanopoulos G. Simple glycolipids of microbes: Chemistry, biological activity and metabolic engineering. Synth Syst Biotechnol 2018;3(1):3–19. [35] Franzetti A, Gandolfi I, Bestetti G, Smyth TJP, Banat IM. Production and applications of trehalose lipid biosurfactants. Eur J Lipid Sci Technol 2010;112(6):617–27. [36] Baccile N, Babonneau F, Banat IM, Ciesielska K, Cuvier A-S, Devreese B, et al. Development of a cradle-tograve approach for acetylated acidic sophorolipid biosurfactants. ACS Sustain Chem Eng 2017;5:1186–98. [37] Saerens KMJ, Roelants SL, Van Bogaert IN, Soetaert W. Identification of the UDP-glucosyltransferase gene UGTA1, responsible for the first glucosylation step in the sophorolipid biosynthetic pathway of Candida bombicola ATCC 22214. FEMS Yeast Res 2011;11(1):123–32. [38] Lodens S, De GM, Roelants SLKW, De MSL, Soetaert W. Transformation of an exotic yeast species into a platform organism: a case study for engineering glycolipid production in the yeast Starmerella bombicola. Methods Mol Biol 2018;1772:95–123. Braman J, editor. [39] Roelants SLKW, Ciesielska K, De Maeseneire SL, Moens H, Everaert B, Verweire S, et al. Towards the industrialization of new biosurfactants: Biotechnological opportunities for the lactone esterase gene from Starmerella bombicola. Biotechnol Bioeng 2016;113(3):550–9. [40] Ben Messaoud G, Baccile N. The biosynthetic gene cluster for sophorolipids: a biotechnological interesting biosurfactant produced by Starmerella bombicola, Molecular microbiology. vol. 88; 2019. p. 501–9. [41] Dierickx S, Remmery J, Lodens S, De Clercq V, Baccile N, Maeseneire D, et al. From beehive to bioeconomy: a critical review of recent developments and perspectives for sophorolipids and its producing organisms. Biotechnol Adv 2022;54:107788.

24

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

[42] Baccile N, Seyrig C, Poirier A, Castro SA, Roelants SLKW, Abel S. Self-assembly, interfacial properties, interactions with macromolecules and molecular modelling and simulation of microbial bio-based amphiphiles (biosurfactants). A tutorial review. Green Chem 2021;23:3842–944. [43] Griffin WC. Classification of surface-active agents by “HLB”. J Soc Cosmet Chem 1946;1:311–26. [44] Davies J.T.. Proc 2nd int congress surface activity, vol. 1. London; 1957. [45] Yamashita Y, Sakamoto K, Ohshima H, Yamashita Y, Sakamoto K. Hydrophilic-lipophilic balance (HLB): classical indexation and novel indexation of surfactant. In: Ohshima H, editor. Encyclopedia of biocolloid and biointerface science. Hoboken, New Jersey: John Wiley & Sons, Inc.; 2016. p. 570–4. [46] Salager J-LJ, Anton R, Aubry J-M, Anto´n R. Formulation des micro
emulsions par la m
ethode HLD. Tech l’ing
enieur Principes Formul 2001;(J2157):1–16. [47] Salager JL, Bourrel M, Schechter RS, Wade WH. Mixing rules for optimum phase-behavior formulations of surfactant/oil/water systems. Soc Pet Eng J 1979;19(5):271–8. [48] Vasquez E, Salager JL, Morgan JC, Schechter RS, Wade WH. Optimum formulation of surfactant-water-oil systems for minimum interfacial tension or phase behaviour. Soc Pet Eng J 1979;19:107–15. [49] Anon. The HLB system—a time saving guide to emulsfier selection. Wilmington, Delaware: ICI americas Inc.; 1976. p. 1–20. [50] Dehghan-noudeh G, Housaindokht M, Sedigeh B, Bazzaz F. Isolation, characterization and investigation of surface and hemolytic activities of a lipopeptide biosurfactant produced by Bacillus subtilis ATCC 6633. J Microbiol 2005;43(3):272–6. [51] Goossens E, Wijnants M, Packet D, Lemie`re F. Enhanced separation and analysis procedure reveals production of tri-acylated mannosylerythritol lipids by Pseudozyma aphidis. J Ind Microbiol Biotechnol 2016;43(11):1537–50. [52] Fukuoka T, Morita T, Konishi M, Imura T, Sakai H, Kitamoto D. Structural characterization and surfaceactive properties of a new glycolipid biosurfactant, mono-acylated mannosylerythritol lipid, produced from glucose by Pseudozyma antarctica. Appl Microbiol Biotechnol 2007;76(4):801–10. [53] Alimadadi N, Soudi MR, Talebpour Z. Efficient production of tri-acetylated mono-acylated mannosylerythritol lipids by sporisorium SAM20. Int J Lab Hematol 2016;38(1):42–9. [54] Burch AY, Shimada BK, Browne PJ, Lindow SE. Novel high-throughput detection method to assess bacterial surfactant production. Appl Environ Microbiol 2010;76(16):5363–72. [55] Zhao F, Han S, Zhang Y. Comparative studies on the structural composition, surface/interface activity and application potential of rhamnolipids produced by Pseudomonas aeruginosa using hydrophobic or hydrophilic substrates. Bioresour Technol 2020;295. [56] Daverey A, Pakshirajan K. Sophorolipids from Candida bombicola using mixed hydrophilic substrates: production, purification and characterization. Colloids Surfaces B Biointerfaces 2010;79(1):246–53. [57] Marqu
es AM, Pinazo A, Farfan M, Aranda FJ, Teruel JA, Ortiz A, et al. The physicochemical properties and chemical composition of trehalose lipids produced by Rhodococcus erythropolis 51T7. Chem Phys Lipids 2009;158(2):110–7. [58] Koh A, Linhardt RJ, Gross R. Effect of sophorolipid n-alkyl ester chain length on its interfacial properties at the almond oil-water interface. Langmuir 2016;32(22):5562–72. [59] Vaz DA, Gudin˜a EJ, Alameda EJ, Teixeira JA, Rodrigues LR. Performance of a biosurfactant produced by a Bacillus subtilis strain isolated from crude oil samples as compared to commercial chemical surfactants. Colloids Surf B Biointerfaces 2012;89(1):167–74. [60] Infante MR, Moses V. Synthesis and surface activity properties of hydrophobic/hydrophilic peptides. Int J Pept Protein Res 1994;43(2):173–9. [61] D
ejugnat C, Diat O, Zemb T. Surfactin self-assembles into direct and reverse aggregates in equilibrium and performs selective metal cation extraction. ChemPhysChem 2011;12(11):2138–44. [62] Liu JF, Mbadinga SM, Yang SZ, Gu JD, Mu BZ. Chemical structure, property and potential applications of biosurfactants produced by Bacillus subtilis in petroleum recovery and spill mitigation. Int J Mol Sci 2015;16(3):4814–37.

References

25

[63] Arutchelvi J, Sangeetha J, Philip J, Doble M. Self-assembly of surfactin in aqueous solution: Role of divalent counterions. Colloids Surf B Biointerfaces 2014;116:396–402. [64] Navascues G. Liquid surfaces: Theory of surface tension. Reports Prog Phys 1979;42(7):1131–86. [65] Holmberg K., J€onsson B., Kronberg B., Lindman B. Chapter 1—Introduction to surfactants. In: Surfactants and polymers in aqueous solution. John Wiley & Sons Ltd.; 2002. [66] Bergstr€om LM. A theoretical investigation of the influence of the second critical micelle concentration on the solubilization capacity of surfactant micelles. AIP Adv 2018;8(5), 055136. [67] Holmberg K, J€onsson B, Kronberg B, Lindman B. Chapter 2—Surfactant micellization. In: Surfactants and polymers in aqueous solutions. John Wiley & Sons, Ltd.; 2002. p. 39–66. [68] Fuhrhop J-H, Wang T. Bolaamphiphiles. Chem Rev 2004;104(6):2901–38. [69] Nagarajan R. Self-assembly of bola amphiphiles. Chem Eng Commun 1987;55:251–73. [70] Arima K, Kakinuma A, Tamura G. Surfactin, a crystalline peptidelipid surfactant produced by Bacillus subtilis: Isolation, characterization and its inhibition of fibrin clot formation. Biochem Biophys Res Commun 1968;31(3):488–94. [71] Zuckerberg A, Diver A, Peeri Z, Gutnick DL, Rosenberg E. Emulsifier of arthrobacter RAG-1: chemical and physical properties. Appl Environ Microbiol 1979;37(3):414–20. [72] Reisfeld A, Rosenberg E, Gutnick D. Microbial degradation of crude oil: factors affecting the dispersion in sea water by mixed and pure cultures. Appl Microbiol 1972;24(3):363–8. [73] Syldatk C, Lang S, Wray V, Witte L. Chemical and physical characterization of four interfacial-active rhamnolipids from pseudomonas spec. dsm 2874 grown on -alkanes. Zeitsch Naturforsch Sect C J Biosci 1985;40 (1–2):51–60. [74] Hommel R, Stiiwer O, Stuber W, Haferburg D, Kleber HP. Production of water-soluble surface-active exolipids by Torulopsis apicola. Appl Microbiol Biotechnol 1987;26(3):199–205. [75] Kretschmer A, Bock H, Wagner F. Chemical and physical characterization of interfacial-active lipids from Rhodococcus erythropolis grown on n-alkanes. Appl Environ Microbiol 1982;44(4):864–70. [76] Kitamoto D, Yanagishita H, Shinbo T, Nakane T, Kamisawa C, Nakahara T. Surface active properties and antimicrobial activities of mannosylerythritol lipids as biosurfactants produced by Candida antarctica. J Biotechnol 1993;29(1–2):91–6. [77] Shen L, Zhu J, Lu J, Gong Q, Jin M, Long X. Isolation and purification of biosurfactant mannosylerythritol lipids from fermentation broth with methanol/water/n-hexane. Sep Purif Technol 2019;219:1–8. [78] Lemieux RU, Thorn JA, Brice C, Haskins RH. The ustilaginales. Can J Chem 1951;29:409–14. [79] Puchkov EO, Z€ahringer U, Lindner B, Kulakovskaya TV, Seydel U, Wiese A. The mycocidal, membraneactive complex of Cryptococcus humicola is a new type of cellobiose lipid with detergent features. Biochim Biophys Acta Biomembr 2002;1558(2):161–70. [80] Uzoigwe C, Burgess JG, Ennis CJ, Rahman PKSM. Bioemulsifiers are not biosurfactants and require different screening approaches. Front Microbiol 2015;6:1–6. [81] Haba E, Abalos A, Ja´uregui O, Espuny MJ, Manresa A. Use of liquid chromatography-mass spectroscopy for studying the composition and properties of rhamnolipids produced by different strains of Pseudomonas aeruginosa. J Surfactants Deterg 2003;6(2):155–61. [82] Parra JL, Guinea J, Manresa MA, Robert M, Mercad
e ME, Comelles F, et al. Chemical characterization and physicochemical behavior of biosurfactants. J Am Oil Chem Soc 1989;66(1):141–5. [83] Gudin˜a EJ, Rodrigues AI, Alves E, Domingues MR, Teixeira JA, Rodrigues LR. Bioconversion of agroindustrial by-products in rhamnolipids toward applications in enhanced oil recovery and bioremediation. Bioresour Technol 2015;2015(177):87–93. [84] Nitschke M, Costa SGVAO, Contiero J. Rhamnolipid surfactants: an update on the general aspects of these remarkable biomolecules. Biotechnol Prog 2005;21(6):1593–600. [85] Wu LM, Lai L, Lu Q, Mei P, Wang YQ, Cheng L, et al. Comparative studies on the surface/interface properties and aggregation behavior of mono-rhamnolipid and di-rhamnolipid. Colloids Surf B Biointerfaces 2019;181:593–601.

26

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

[86] Chen ML, Penfold J, Thomas RK, Smyth TJPP, Perfumo A, Marchant R, et al. Solution self-assembly and adsorption at the air-water interface of the monorhamnose and dirhamnose rhamnolipids and their mixtures. Langmuir 2010;26(23):18281–92. [87] Kim HS, Kim YB, Lee BS, Kim EK. Sophorolipid production by Candida bombicola ATCC 22214 from a corn-oil processing byproduct. J Microbiol Biotechnol 2005;15(1):55–8. [88] Tokumoto Y, Nomura N, Uchiyama H, Imura T, Morita T, Fukuoka T, et al. Structural characterization and surface-active properties of a succinoyl trehalose lipid produced by Rhodococcus sp. SD-74. J Oleo Sci 2009;58(2):97–102. [89] Shao Z. Trehalolipids. In: S-C G, editor. Biosurfactants microbiology monographs, 20. Berlin, Heidelberg: Springer; 2010. p. 121–43. [90] Morita T, Ishibashi Y, Hirose N, Wada K, Takahashi M, Fukuoka T, et al. Production and characterization of a glycolipid biosurfactant, mannosylerythritol lipid B, from sugarcane juice by Ustilago Scitaminea NBRC 32730. Biosci Biotechnol Biochem 2011;75(7):1371–6. [91] Arutchelvi J, Doble M. Mannosylerythritol lipids: microbial production and their applications; 2010. p. 146–73. [92] Morita T, Konishi M, Fukuoka T, Imura T, Yamamoto S, Kitagawa M, et al. Identification of Pseudozyma graminicola CBS 10092 as a producer of glycolipid biosurfactants mannosylerythritol lipids. J Oleo Sci 2008;57(2):123–31. [93] Imura T, Yamamoto S, Yamashita C, Taira T, Minamikawa H, Morita T, et al. Aqueous gel formation from sodium salts of cellobiose lipids. J Oleo Sci 2014;63(10):1005–10. [94] Morita T, Ishibashi Y, Fukuoka T, Imura T, Sakai H, Abe M, et al. Production of glycolipid biosurfactants, cellobiose lipids, by Cryptococcus humicola JCM 1461 and their interfacial properties. Biosci Biotechnol Biochem 2011;75(8):1597–9. [95] Hirata Y, Ryu M, Igarashi K, Nagatsuka A, Furuta T, Kanaya S, et al. Natural synergism of acid and lactone type mixed sophorolipids in interfacial activities and cytotoxicities (a). J Oleo Sci 2009;58(11):565–72. [96] Ishigami Y, Gama Y, Nagahora H, Yamaguchi M, Nakahara H, Kamata T. The pH-sensitive conversion of molecular aggregates of rhamnolipid biosurfactant. Chem Lett 1987;16(5):763–6. [97] Amani H. Synergistic effect of biosurfactant and nanoparticle mixture on microbial enhanced oil recovery. J Surfact Deterg 2017;20(3):589–97. [98] Zhang J, Lee SH, Gross RA, Kaplan D. Surface properties of emulsan-analogs. J Chem Technol Biotechnol 1999;74(8):759–65. [99] Gurjar M, Khire JM, Khan MI. Bioemulsifier production by Bacillus stearothermophilus VR-8 isolate. Lett Appl Microbiol 1995;21(2):83–6. [100] Cirigliano MC, Carman GM. Purification and characterization of liposan, a bioemulsifier from Candida lipolytica. Appl Environ Microbiol 1985;50(4):846–50. [101] Navon-Venezia S, Zosim Z, Gottlieb A, Legmann R, Carmeli S, Ron EZ, et al. Alasan, a new bioemulsifier from Acinetobacter radioresistens. Appl Environ Microbiol 1995;61(9):3240–4. [102] Jimoh AA, Lin J. Biosurfactant:a new frontier for greener technology and environmental sustainability. Ecotoxicol Environ Saf 2019;184, 109607. [103] Kitamoto D, Isoda H, Nakahara T. Functions and potential applications of glycolipid biosurfactants—from energy-saving materials to gene delivery carriers. J Biosci Bioeng 2002;94(3):187–201. [104] Jezierska S, Claus S, Van Bogaert I. Yeast glycolipid biosurfactants. FEBS Lett 2018;592(8):1312–29. [105] Amani H, Mehrnia MR, Sarrafzadeh MH, Haghighi M, Soudi MR. Scale up and application of biosurfactant from Bacillus subtilis in enhanced oil recovery. Appl Biochem Biotechnol 2010;162(2):510–23. [106] Su WT, Chen WJ, Lin YF. Optimizing emulsan production of A. venetianus RAG-1 using response surface methodology. Appl Microbiol Biotechnol 2009;84(2):271–9. [107] Maget-Dana R, Ptak M. Interfacial properties of surfactin. J Colloid Interface Sci 1992;153(1):285–91. [108] Zdziennicka A, Ja
nczuk B. Thermodynamic parameters of some biosurfactants and surfactants adsorption at water-air interface. J Mol Liq 2017;243:236–44.

References

27

[109] Song LD, Rosen MJ. Surface properties, micellization, and premicellar aggregation of gemini surfactants with rigid and flexible spacers. Langmuir 1996;12(5):1149–53. [110] Gao Y, Yang X, Bai L, Zhang J. Preparation and physiochemical properties of disodium lauryl glucoside sulfosuccinate. J Surfactants Deterg 2014;17(4):603–8. [111] Shinoda K, Yamaguchi T, Hori R. The surface tension and the critical micelle concentration in aqueous solution of β-D-alkyl glucosides and their mixtures. Bull Chem Soc Jpn 1961;34(2):237–41. [112] Marchant R, Banat IM. Biosurfactants: a sustainable replacement for chemical surfactants? Biotechnol Lett 2012;34(9):1597–605. [113] Pathania AS, Jana AK. Improvement in production of rhamnolipids using fried oil with hydrophilic co-substrate by indigenous Pseudomonas aeruginosa NJ2 and characterizations. Appl Biochem Biotechnol 2020;191:1223–46. _ [114] Ikizler B, Arslan G, Kipcak E, Dirik C, C ¸ elenk D, Aktu glu T, et al. Surface adsorption and spontaneous aggregation of rhamnolipid mixtures in aqueous solutions. Colloids Surf A Physicochem Eng Asp 2017;519:125–36. [115] Solaiman DKY, Ashby RD, Nun˜ez A, Foglia TA. Production of sophorolipids by Candida bombicola grown on soy molasses as substrate. Biotechnol Lett 2004;26(15):1241–5. [116] Morita T, Ogura Y, Takashima M, Hirose N, Fukuoka T, Imura T, et al. Isolation of Pseudozyma churashimaensis sp. nov., a novel ustilaginomycetous yeast species as a producer of glycolipid biosurfactants, mannosylerythritol lipids. J Biosci Bioeng 2011;112(2):137–44. [117] Kowall M, Vater J, Kluge B, Stein T, Franke P, Ziessow D. Separation and characterization of surfactin isoforms produced by Bacillus subtilis OKB 105. J Colloid Interface Sci 1998;204(1):1–8. [118] Bonmatin J, Genest M, Labb
e H, Ptak M. Solution three-dimensional structure of surfactin: a cyclic lipopeptide studied by 1H-NMR, distance geometry, and molecular dynamics. Biopolymers 1994;34(7):975–86. [119] Shen H-H, Thomas RK, Chen C-Y, Darton RC, Baker SC, Penfold J. Aggregation of the naturally occurring lipopeptide, surfactin, at interfaces and in solution: an unusual type of surfactant? Langmuir 2009;25 (7):4211–8. [120] Ishigami Y, Osman M, Nakahara H, Sano Y, Ishiguro R, Matsumoto M. Significance of β-sheet formation for micellization and surface adsorption of surfactin. Colloids Surf B Biointerfaces 1995;4(6):341–8. [121] Oremusova´ J. Micellization of alkyl trimethyl ammonium bromides in aqueous solutions-part 1: critical micelle concentration (cmc) and ionization degree. Tenside Surfact Deterg 2012;49(3):231–40. [122] Ogino K, Kakihara T, Abe M. Estimation of the critical micelle concentrations and the aggregation numbers of sodium alkyl sulfates by capillary-type isotachophoresis. Colloid Polym Sci 1987;265(7):604–12. [123] Li X, Zhang G, Dong J, Zhou X, Yan X, Luo M. Estimation of critical micelle concentration of anionic surfactants with QSPR approach. J Mol Struct THEOCHEM 2004;710(1–3):119–26. [124] Berthod A, Tomer S, Dorsey JG. Polyoxyethylene alkyl ether nonionic surfactants: physicochemical properties and use for cholesterol determination in food. Talanta 2001;55(1):69–83. [125] El-Sukkary MMA, Syed NA, Aiad I, El-Azab WIM. Synthesis and characterization of some alkyl polyglycosides surfactants. J Surfact Deterg 2008;11(2):129–37. [126] Imura T, Yanagishita H, Kitamoto D. Coacervate formation from natural glycolipid: one acetyl group on the headgroup triggers coacervate-to-vesicle transition. J Am Chem Soc 2004;126:10804–5. [127] Kitamoto D, Ghosh S, Ourisson G, Nakatani Y. Formation of giant vesicles from diacylmannosylerythritols, and their binding to concanavalin A. Chem Commun 2000;10:861–2. [128] Imura T, Ohta N, Inoue K, Yagi N, Negishi H, Yanagishita H, et al. Naturally engineered glycolipid biosurfactants leading to distinctive self-assembled structures. Chem - A Eur J 2006;12(9):2434–40. [129] Dhasaiyan P, Le Griel P, Roelants S, Redant E, Van Bogaert INA, Prevost S, et al. Micelles versus ribbons: how congeners drive the self-assembly of acidic sophorolipid biosurfactants. ChemPhysChem 2017;18:643–52. [130] Otto RT, Daniel HJ, Pekin G, M€uller-Decker K, F€ urstenberger G, Reuss M, et al. Production of sophorolipids from whey. II. Product composition, surface active properties, cytotoxicity and stability against hydrolases by enzymatic treatment. Appl Microbiol Biotechnol 1999;52(4):495–501.

28

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

[131] Jana S, Kulkarni SS. Synthesis of trehalose glycolipids. Org Biomol Chem 2020;18(11):2013–37. [132] Teichmann B, Linne U, Hewald S, Marahiel MA, Bolker M, B€ olker M. A biosynthetic gene cluster for a secreted cellobiose lipid with antifungal activity from Ustilago maydis. Mol Microbiol 2007;66(2):525–33. € [133] Helvacı SS, Peker S, Ozdemir G. Effect of electrolytes on the surface behavior of rhamnolipids R1 and R2. Colloids Surf B Biointerfaces 2004;35(3–4):225–33. € [134] Ozdemir G, Peker S, Helvacı SS. Effect of pH on the surface and interfacial behavior of rhamnolipids R1 and R2. Colloids Surf A Physicochem Eng Asp 2004;234(1–3):135–43. [135] Tanford C. The hydrophobic effect: formation of micelles and biological membranes. New York: John Wiley & Sons Inc; 1973. [136] Israelachvili JN, Mitchell DJ, Ninham BW. Theory of self-assembly of hydrocarbon amphiphiles into micelles and bilayers. J Chem Soc Faraday Trans 2 1976;72:1525. [137] Blackmore ES, Tiddy GJT. Phase behaviour and lyotropic liquid crystals in cationic surfactant-water systems. J Chem Soc Faraday Trans 2 Mol Chem Phys 1988;2:1115–27. [138] Srinivasan V, Blankschtein D. Effect of counterion binding on micellar solution behavior. 1. Molecularthermodynamic theory of micellization of ionic surfactants. Langmuir 2003;19:9946. [139] Srinivasan V, Blankschtein D. Effect of counterion binding on micellar solution behavior. 2. Prediction of micellar solution properties of ionic surfactant-electrolyte systems. Langmuir 2003;19:9946. [140] Mitchell DJ, Tiddy GJT, Waring L, Bostock T, McDonald MP. Phase behaviour of polyoxyethylene surfactants with water. Mesophase structures and partial miscibility (cloud points). J Chem Soc Faraday Trans 1 Phys Chem Condens Phases 1983;79:975–1000. [141] Bergstr€om LM. Bending energetics of tablet-shaped micelles: a novel approach to rationalize micellar systems. ChemPhysChem 2007;8(3):462–72. [142] Oda R, Huc I, Schmutz M, Candau SJ, MacKintosh FC. Tuning bilayer twist using chiral counterions. Nature 1999;399:566–9. [143] Svenson S. Self-assembly and self-organization: Important processes, but can we predict them? J Dispers Sci Technol 2004;25(2):101–18. [144] Eriksson JC, Ljunggren S. Model calculations on the transitions between surfactant aggregates of different shapes. Langmuir 1990;6(5):895–904. [145] Eriksson JC, Ljunggren S, Henriksson U. A novel approach to the mechanics and thermodynamics of spherical micelles. J Chem Soc Faraday Trans 2 Mol Chem Phys 1985;81(6):833–68. [146] Puvvada S, Blankschtein D. Molecular-thermodynamic approach to predict micellization, phase behavior and phase separation of micellar solutions. I. Application to nonionic surfactants. J Chem Phys 1990;92:3710. [147] Nagarajan R, Ruckenstein E. Theory of surfactant self-assembly: a predictive molecular thermodynamic approach. Langmuir 1991;7(12):2934–69. [148] Nagarajan R. Molecular packing parameter and surfactant self-assembly: the neglected role of the surfactant tail self-assembly. Langmuir 2002;18:31–8. [149] Nagarajan R. Constructing a molecular theory of self-assembly: Interplay of ideas from surfactants and block copolymers. Adv Colloid Interface Sci 2016;244:113–23. [150] Bergstr€om M. Thermodynamics of anisotropic surfactant micelles. I. The influence of curvature free energy on the micellar size and shape. J Chem Phys 2000;113(13):5559–68. [151] Bergstr€om M. Thermodynamics of anisotropic surfactant micelles. II. A molecular interpretation of the micellar curvature free energy. J Chem Phys 2000;113(13):5569–79. [152] Bergstr€om LM. Explaining the growth behavior of surfactant micelles. J Colloid Interface Sci 2015;440:109–18. [153] Yan Y, Xiong W, Li X, Lu T, Huang J, Li Z, et al. Molecular packing parameter in bolaamphiphile solutions: adjustment of aggregate morphology by modifying the solution conditions. J Phys Chem B 2007;111 (9):2225–30. [154] Dhasaiyan P, Prasad BLV. Self-assembly of bolaamphiphilic molecules. Chem Rec 2017;17(6):597–610.

References

29

[155] Masuda M, Yoza K, Shimizu T. Polymorphism of monolayer lipid membrane structures made from unsymmetrical bolaamphiphiles. Carbohydr Res 2005;340(16):2502–9. [156] Zhou S, Xu C, Wang J, Gao W, Akhverdiyeva R, Shah V, et al. Supramolecular assemblies of a naturally derived sophorolipid. Langmuir 2004;20(19):7926–32. [157] Imura T, Kawamura D, Ishibashi Y, Morita T, Sato S, Fukuoka T, et al. Low molecular weight gelators based on biosurfactants, cellobiose lipids by Cryptococcus humicola. J Oleo Sci 2012;61(11):659–64. [158] Israelachvili JN, Ninham BW, Mitchell DJ. Theory of self-assembly of lipids and vesicles. Biochim Biophys Acta 1977;470:185–201. [159] Penfold J, Chen M, Thomas RK, Dong C, Smyth TJPP, Perfumo A, et al. Solution self-assembly of the sophorolipid biosurfactant and its mixture with anionic surfactant sodium dodecyl benzene sulfonate. Langmuir 2011;27(14):8867–77. [160] Baccile N, Selmane M, Le Griel P, Pr
evost S, Perez J, Stevens CV, et al. PH-driven self-assembly of acidic microbial glycolipids. Langmuir 2016;32(25):6343–59. [161] Baccile N, Nassif N, Malfatti L, Van Bogaert INA, Soetaert W, P
ehau-Arnaudet G, et al. Sophorolipids: a yeast-derived glycolipid as greener structure directing agents for self-assembled nanomaterials. Green Chem 2010;12(9):1564. [162] Cuvier AS, Babonneau F, Berton J, Stevens CV, Fadda GC, P
ehau-Arnaudet G, et al. Nanoscale platelet formation by monounsaturated and saturated sophorolipids under basic pH conditions. Chem - A Eur J 2015;21(52):19265–77. [163] Baccile N, Van Renterghem L, Le Griel P, Ducouret G, Brennich M, Cristiglio V, et al. Bio-based glycobolaamphiphile forms a temperature-responsive hydrogel with tunable elastic properties. Soft Matter 2018;14:7859–72. [164] Cuvier A-SS, Berton J, Stevens CV, Fadda GC, Babonneau F, INA VB, et al. pH-triggered formation of nanoribbons from yeast-derived glycolipid biosurfactants. Soft Matter 2014;10(22):3950–9. [165] Dhasaiyan P, Banerjee A, Visaveliya N, BLV P. Influence of the sophorolipid molecular geometry on their self-assembled structures. Chem Asian J 2013;8(2):369–72. [166] Dhasaiyan P, Pr
evost S, Baccile N, Prasad BLV. pH- and time-resolved in-situ SAXS study of selfassembled twisted ribbons formed by elaidic acid sophorolipids. Langmuir 2018;34:2121–31. [167] Baccile N, Ben MG, Le GP, Cowieson N, Perez J, Geys R, et al. Palmitic acid sophorolipid biosurfactant: from self-assembled fibrillar network (SAFiN) to hydrogels with fast recovery. Philos Trans A 2021;379, 20200343. [168] Dhasaiyan P, Pandey PR, Visaveliya N, Roy S, BLV P. Vesicle structures from bolaamphiphilic biosurfactants: Experimental and molecular dynamics simulation studies on the effect of unsaturation on sophorolipid self-assemblies. Chem - A Eur J 2014;20(21):6246–50. [169] Baccile N, Le Griel P, Pr
evost S, Everaert B, Van Bogaert INA, Roelants S, et al. Glucosomes: glycosylated vesicle-in-vesicle aggregates in water from pH-responsive microbial glycolipid. ChemistryOpen 2017;6:526–33. [170] Baccile N, Cuvier A-S, Pr
evost S, Stevens CV, Delbeke E, Berton J, et al. Self-assembly mechanism of pH-responsive glycolipids: micelles, fibers, vesicles, and bilayers. Langmuir 2016;32(42):10881–94. [171] Van RL, Guzzetta F, Le Griel P, Selmane M, Ben MG, Teng TTS, et al. Easy formation of functional liposomes in water using a pH-responsive microbial glycolipid: encapsulation of magnetic and upconverting nanoparticles. ChemNanoMat 2019;5:1188–201. [172] Ben Messaoud G, Le GP, Pr
evost S, Merino DH, Soetaert W, Roelants SLKW, et al. Single-molecule lamellar hydrogels from bolaform microbial glucolipids. Soft Matter 2020;16:2528–39. [173] Shin K-H, Kim K-W, Kim J-Y, Lee K-E, Han S-S. Rhamnolipid morphology and phenanthrene solubility at different pH values. J Environ Qual 2008;37(2):509–14. [174] Champion JT, Gilkey JC, Lamparski H, Retterer J, Miller RM. Electron microscopy of rhamnolipid morphology. J Colloid Interface Sci 1995;170:569–74. [175] Zdziennicka A, Krawczyk J, Ja
nczuk B. Volumetric properties of rhamnolipid and surfactin at different temperatures. J Mol Liq 2018;255:562–71.

30

Chapter 1 Microbial bio-based amphiphiles (biosurfactants)

[176] Rodrigues AI, Gudin˜a EJ, Teixeira JA, Rodrigues LR. Sodium chloride effect on the aggregation behaviour of rhamnolipids and their antifungal activity. Sci Rep 2017;7(1):1–9. [177] Shen H-HH, Lin T-WW, Thomas RK, Taylor DJFF, Penfold J. Surfactin structures at interfaces and in solution: the effect of pH and cations. J Phys Chem B 2011;115(15):4427–35. [178] Imura T, Hikosaka Y, Worakitkanchanakul W, Sakai H, Abe M, Konishi M, et al. Aqueous-phase behavior of natural glycolipid biosurfactant mannosylerythritol lipid A: sponge, cubic, and lamellar phases. Langmuir 2007;23(4):1659–63. [179] Fukuoka T, Yanagihara T, Ito S, Imura T, Morita T, Sakai H, et al. Reverse vesicle formation from the yeast glycolipid biosurfactant mannosylerythritol lipid-D. J Oleo Sci 2012;61(5):285–9. [180] Worakitkanchanakul W, Imura T, Fukuoka T, Morita T, Sakai H, Abe M, et al. Aqueous-phase behavior and vesicle formation of natural glycolipid biosurfactant, mannosylerythritol lipid-B. Colloids Surf B Biointerfaces 2008;65(1):106–12. [181] Fukuoka T, Yanagihara T, Imura T, Morita T, Sakai H, Abe M, et al. The diastereomers of mannosylerythritol lipids have different interfacial properties and aqueous phase behavior, reflecting the erythritol configuration. Carbohydr Res 2012;351:81–6. [182] Konishi M, Morita T, Fukuoka T, Imura T, Kakugawa K, Kitamoto D. Efficient production of mannosylerythritol lipids with high hydrophilicity by Pseudozyma hubeiensis KM-59. Appl Microbiol Biotechnol 2008;78(1):37–46. [183] Morita T, Konishi M, Fukuoka T, Imura T, Kitamoto D. Production of glycolipid biosurfactants, mannosylerythritol lipids, by Pseudozyma siamensis CBS 9960 and their interfacial properties. J Biosci Bioeng 2008;105(5):493–502. [184] Fukuoka T, Yanagihara T, Imura T, Morita T, Sakai H, Abe M, et al. Enzymatic synthesis of a novel glycolipid biosurfactant, mannosylerythritol lipid-D and its aqueous phase behavior. Carbohydr Res 2011;346 (2):266–71. [185] Ba AA, Everaert J, Poirier A, Le GP, Soetaert W, Roelants SLKW, et al. Synthesis and self-assembly of aminyl and alkynyl substituted sophorolipids. Green Chem 2020;22:8323–36. [186] Uhde J, Keller M, Sackmann E, Parmeggiani A, Frey E. Internal motility in stiffening actin-myosin networks. Phys Rev Lett 2004;93(26 I):1–4. [187] Iwaura R, Yoshida K, Masuda M, Yase K, Shimizu T. Spontaneous fiber formation and hydrogelation of nucleotide bolaamphiphiles. Chem Mater 2002;14(7):3047–53. [188] Graf G, Drescher S, Meister A, Dobner B, Blume A. Self-assembled bolaamphiphile fibers have intermediate properties between crystalline nanofibers and wormlike micelles: formation of viscoelastic hydrogels switchable by changes in pH and salinity. J Phys Chem B 2011;115(35):10478–87. [189] Oliveira IS, Lo M, Arau´jo MJ, Marques EF. Temperature-responsive self-assembled nanostructures from lysine-based surfactants with high chain length asymmetry: from tubules and helical ribbons to micelles and vesicles. Soft Matter 2019;15(18):3700–11. [190] Ben Messaoud G, Le Griel P, Hermida-Merino D, Roelants SLKW, Soetaert W, Stevens CV, et al. pHcontrolled self-assembled fibrillar network (SAFiN) hydrogels: evidence of a kinetic control of the mechanical properties. Chem Mater 2019;31:4817–30. [191] Baccile N, Delbeke EIP, Brennich M, Seyrig C, Everaert J, Roelants SLKW, et al. Asymmetrical, symmetrical, divalent and Y-shaped (bola)amphiphiles: the relationship between molecular structure and selfassembly in amino derivatives of sophorolipid biosurfactants. J Phys Chem B 2019;123:3841–58. [192] Mekala S, Peters KC, Singer KD, Gross RA. Biosurfactant-functionalized porphyrin chromophore that forms: J-aggregates. Org Biomol Chem 2018;16(39):7178–90. [193] Vermathen M, Marzorati M, Bigler P. Self-assembling properties of porphyrinic photosensitizers and their effect on membrane interactions probed by NMR spectroscopy. J Phys Chem B 2013;117(23):6990–7001. [194] Shimizu T, Masuda M, Minamikawa H. Supramolecular nanotube architectures based on amphiphilic molecules. Chem Rev 2005;105(4):1401–44.

References

31

[195] Tadros T. Encyclopedia of colloid and interface science. In: Encyclopedia of colloid and interface science. Berlin Heidelberg: Springer-Verlag; 2013. [196] Imura T, Ikeda S, Aburai K, Taira T, Kitamoto D. Interdigitated lamella and bicontinuous cubic phases formation from natural cyclic surfactin and its linear derivative. J Oleo Sci 2013;62(7):499–503. [197] Peters KC, Mekala S, Gross RA, Singer KD. Cooperative self-assembly of helical exciton-coupled biosurfactant-functionalized porphyrin chromophores. ACS Appl Bio Mater 2019;2(4):1703–13. [198] Peters KC, Mekala S, Gross RA, Singer KD. Chiral inversion and enhanced cooperative self-assembly of biosurfactant-functionalized porphyrin chromophores. J Mater Chem C 2020;8(14):4675–9. [199] Baccile N, Babonneau F, Jestin J, Pehau-Arnaudet G, Van Bogaert I, P
ehau-Arnaudet G, et al. Unusual, pHinduced, self-assembly of sophorolipid biosurfactants. ACS Nano 2012;6(6):4763–76. [200] Dahrazma B, Mulligan CN, Nieh MP. Effects of additives on the structure of rhamnolipid (biosurfactant): a small-angle neutron scattering (SANS) study. J Colloid Interface Sci 2008;319(2):590–3. [201] Draper ER, Adams DJ. Low-molecular-weight gels: the state of the art. Chem 2017;3(3):390–410. [202] Du X, Zhou J, Shi J, Xu B. Supramolecular hydrogelators and hydrogels: from soft matter to molecular biomaterials. Chem Rev 2015;115(24):13165–307. [203] Baccile N, Ben Messaoud G, Zinn T, Fernandes FM. Soft lamellar solid foams from ice-templating of selfassembled lipid hydrogels: organization drives the mechanical properties. Mater Horizons 2019;6:2073–86. [204] Raeburn J, Pont G, Chen L, Cesbron Y, L
evy R, Adams DJ. Fmoc-diphenylalanine hydrogels: Understanding the variability in reported mechanical properties. Soft Matter 2012;8(4):1168–74. [205] Ben Messaoud G, Le GP, Merino DH, Baccile N. Effect of pH, temperature and shear on the structureproperty relationship of lamellar hydrogels from microbial glycolipid probed by in-situ rheo-SAXS. Soft Matter 2020;16:2540–51. [206] Warriner HE, Idziak SH, Slack NL, Davidson P, Safinya CR. Lamellar biogels: fluid-membrane-based hydrogels containing polymer lipids. Science 1996;271(5251):969–73. [207] Niu J, Wang D, Qin H, Xiong X, Tan P, Li Y, et al. Novel polymer-free iridescent lamellar hydrogel for twodimensional confined growth of ultrathin gold membranes. Nat Commun 2014;5:3313.

CHAPTER

New insights in biosurfactants research

2

Gloria Sobero´n-Cha´veza, Martı´n P. Soto-Acevesa,b, and Luis Servı´n-Gonza´leza a

Departamento de Biologı´a Molecular y Biotecnologı´a, Instituto de Investigaciones Biom
edicas, Universidad Nacional Auto´noma de M
exico, Ciudad Universitaria, CDMX, Mexico, bDepartment of Microbiology, University of Washington, Seattle, WA, United States

1. Introduction This chapter is not intended as an extensive review of the field of biosurfactants (BS) research but instead pretends to highlight some recent and interesting research results obtained using different microorganisms and experimental approaches in the study of BS. The BS research field is a rather new scientific area, which is still not very large but is growing vigorously. Publication of the first article referring to BS dates to 1979 as consigned in PubMed, while 40 years later 301 articles were published, in 2020 there were 345 publications and by October of 2021, 333 articles were consigned (Fig. 1). However, a problem in the field of BS research is that many publications do not have the required rigor to constitute a solid contribution mainly due to the use of unsuited methods and techniques [1]. Therefore, reports that provide standardized techniques and protocols that do not rely on sophisticated equipment, like the quantification of BS using the solubilization of a dye, for example [2], can be a driving force to the advancement of this field. BS reached the market about 20 years ago when sophorolipids (SLs) produced by the yeast Starmerella bombicola began to be commercialized, but they still represent a small share of the surfactant market. To increase BS commercialization, it is important to innovate in all aspects of BS research, from describing new microorganisms that produce BS and developing genetically engineered ones to identifying new molecules, novel applications and improving scale-up and downstream processes to produce BS. Some recent examples of these types of reports will be presented here. The best-studied BSs are the glycolipids known as rhamnolipids (RLs) that are naturally produced by the opportunistic pathogenic bacterium Pseudomonas aeruginosa [3], the SL produced by the yeasts S. bombicola, Pseudohyphozyma, and other fungi [4], as well as lipopeptides like surfactin produced by several Bacillus species [5] and other lipopeptides produced by Pseudomonas [6]. In addition to producing SL, some fungal species produce other glycolipids like mannosylerythritol lipids that are also the focus of current research [7].

Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00002-8 Copyright # 2023 Elsevier Inc. All rights reserved.

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Chapter 2 New insights in biosurfactants research

FIG. 1 Number of articles related to biosurfactants published per year (PubMed October 2021).

RL and SL are currently in the market and are primarily used in the petrochemical industry, for the bioremediation of different pollutants, in household products, agricultural chemicals, and personal care products [8]. In this chapter, some recent publications will be highlighted to provide insights into novel approaches and findings related to these traditionally studied BS and the microorganisms that produce them. Some publications that represent innovations in this field will also be covered. Thus, it is aimed to provide the reader with a brief overview of the exciting BS research field.

2. Novel and traditional BS and their producers revisited As already stated above, RLs are some of the best-studied BSs which have been in the market for several years. However, there are still important challenges that drive the research on these molecules to attain their widespread use in all the industries where they have a potential application. It has been recently reported that RL can be produced anaerobically using glycerol as substrate avoiding the problem of excessive foam formation [9], which is one of the drawbacks of BS large-scale production. These BS are naturally produced by P. aeruginosa, which is the most efficient RL producer [3], and by some Burkholderia species [10], but with low productivity [11].

2 Novel and traditional BS and their producers revisited

35

The main drawback for the use of P. aeruginosa strains to produce RL at a large scale is their pathogenicity, but there have been reports on the construction of nonvirulent P. aeruginosa RLhyperproducing strains [12–14]. In the case of Burkholderia thailandensis, the molecular genetics of its RL production has started to be described [11], which represents an important precedent for further development of RL-hyperproducing strains. It has been reported recently that the production of this BS is inversely regulated, at the transcriptional level, with the production of the fatty acid polymer polyhydroxyalkanoates (PHA) by ScmR [15]. RL and PHA production use the same fatty acids as precursors, so this result with B. thailandensis opens the door to develop a metabolic engineering strategy to redirect the carbon flux to RL synthesis as has been done with P. aeruginosa RL-hyperproducing strains [13]. Occasionally strains not belonging to P. aeruginosa or Burkholderia that produce RL have been isolated [9]. In particular, some marine bacteria have been reported to be RL producers [16,17]. A special mention for its relatively high production and stability is deserved by the nonpathogenic marine bacterium Marinobacter sp. MCTG107b [18], which might be a good alternative for large-scale RL production. It is also noteworthy that the deep-sea Gram-positive bacterium Dietzia maris As-133 was reported to produce RL with two rhamnose moieties (di-RL) and to be a good oil degrader [19]. The genus Planococcus also corresponds to BS-producing marine bacteria, that can synthesize both glycolipids and exopolysaccharides BS and has been proposed as an important source for developing strains that can be used to produce these tensio-active compounds commercially [20]. Another niche where it has been reported that BS producers are enriched is soil contaminated with crude oil [21,22]. For example, D. maris AURCCBT01 (which belongs to the same species as the deepsea strain) isolated from a crude oil-contaminated soil was shown to produce an uncharacterized BS and to efficiently degrade n-alkanes [21]. The structurally heterogeneous BS produced by lactic acid bacteria (LAB) including the genera Lactobacillus and Pediococcus, have remarkable antimicrobial and antibiofilm properties [22–25] and therefore have a great potential for application in the food and healthcare industries, where LAB have been traditionally used. In addition, an approach using the metagenomic analysis of a consortium isolated from a production water sample from an oil reservoir and selected using crude oil as the carbon source showed that genes involved in BS production were enriched [26]. The predominant members of this consortium were Brevibacillus strains [26], which are spore-forming Gram-positive bacteria with high agroecology significance as potential plant growth-promoting rhizobacteria, biocontrol agents against plant diseases, and soil bioremediation agents for the removal of toxic heavy metals [27]. With respect to lipopeptide BS, there have been reports of bacteria producing them which are different from Bacillus [5] and Pseudomonas [28], the traditional producers of these compounds, such as the alpha-proteobacterium Roseomonas cervicalis [29]. Other recent reports have highlighted some novel aspects of the lipopeptide BS surfactin produced by Bacillus. For example, it was shown that surfactin produced by Bacillus velezensis H2O-1 efficiently maintains its interfacial properties in extreme conditions such as postsalt and presalt oil reservoirs [30], and the conditions to anaerobically produce surfactin were reported, avoiding the problem of foam production during fermentation [31]. It has been proposed that surfactin plays an important role in horizontal gene transfer in B. subtilis suggesting a role of this BS in the evolution of this species [32].

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Chapter 2 New insights in biosurfactants research

3. Biocatalysis, chemical, and genetic engineering strategies in BS research Carbohydrate fatty acid esters are nonionic BS produced by enzymatic synthesis that are currently used as low caloric sweeteners, and as additives in the food, pharmaceutical, and cosmetic industries [33]. Different approaches, like the overproduction of enzymes used for their synthesis and the use of protein engineering, are being followed to increase yield and to obtain molecules with improved characteristics [34,35]. The case of the surfactin family of lipopeptide BS produced by Bacillus represents an exciting and remarkable example of research and innovation which has been thoroughly reviewed recently [5]. These BSs were first reported in 1969 and their biosynthetic pathway has been described for several years. The peptide part of molecules of the surfactin family of BS is synthesized by a modular multiprotein complex belonging to the nonribosomal peptide synthetases. Since any of 15 isoforms of lipids (C12–C17) can be incorporated into a particular surfactin, each surfactin molecule produced by a particular Bacillus strain has a special structure that determines its physicochemical characteristics and potential applications. Surfactin variants with an altered structure have been produced, for example, by chemical modification of the natural BS synthesized by Bacillus [36] or by genetic engineering strategies that alter some modules of the nonribosomal peptide synthetase gene cluster [37]. Semisynthetic SL derivatives with improved characteristics have been produced using these glycolipids produced by the yeast S. bombicola as substrates for chemical modification [38]. In addition, S. bombicola has been genetically modified to produce new SL derivatives with modified structures and physicochemical characteristics [39–41]. The isolation of SL-producing halotolerant yeast from extreme environments has been recently reported [42], raising the possibility of using genes coding for more robust enzymes in the synthesis of this BS. An alternative to the use of P. aeruginosa for RL production has been the heterologous production of these BS using the nonpathogenic Pseudomonas putida strain KT2440 [43]. This heterologous system for RL production relies on the induction of the rhlAB operon of P. aeruginosa that encodes the enzymes to produce RL with one rhamnose moiety (mono-RL) and has been subject to multiple genetic modifications to develop derivatives with improved characteristics for large scale production of this BS. In addition, other microorganisms besides P. putida have been used for the heterologous production of RL [44]. As stated earlier, one of the main problems for large-scale BS production is excessive foaming during fermentation that causes loss of biomass [45]. To address this problem, three strategies based on genetic modification of P. putida KT2440, which produces RL, were recently reported [46–48]. In one case, a derivative that efficiently used ethanol (a solvent that reduces foaming) as the carbon source was selected using experimental evolution. The resulting strain showed increased RL production in an ethanol fed-batch fermentation with less foam produced [46,47]. An alternative strategy consisted of the stable integration of cassettes encoding the RL biosynthetic genes in the P. putida KT2440 chromosome and the redirection of the carbon flow towards the production of this BS by deleting genes involved in flagella and PHA synthesis; RL production by this genetically modified P. putida KT2440 derivative was improved by using a fermentation process in which the foam produced was recycled [48]. The third strategy consisted of fractionation of the foam to separate RL and bacteria from the

4 Novel applications of BS

37

culture medium; mutants with deletion of genes encoding hydrophobic membrane proteins were constructed to reduce the partitioning of cells to the gas-liquid interphase since it was shown that biomass enrichment in the foam of a derivative that does not produce flagella is reduced by 46% compared to the original P. putida KT2440 strain [49]. Expression of the P. aeruginosa rhlAB operon is posttranscriptionally thermoregulated by a ROSEtype RNA thermometer causing reduced RL production at 30°C, compared to its production at 37°C where the RNA thermometer is in an open conformation enabling translation initiation [50]. The impact on RL production of this ROSE-type RNA thermometer located in the 50 UTR of rhlAB was evaluated in P. putida KT2440 cultured at 30°C and 37°C. It was found that the slight increment in production of this BS at 37°C resulted from the increased metabolic rate at this temperature and not by the presence of the RNA thermometer [51]. However, the Salmonella 4U-RNA thermometer placed upstream of rhlAB was successfully used recently to increase RL production in P. putida KT2440, controlling its production during fermentation; cells were grown at 25°C maintaining rhlAB expression “closed” and, when the desired biomass was reached, the temperature was shifted to 38°C and RL production was induced [52].

4. Novel applications of BS The application of a particular BS is dependent on its physicochemical characteristics and the requirements for each specific case. For example, in their application in biomedical products such as pharmaceuticals and in therapeutics, where their antimicrobial, antibiofilm, anticancer, or drug delivery abilities are important [22], high purity and biocompatibility with other compounds are required. While regulations for BS applications in the cosmetic industry are not very strict, high purity standards are required [53]. On the other hand, for their application in oil recovery and industry, large quantities of these compounds are needed for increasing their efficiency; furthermore, they must show low environmental toxicity and should be preferably biodegradable [54]. BS have been applied in bioremediation and the biodegradation and detoxification of industrial effluents for several years. However, there is no rule about which technology and BS should be used. Selection of the most appropriate remediation technology must correspond with the environmental characteristics of the site and the BS should be selected based on the characteristics of the pollutant, effectiveness, regulation, and time constraints, among other factors [55]. It is also important to consider the possibility of using renewable or waste substrates and the possibility of producing BS in situ. Some recent reports have addressed the use of BS for the bioremediation of contamination by petroleum oils [56–59]. The toxicity and product formulation using the BS produced by Bacillus cereus UCP1615 for the treatment of motor oil contamination of sand and rock walls was reported [56], while a P. aeruginosa strain (ENO14-MH271625) was shown to degrade up to 73% of crude oil from soil in laboratory conditions while producing uncommon RL congeners [57]. Candida tropicalis UCP0996 produced a glycolipid BS when grown using molasses, residual frying oil, and corn steep liquor that could remove up to 66% of the motor oil adsorbed in marine stones [58], while Candida lipolytica produced a BS using a similar culture medium that removed motor oil from soil under static conditions [59]. RL have been reported to be useful for the treatment of arsenic-containing wastes [60].

38

Chapter 2 New insights in biosurfactants research

BS have been reported to be useful in industrial settings. For example, they have been used to prevent biofouling [61], and RL have been reported to be useful for the fungal elimination of toluene vapor in a biotrickling filter under stressed operational conditions [62]. Recently, the use of surfactants in paper-making industry has been reviewed, and the importance of substituting chemically synthetized surfactants with “green surfactants”, not only BS, was highlighted [63]. The application of BS in healthcare-related applications is a vigorous research field [22,64–74]. The glycolipopeptides BS produced by Lactobacillus have been reported to have antimicrobial properties that can eliminate skin pathogens [64,65] and Chlamydia trachomatis elementary bodies [66], while they have also been shown to be useful for the elimination of biofilms using a microfluidic approach [67], and of removing pathogens from PDMS-based implants [68]. SL are active against Staphylococcus aureus catheter-related infections [69]. A lipopeptide BS produced by Acinetobacter junii has been reported as useful in wound healing when applied as a hydrogel presentation [70]. In the case of RL, they have been proposed as natural anticancer agents and autophagy inhibitors [71]. The potential use of BS in the context of the present coronavirus disease 2019 (COVID-19) pandemic has been recently reviewed [72,73] and RL have been recently reported as useful hand sanitizers, capable of eliminating SARS-CoV-2 and antibiotic-resistant bacteria [74]. The use of BS in agriculture shows great potential. One of their applications relates to their antimicrobial properties that are effective against many plant pathogens [75–79]. In the case of RL and lipopeptide BS, it has been reported that they are useful for eliminating plant pathogens not only due to their antimicrobial activity but also by modulating plant immune response [76]. SL have been shown to have antimicrobial activity against phytopathogens of cherry tomatoes [77] and different fungal and oomycete plant pathogens [78]. It has recently been reported that RL and their 3-hydroxyalkanoate precursor act as signals that can be detected by Arabidopsis thaliana through independent pathways which modify its immune response [79]; the transcriptome of A. thaliana seedlings and plants in the presence of mono-RL is available [80]. Thus, it seems that these BS can act as signals that establish bacteria-plant communication. Other potential applications related to agriculture are just starting to be envisioned. It has been recently reported, for example, that RL promote anaerobic co-digestion of excess sludge and plant waste [81]. An outstanding example of RL activity that could be useful in agriculture is their ability to enhance the nitrogen fixation activity of the soil bacterium Azotobacter chroococcum [82].

5. Concluding remarks In this chapter, we have included some examples of exciting recent reports that aim to give a wide picture of the dynamism and importance of the BS research field. The study of the diverse tensio-active molecules, the microorganisms that produce them and the conditions for their large-scale production, among other themes, presents important challenges that are being addressed, so we will continue learning about these remarkable molecules and their applications (Fig. 2).

References

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FIG. 2 Some advantages and challenges of biosurfactants-related research.

References [1] Twigg MS, Baccile N, Banat IM, D
eziel E, Marchant R, Roelants S, Van Bogaert INA. Microbial biosurfactant research: time to improve the rigour in the reporting of synthesis, functional characterization and process development. Microb Biotechnol (MBT) 2021;14(1):147–70. [2] Kubicki S, Bator I, Jankowski S, Schipper K, Tiso T, et al. A straightforward assay for screening and quantification of biosurfactants in microbial culture supernatants. Front Bioeng Biotechnol 2020;8:958. [3] Sobero´n-Cha´vez G, Gonza´lez-Valdez A, Soto-Aceves MP, Cocotl-Yan˜ez M. Rhamnolipids produced by Pseudomonas: from molecular genetics to the market. Microbial Biotechnol (MBT) 2021;14(1):136–46. [4] Da Silva AF, Banat IM, Giachini AJ, Robl D. Fungal biosurfactants, from nature to biotechnological product: bioprospection, production and potential applications. Bioprocess Biosyst Eng 2021;44:2003–34. [5] Th
eatre A, Cano-Prieto C, Bartolini M, Laurin Y, Deleu M, et al. The surfactin-like lipopeptides from Bacillus sp: natural biodiversity and synthetic biology for a broader application range. Front Bioeng Biotechnol 2021;9:118. [6] Biniarz P, Henkel M, Hausmann R, Łukaszewicz M. Development of a bioprocess for the production of cyclic lipopeptides pseudofactins with efficient purification from collected foam. Front Bioeng Biotechnol 2020;8, 565619. [7] Beck A, Haitz F, Their I, Siems K, Jakupovic S, et al. Novel mannosylerythritol lipid biosurfactant structures from castor oil revealed by advanced structure analysis. J Ind Microbiol Biotechnol 2021;48:kuab042. [8] Sekhon Randhawa KK, Rahman PK. Rhamnolipid biosurfactants—past, present and future scenario of global market. Front Microbiol 2014;5:454. [9] Zhao F, Wu Y, Wang Q, Zheng M, Cui Q. Glycerol or crude glycerol as substrates make Pseudomonas aeruginosa achieve anaerobic production of rhamnolipids. Microbial Cell Fact 2021;20:185. [10] Toribio J, Escalante AE, Sobero´n-Cha´vez G. Production of rhamnolipids in bacteria other than Pseudomonas aeruginosa. European J Lipid Sci Technol 2010;112:1082–7. [11] Dubeau D, D
eziel E, Woods DE, L
epine F. Burkholderia thailandensis harbors two identical rhl gene clusters responsible for the biosynthesis of rhamnolipids. BMC Microbiol 2009;9:263.

40

Chapter 2 New insights in biosurfactants research

[12] Grosso-Becerra MV, Gonza´lez-Valdez A, Granados-Martı´nez M-J, Morales E, Servı´n-Gonza´lez L, M
endez J-L, Delgado G, Morales-Espinosa R, Ponce-Soto G-Y, Cocotl-Yan˜ez M, Sobero´n-Cha´vez G. Pseudomonas aeruginosa ATCC 9027 is a non-virulent strain suitable for mono-rhamnolipids production. Appl Microbiol Biotech 2016;100(23):9995–10004. [13] Guti
errez-Go´mez U, Soto-Aceves MP, Servı´n-Gonza´lez L, Sobero´n-Cha´vez G. Overproduction of rhamnolipids in Pseudomonas aeruginosa PA14 by redirection of the carbon flux from polyhydroxyalcanoate synthesis and overexpression of the rhlAB-R operon. Biotechnol Lett 2018;40(11):1561–6. [14] Guti
errez-Go´mez U, Sobero´n-Cha´vez G. M
etodo para la construccio´n de cepas del g
enero Pseudomonas para disminuir su virulencia e incrementar su produccio´n de ramnolı´pidos, y productos obtenidos con el mismo. Mexican patent submission MX/a/2019/006840; June 2019. [15] Martinez S, Humery A, Groleau M-C, D
eziel E. Quorum sensing controls both rhamnolipid and polyhydroxyalkanoate production in Burkholderia thailandensis through ScmR regulation. Front Bioeng Biotechnol 2020;8:1033. [16] Twigg M, Tripathi L, Zompra K, Salek K, Irorere V, Gutierrez T, et al. Surfactants from the sea: rhamnolipid production by marine bacteria. Access Microbiol 2019;1:192. [17] Tripathi L, Irorere VU, Marchant R, Banat IM. Marine derived biosurfactants: a vast potential future resource. Biotechnol Lett 2018;40:1441–57. [18] Tripathi L, Twigg MS, Zompra A, Salek K, Irorere VU, et al. Biosynthesis of rhamnolipid by Marinobacter species expands the paradigm of biosurfactant synthesis to a new genus of marine microflora. Microb Cell Fact 2019;18:164. [19] Wang W, Bobo C, Zongze S. Oil degradation and biosurfactant production by the deep-sea bacterium Dietzia maris As-13-3. Front Microbiol 2014;5:711. [20] Waghmode S, Suryavanshi M, Sharma D, Satpute SK. Planococcus species—an imminent resource to explore biosurfactants and bioactive metabolites for industrial applications. Front Bioeng Biotechnol 2020;8:996. [21] Venil CK, Malathi M, Devi PR. Characterization of Dietzia maris AURCCBT01 from oil-contaminated soil for biodegradation of crude oil. 3 Biotech 2021;11(6):291. [22] Ceresa C, Fracchia L, Fedeli E, Porta C, Banat IM. Recent advances in biomedical, therapeutic and pharmaceutical applications of microbial surfactants. Pharmaceutics 2021;13:466. [23] Patel M, Siddiqui AJ, Hamadou WS, Surti M, Awadelkareem AM, et al. Inhibition of bacterial adhesion and antibiofilm activities of a glycolipid biosurfactant from Lactobacillus rhamnosus with its physicochemical and functional properties. Antibiotics 2021;10:1546. [24] Sharma V, Singh D, Manzoor M, et al. Characterization and cytotoxicity assessment of biosurfactant derived from Lactobacillus pentosus NCIM 2912. Braz J Microbiol 2021; [Published online November 8]. [25] Adnan M, Siddiqui AJ, Hamadou WS, Ashraf SA, et al. Functional and structural characterization of Pediococcus pentosaceus-derived biosurfactant and its biomedical potential against bacterial adhesion, quorum sensing, and biofilm formation. Antibiotics 2021;10:1371. [26] Arau´jo WJ, Oliveira JS, Arau´jo SCS, Minicelli CF, et al. Microbial culture in minimal medium with oil favours enrichment of biosurfactant producing genes. Front Bioeng Biotechnol 2020;8:962. [27] Ray S, Patel N, Amin D. Brevibacillus (Chapter 9). In: Amaresan N, Senthil Kumar M, Annapurna K, Kumar K, Sankaranarayanan A, editors. Beneficial microbes in agro-ecology. Academic Press; 2020. [28] Chauhan V, Dhiman V, Kanwar SS. Combination of classical and statistical approach to enhance the fermentation conditions and increase the yield of lipopeptide(s) by Pseudomonas sp. OXDC12: its partial purification and determining antifungal property. Turk. J Biol 2021;45:695–710. [29] Mukherjee AK, Chanda A, Mukherjee I, Kumar P. Characterization of lipopeptide biosurfactant produced by a carbazole degrading bacterium Roseomonas cervicalis: the role of biosurfactant in carbazole solubilization. J Appl Microbiol 2021. https://doi.org/10.1111/JAM.15258 [published online August 20].

References

41

[30] Guimara˜es CR, Pasqualino IP, de Sousa JS, Sousa Nogueira FC, Seldin L, et al. Bacillus velezensis H2O-1 surfactin efficiently maintains its interfacial properties in extreme conditions found in post-salt and pre-salt oil reservoirs. Colloids Surf B: Biointerfaces 2021;208, 112072. [31] Hoffman M, Fernandez Cano Luna SD, Xiao S, Stegem€ uller L, Rief K, et al. Towards the anaerobic production of surfactin using Bacillus subtilis. Front Bioeng Biotechnol 2020;8:1306. [32] Danevcic T, Dragosˇ A, Spacapan M, Stefanic P, Dogsa I, Mandic-Mulec I. Surfactin facilitates horizontal gene transfer in Bacillus subtilis. Front Microbiol 2021;12, 657407. [33] Chang SW, Shaw JF. Biocatalysis for the production of carbohydrate esters. New Biotechnol 2009;26(34):109–16. [34] Kovalenko G, Perminova L, Beklemishev A. Heterogeneous biocatalytical esterification by recombinant Thermomyces lanuginosus lipase immobilized on macroporous carbon aerogel. Catal Today 2021;379:36–41. [35] Intasian P, Prakinee K, Phintha A, Trisrivirat D, Weeranoppanant N, Wongnate T, Chaiyen P. Enzymes, in vivo biocatalysis, and metabolic engineering for enabling a circular economy and sustainability. Chem Rev 2021;121(17):10367–451. [36] Shao C, Liu L, Gang H, Yang S, Mu B. Structural diversity of the microbial surfactin derivatives from selective esterification approach. Int J Mol Sci 2015;16:1855–72. [37] KAJ B, Linck A, Tietze A, Kranz J, Wesche F, Nowak S, et al. Modification and de novo design of nonribosomal peptide synthetases using specific assembly points within condensation domains. Nat Chem 2019;11:653–61. [38] Baccile N, Delbeke EIP, Brennich M, Seyrig C, Everaert J, et al. Asymmetrical, symmetrical, divalent, and Y-shaped (bola)amphiphiles: the relationship between the molecular structure and self-assembly in amino derivatives of sophorolipid biosurfactants. J Phys Chem B 2019;123(17):3841–58. [39] Dierickx S, Castelein M, Remmery J, De Clercq V, Lodens S, et al. From bumblebee to bioeconomy: recent developments and perspectives for sophorolipid biosynthesis. Biotechnol Adv 2021. https://doi.org/10.1016/ j.biotechadv.2021.107788 [published online June 21]. [40] Van Bogaert INA, Buyst D, Martins JC, Roelants SLKW, Soetaert WK. Synthesis of bolaform biosurfactants by an engineered Starmerella bombicola yeast. Biotechnol Bioeng 2016;113:2644–51. [41] Van Renterghem L, Roelants SLKW, Baccile N, Uyttersprot K, Taelman MC, Everaert B, et al. From lab to market: An integrated bioprocess design approach for new-to-nature biosurfactants produced by Starmerella bombicola. Biotechnol Bioeng 2018;115:1195–206. [42] Loeto D, Jongman M, Lekote L, Muzila M, Mokomane M, Motlhanka K, Ndlovu T, Zhou N. Biosurfactant production by halophilic yeasts isolated from extreme environments in Botswana. FEMS Microbiol Lett 2021;368(20), fnab146. [43] Wittgens A, Tiso T, Arndt TT, Wenk P, Hemmerich J, et al. Growth independent rhamnolipid product from glucose using the non-pathogenic Pseudomonas putida KT2440. Microb Cell Fact 2011;10:80. [44] Wittgens A, Rosenau F. Heterologous rhamnolipid biosynthesis: advantages, challenges, and the opportunity to produce tailor-made rhamnolipids. Front Bioeng Biotechnol 2020;8:1263. [45] Henkel M, Geissler M, Weggenmann F, Hausmann R. Production of microbial surfactants: status quo of rhamnolipid and surfactin towards large-scale production. Biotechnol J 2017;12, 1600561. [46] Bator I, Karmainsky T, Tiso T, Blank LM. Killing two birds with one stone—strain engineering facilitates the development of a unique rhamnolipid production process. Front Bioeng Biotechnol 2020;8:899. [47] Bator I, Karmainsky T, Tiso T, Blank LM. Corrigendum to: killing two birds with one stone—strain engineering facilitates the development of a unique rhamnolipid production process. Front Bioeng Biotechnol 2020;8:1101. [48] Tiso T, Ihling N, Kubicki S, Biselli A, Shonhoff A, et al. Integration of genetic and process engineering for optimized rhamnolipid production using Pseudomonas putida. Front Bioeng Biotechnol 2020;8:976.

42

Chapter 2 New insights in biosurfactants research

[49] Blesken CC, Bator I, Eberlein C, Heipieper HJ, Tiso T, Blank LM. Genetic cell-surface modification for optimized foam fractionation. Front Bioeng Biotechnol 2020;8, 572892. [50] Grosso-Becerra MV, Croda-Garcı´a G, Merino E, Servı´n-Gonza´lez L, Mojica-Espinosa R, Sobero´n-Cha´vez G. Regulation of Pseudomonas aeruginosa virulence factors by two novel RNA-thermometers. Proc Natl Acad Sci USA 2014;111(43):15562–7. [51] Noll P, Treinen C, M€uller S, Senkalla S, Lilge L, Hausmann R, Henkel M. Evaluating temperature-induced regulation of a ROSE-like RNA-thermometer for heterologous rhamnolipid production in Pseudomonas putida KT2440. AMB Express 2019;9(1):154. [52] Noll P, Treinen C, M€uller S, Lilge L, Hausmann R, Henkel M. Exploiting RNA thermometer-driven molecular bioprocess control as a concept for heterologous rhamnolipid production. Sci Rep 2021;11:14802. [53] Moldes AB, Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Lo´pez-Prieto A, Vecino X, Cruz JM. Synthetic and bioderived surfactants versus microbial biosurfactants in the cosmetic industry: an overview. Int J Mol Sci 2021;22:2371. [54] Nikolova C, Gutierrez T. Biosurfactants and their application in the oil and gas industry: current state of knowledge and future perspectives. Front Bioeng Biotechnol 2021;9:46. [55] Mulligan CN. Sustainable remediation of contaminated soils. Front Bioeng Biotechnol 2021;9:195. [56] Durval I, Rufino R, Sarubbo L. Biosurfactant as an environmental remediation agent: toxicity, formulation, and application in the removal of petroderivate in sand and rock walls. Biointerf Res Appl Chem 2022;12 (1):34–48. [57] Haque E, Bin Riyaz MA, Shankar S, Hassan S. Compositional characterization of biosurfactant produced from Pseudomonas aeruginosa ENO14-MH271625 and its application in crude oil bioremediation. Int J Pharm Investig 2021;11(2):204–7. [58] Almeida DG, da Silva RdCF S, Meira HM, PPF B, Silva EJ. Production, characterization and commercial formulation of a biosurfactant from Candida tropicalis UCP0996 and its application in decontamination of petroleum pollutants. Processes 2021;9:885. [59] Dos Santos JCV, da Santos SEM, da Silva YA, Lira IRADS, Raianny Silva R, et al. Application of Candida lipolytica biosurfactant for bioremediation of motor oil from contaminated environment. Chem Eng Trans 2021;86:649–54. [60] Pawlowska A, Sadowski Z, Winiarska K. Effect of rhamnolipids and lipopolysaccharides on the bioleaching of arsenic-bearing waste. Minerals 2021;11:1303. [61] Da Silva M, da GC, Sarubbo LA. Synthetic and biological surfactants used to mitigate biofouling on industrial facilities surfaces. Biointerf Res Appl Chem 2022;12(2):2560–85. [62] Dewidar AA, Sorial GA. Effect of rhamnolipids on the fungal elimination of toluene vapor in a biotrickling filter under stressed operational conditions. Environ Res 2022;204, 111973. [63] Wang T, Chang D, Huang D, Liu Z, Wu Y, et al. Application of surfactants in papermaking industry and future development trend of green surfactants. Appl Microbiol Biotechnol 2021;105:7619–34. [64] Kachrimanidou V, Papadaki A, Lappa I, Papastergiou S, Kleisiari D, Kopsahelis N. Biosurfactant production from Lactobacilli: an insight on the interpretation of prevailing assessment methods. Appl Biocem Biotechnol 2021. https://doi.org/10.1007/s12010-021-03686-7 [published online September 8th]. [65] Vecino X, Rodrı´guez-Lo´pez L, Ferreira D, Cruz JM, Moldes AB. Rodrigues LR Bioactivity of glycolipopeptide cell-bound biosurfactants against skin pathogens. Int J Biol Macromol 2018;109:971–9. [66] Foschi C, Parolin C, Giordani B, Morselli S, Luppi B, et al. Lactobacillus crispatus BC1 biosurfactant counteracts the infectivity of Chlamydia trachomatis elementary bodies. Microorganisms 2021;9:975. [67] Satpute SK, Mone NS, Das P, Banpurkar AG, Banat IM. Lactobacillus acidophilus derived biosurfactant as a biofilm inhibitor: a promising investigation using microfluidic approach. Appl Sci 2018;8:1555. [68] Satpute SK, Mone NS, Das P, Banat IM, Banpurkar AG. Inhibition of pathogenic bacterial biofilms on PDMS based implants by L. acidophilus derived biosurfactant. BMC Microbiol 2019;19:39.

References

43

[69] Mendes RM, Francisco AP, Carvalho FA, Dardouri M, Costa B, et al. Fighting S. aureus catheter-related infections with sophorolipids: electing an antiadhesive strategy or a release one? Colloids Surf B: Biointerfaces 2021;208, 112057. [70] Afsharipour S, Asadi A, Ohadi M, Ranjbar M, Forootanfar H. Preparation and characterization of nano-lipopeptide biosurfactant hydrogel and evaluation of wound-healing properties. BioNanoScience 2021;11:1061–9. [71] Semkova S, Antov G, Iliev I, Tsoneva I, Lefterov P, et al. Rhamnolipid biosurfactants—possible natural anticancer agents and autophagy inhibitors. Separations 2021;8:92. [72] C ¸ elik PA, Manga EB, C¸abuk A, Banat IM. Biosurfactants’ potential role in combating COVID-19 and similar future microbial threats. Appl Sci 2021;11:334. [73] Smith ML, Gandolfi S, Coshall PM, Rahman PKSM. Biosurfactants: a covid-19 perspective. Front Microbiol 2020;11:1341. [74] Bakkar MR, Faraag AHI, Soliman ERS, Fouda MS, Sarguos AMM, et al. Rhamnolipids nano-micelles as a potential hand sanitizer. Antibiotics 2021;10:751. BioNanoScience [published online 26 August] https://doi. org/10.1007/s12668-021-00896-5. [75] Penha RO, Vandenberghe LPS, Faulds C, Soccol VT, Soccol CR. Bacillus lipopeptides as powerful pest control agents for a more sustainable and healthy agriculture: recent studies and innovations. Planta 2020;251:70. [76] Crouzet J, Arguelles-Arias A, Dhondt-Cordelier S, Prsic J, Hoff G, et al. Biosurfactants in plant protection against diseases: rhamnolipids and lipopeptides case study. Front Bioeng Biotechnol 2020;8:1014. [77] De Oliveira CT, Silveira VAI, Andrade G, Macedo Jr F, Celligoi MAPC. Antimicrobial activity of sophorolipids produced by Starmerella bombicola against phytopathogens from cherry tomato. Sci Food Agric 2021. https://doi.org/10.1002/jsfa.11462 [published on line 10 August]. [78] Chen J, Liu X, Fu S, An Z, Feng Y, et al. Effects of sophorolipids on fungal and oomycete pathogens in relation to pH solubility. J Appl Microbiol 2020;128(6):1754–63. [79] Schellenberger R, Crouzet J, Nickzad A, Shu L-A, Kutschera A, et al. Bacterial rhamnolipids and their 3hydroxyalkanoate precursors activate Arabidopsis innate immunity through two independent mechanisms. Proc Natl Acad Sci 2021;118(39), e2101366118. [80] Monnier N, Sarazin C, Rippa S. Transcriptomic dataset from Arabidopsis thaliana seedlings in response to Pseudomonas aeruginosa mono-rhamnolipids. Data Brief 2021;38, 107397. [81] Wang Y, Zhou X, Dai B, Zhu B. Surfactant rhamnolipid promotes anaerobic codigestion of excess sludge and plant waste. Water Sci Technol 2021. https://doi.org/10.2166/wst.2021.414 [published on line October 1]. [82] Li J, Pan H, Yang H, Wang C, Liu H, et al. Rhamnolipids enhances the nitrogen fixation activity of Azotobacter chroococcum by influencing lysine succinylation. Front Microbiol 2021;12, 697963.

CHAPTER

Bioinspired glycolipids: Metals interactions and aqueous-source metal recovery technologies

3

David E. Hogana, Chett J. Boxleyb, Ryan M. Stolleyb, and Raina M. Maiera a

Department of Environmental Science, University of Arizona, Tucson, AZ, United States, bGlycoSurf, Inc., Salt Lake City, UT, United States

1. Introduction One promising area for commercial use of biosurfactants is for recovery of metals from aqueous systems. These can be contaminant metals, such as lead, or metals with strategic value for the manufacture of myriad electronic products, including rare earth elements. There are a variety of aqueous systems that have potential for “metal mining.” These include naturally occurring water sources (e.g., geothermal brines, seawater) or anthropogenic sources (e.g., acid mine drainage, oil and gas produced water, and desalination brines). This chapter will discuss biosurfactant-facilitated mining of metals from aqueous sources beginning with a brief introduction to glycolipid biosurfactants which have been the initial focus for developing metal recovery technologies. This is followed by a description of the synthetic production of glycolipids that are similar to or inspired by their microbial analogs and how synthetic approaches may advance metal recovery. Finally, we will discuss these glycolipids in terms of possible uses and technologies in the area of metal recovery.

2. Glycolipids 2.1 Microbially produced glycolipids The most widely studied microbially produced surfactants are the glycolipids. Microbial glycolipids are structurally diverse but all contain a carbohydrate moiety that is attached to one or more long chain aliphatic acid or hydroxy-aliphatic acid groups. The best studied glycolipid is rhamnolipid which was first isolated from Pseudomonas aeruginosa 141 and characterized by Jarvis and Johnson [1] in 1949 (Fig. 1). Other examples, which demonstrate the great structural diversity of the glycolipids, include the trehalose lipids produced by Rhodococcus species [2]; the sophorolipids produced by yeast species including Candida apicola and Rhodotorula bogoriensis [3]; and xylolipids produced by Lactobacilli [4] (Fig. 2). Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00003-X Copyright # 2023 Elsevier Inc. All rights reserved.

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Chapter 3 Bioinspired glycolipids

FIG. 1 The major microbially produced rhamnolipids. (A) rhamnosyl-β-hydroxydecanoyl-β-hydroxydecanoate; (B) rhamnosyl-rhamnosyl-β-hydroxydecanoyl-β-hydroxydecanoate (C) rhamnosyl-β-hydroxydecanoate; and (D) rhamnosyl-rhamnosyl-β-hydroxydecanoate. n and m range generally from 4 to 8.

2.2 Bioinspired glycolipids One of the roadblocks to commercialization of promising biosurfactant-based technologies is that there has been minimal commercial production of rhamnolipid as well as a significantly higher price point relative to petroleum-based surfactants. To address this concern, there are several companies currently innovating in the glycolipid market with the aim of bringing biosynthetic glycolipids (especially rhamnolipid) to a commercial scale thereby enabling their use for commercial applications. These include companies such as Evonik and Agae Tech. An alternative approach is to create these materials synthetically using green chemistry principles. A library of rhamnolipid molecules was first successfully prepared in 2006 by Bauer et al. [5], using a solid phase system, to study the specificity of immunostimlation against human monocyte cells. This synthetic strategy was subsequently improved upon and patented [6,7], opening the door to large-scale synthetic production of rhamnolipid. GlycoSurf, Inc. licensed this patent in 2016 and uses this synthetic process in combination with green chemistry approaches including the use of biorenewable organic solvents, use of renewable feedstocks as chemical reagents, concentrated reaction schemes (i.e., less solvent use), low temperature operations, and highly efficient glycosylation promotors. There are several advantages of using the synthetic pathway, particularly for applications like metal recovery. These include: (1) Structure-specific production: Biosynthetic pathways for rhamnolipid production are known to produce as many as 60 unique congeners, all generically classified as “rhamnolipid” [8]. This

2 Glycolipids

49

FIG. 2 (A) Trehalose lipid, where x + y ¼ 27 to 31; (B) sophorolipid, where R1 and R2 ¼ H or COCH3; (C) glycoglycerolipid; (D) xylolipid.

complex mixture of nearly inseparable chemical structures is difficult to characterize and explore in a systematic way and there is often batch-to-batch variability in the production of the rhamnolipid [9]. Even modest shifts in the congener mixture can impart changes in surfactant characteristics [e.g., surface tension and critical micelle concentration (CMC)]. On the other hand, in synthetic production of rhamnolipid, the raw material inputs for each step of the synthetic process are known and yields single congeners.

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Chapter 3 Bioinspired glycolipids

Table 1 Cost per kilogram of various carbohydrates. Sugar

CAS number

$/kg

D-Glucose

50-99-7 57-48-7 58-86-6 64,044-51-5 6138-23-4 59-23-4 50-69-1 3458-28-4 10,030-85-0 10,323-20-3 528-50-7 533-67-5 66,009-10-7 2438-80-4

10.50 11.55 17.00 40.00 42.50 82.50 99.00 139.60 181.90 315.30 420.00 750.00 2062.00 5200.00

D-Fructose D-Xylose D-Lactose D-Trehalose D-Galactose D-Ribose D-Mannose L-Rhamnose D-Arabinose D-Cellobiose

2-Deoxy-D-ribose D-Melibiose L-Fucose

(2) Purity: Biosynthetic pathways for rhamnolipid production include numerous secondary fermentation products that are difficult to remove, thus, resulting in purity levels that are around 80%. Further purification of the biosynthetic materials is feasible, but is labor-intensive, solvent-intensive, and generally cost-prohibitive. In contrast, GlycoSurf’s synthetic process has built-in purification steps that produce high purity single-congener molecules (>95% pure). (3) Tailorability: One major advantage of synthetic approaches to making rhamnolipid is that by altering starting materials (type of carbohydrate, type, and number of lipid tails), the structure of the resulting glycolipid can be tailored. This can be expected to alter the performance and cost of the glycolipid produced. The effect of structure on performance is discussed later in this chapter. Table 1 shows a comparison of cost per kg of different sugars. As shown, replacement of rhamnose sugar with glucose decreases the cost of the sugar by 17-fold. (4) Use of biorenewable raw materials: Another advancement in the synthetic process is the design of single-chain glycolipid structures using bio-based starting materials. In order to simplify the production process, and enhance the green chemistry aspects, a reduction of synthetic steps has been achieved by incorporation of beta-hydroxy acids made from biological sources. Not only does this increase the amount of biorenewable materials included in the final product synthesis, but it also reduces the number of synthetic steps, and reduces the overall cost of the final product.

3. Complexation of metals by rhamnolipid In subsequent sections, the glycolipids discussed were biologically derived, unless noted otherwise.

3 Complexation of metals by rhamnolipid

51

3.1 Metal selectivity In 1994, Tan et al. [10] reported the interaction of monorhamnolipid with Cd2+. Since that report, the literature has established important aspects of biosurfactant/metal interactions that make the possibility of metal remediation and recovery technologies so promising: metal interactions are strong, rapid, and stable yet reversible; interaction strengths exceed those of natural metal ligands present in the environment; and interactions are amenable to a wide range of environmental conditions. The discovery of Cd binding triggered additional studies [11,12] that examined the strength of metal interactions for metals across the periodic table. Fig. 3 summarizes these results, showing the conditional stability constants of biosynthetic monorhamnolipid with 26 metals. Hogan et al. [11] divided these metals into three groups based on their conditional stability constants (log β): strongly bound (log β values of >8), moderately bound (4), and weakly bound ( Zn2+ >Cu2+ was determined. The flotation of La3+, Cd2+, and Cs+ using a monorhamnolipid from P. aeruginosa ATCC 9027 was investigated by Hogan et al. [36]. The metals were tested individually and as mixtures. Individually, the

4 Glycolipid-based mining of metals from aqueous sources

57

maximum removal for Cs+, Cd2+, and La3+ was 46.2%, 99.8%, and 98.6%, respectively. When the three metals were tested together at equimolar concentrations, the removals were 39.4%, 98.4%, and 88.1%, respectively. When mixed at a ratio of 200:10:1 (Cs:Cd:La), the removal efficiencies were 49.9%, 99.5%, and 51.5%, respectively. Shetty et al. [37] expanded upon the studies above by testing monorhamnolipid flotation of the rare earth elements La3+, Ce3+, Gd3+, and Yb3+ as a group and as a group mixed with nontarget metals Al3+, Ca2+, Zn2+, and Mg2+. This approach was selected to better represent conditions likely to be encountered in real-world solutions. In their system, monorhamnolipid recovered 90%–95% of the rare earth elements at pH 9. When mixed with the nontarget metals at this pH, the recovery of the rare earth elements remained high, but substantial recovery of Al3+ (80%) and Zn2+ (70%) was also observed; recovery of Ca2+ and Mg2+ was nanofiltration > reverse osmosis), but decreasing the porosity exacerbates problems characteristic of filtration technologies: large energy demand, high pressure requirements, and expensive membranes that are easily fouled [39,40]. Micellar-enhanced ultrafiltration (MEUF) can be used to improve removal efficiencies without decreasing membrane porosity. In this process, a surfactant is added to a solution at concentrations higher than its CMC. When present above the CMC, surfactants aggregate into multimolecular structures such as micelles. These micelles can then bind metals and other contaminants, and the larger size of the aggregates enables their removal by higher-porosity membranes that would otherwise be ineffective for metal removal [40]. Metal-laden aggregates are retained by the filter to form a concentrated retentate, while cleaned water (permeate) passes through the filter. Because some surfactant will be carried with the permeate, the environmental compatibility of biosurfactants makes these molecules ideal for this application [41]. This approach has been demonstrated for both glycolipids as well as the lipopeptide biosurfactant surfactin [42]. El Zeftawy and Mulligan [43] investigated a mixture of mono- and dirhamnolipid for the removal of copper, zinc, nickel, lead, and cadmium using MEUF with 10 and 30 kDa molecular weight cutoff membranes. High rejection ratios (>99%) were found for all the metals with both membranes. Optimal operating conditions of the MEUF system were identified using a response surface methodology: pH 6.9  0.1, 25  1°C, rhamnolipid-to-metal molar ratio of 2:1, and transmembrane pressure of 69 kPa. Using these conditions, six wastewaters from metal refining industries were successfully treated to

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Chapter 3 Bioinspired glycolipids

achieve metal concentrations compliant with federal regulations in Canada. Using the same rhamnolipid, chromium was also successfully removed from solution [44]. 96.2% rejection of Cr(III) was achieved using a rhamnolipid concentration of 0.1% (v/v), a rhamnolipid-to-metal ratio of 36:1. Rhamnolipid itself had up to a 99.4% rejection ratio, indicating few monomers were passed into the permeate. In this study rhamnolipid was also shown to reduce 98.7% of Cr(VI) to Cr(III) using 2% (v/v) rhamnolipid at pH 6. MEUF using rhamnolipid and sophorolipid biosurfactants as collectors for cadmium was optimized by Chai et al. [45] using a response surface methodology and 10 kDa membrane. A model developed to predict system performance was validated, and a maximum Cd2+ rejection of 98.74% was reported under optimal conditions. The model also showed pH had the greatest effect on the filtration efficacy. pH influences both metal chemistry and the nature (e.g., size or shape) of surfactant aggregates [46,47], which supports this finding. Because micelles have both hydrophobic and hydrophilic regions, MEUF can be used to remediate solutions contaminated with both inorganic and organic contaminants [40,41]; charged metals bind the hydrophilic headgroups present on the surface of the aggregate, while organics partition into the hydrophobic core of the aggregate with the lipid tails. Verma and Sarkar [48] removed 98.8% of Cd2+ (60 mg L1) and 25% of p-cresol (75 mg L1) using a mono- and dirhamnolipid mixture and 10 kDa membrane under optimized conditions. These removals for rhamnolipid were equivalent to those found for the commonly used synthetic surfactant sodium dodecyl sulfate. Also using a rhamnolipid mixture, Ridha [49] achieved 100% removal of copper and benzene both separately and when mixed at molar ratios of 6.25 (rhamnolipid:contaminant) for copper and as low as 0.56 for benzene.

4.2.3 Precipitation-based technologies Precipitation of metal contaminants is both effective and widely utilized due to its simplicity and low cost. Common chemical precipitants include hydroxides [e.g., CaO, MgO, NaOH, Ca(OH)2] and sulfides (e.g., FeS or H2S). Hydroxides are low cost and easily handled, but they produce large volumes of low density sludge that can pose disposal challenges. Furthermore, hydroxides are not selective in their activity, and in high enough concentration can lead to the resolubilization of amphoteric metals. Sulfide precipitation yields low-solubility precipitates that are not amphoteric and sludges that are more dense and easily managed. As drawbacks, sulfide precipitates tend to form colloidal dispersions that are difficult to separate, there is little metal selectivity, and they can release toxic H2S under certain pH conditions. Chelating precipitants are highly effective and offer more metal selectivity over chemical precipitants, but many of the commercially available materials (e.g., trimercaptotriazine) pose environmental risks [40]. Glycolipids have been observed precipitating metals in multiple studies. Luna et al. [50] noted the complexation (by conductivity measurements) and precipitation (metal loss from solution measured by atomic absorption spectroscopy) of Cd and Pb when mixed with a biosurfactant produced by Candida sphaerica (likely sophorolipid). The precipitates were removed by centrifugation and recovered as an off-white powder. This precipitation reaction was able to reduce both Cd and Pb concentrations by 95% (initially 1000 ppm) using a 0.05% biosurfactant solution. A similar result for Cd and Pb was observed for another (likely) sophorolipid produced by Candida lipolytica [51]. An unspecified biosurfactant isolated from the marine bacterium Bacillus circulans also precipitated as an off-white material with Pb and Cd with substantial removals exceeding 86% for 1000 ppm metal with 5 x CMC of the biosurfactant [52].

4 Glycolipid-based mining of metals from aqueous sources

59

FIG. 7 Precipitation of lanthanum and biosynthetic monorhamnolipid.

The authors have observed precipitation of both synthetic and biosynthetic monorhamnolipid with various metals. Anecdotally, those metals with higher stability constants (see Fig. 3) tend to form precipitates while those with lower stability constants do not. The nature of these precipitates also varies by metal. For example, lanthanum with biosynthetic monorhamnolipid forms white specks as shown in Fig. 7. Other metals form amorphous, fluffy cloud-like precipitate, while still others form crystal-like flakes. The variability in appearance of metal-rhamnolipid precipitates likely speaks to the chemical structure of the complex, but this has yet to be investigated. The ability of glycolipids to precipitate metals offers a recovery opportunity from aqueous sources using simple processes. Precipitates can be collected by gravity in settling tanks or through more direct means, such as filtration, centrifugation, or flotation (see below) processes. Unlike chemical precipitates, there is potential for glycolipids to selectively precipitate target metals; separations may be possible if precipitation stages are organized in series with glycolipids tuned for specific metals utilized in each step. This application for glycolipids would also benefit from their environmental compatibility, which has been identified as an issue for other chelating precipitants. In a process related to ion flotation, precipitate flotation can be used to remove metals from solution as well. In this process, metal ions are precipitated and the precipitate is removed by floating on the surface of bubbles sparged into the solution, with or without the assistance of a surfactant collector. This approach has been successfully demonstrated with chromium contaminated solutions using

60

Chapter 3 Bioinspired glycolipids

rhamnolipid. Cr(VI) is soluble in water, but when reacted with a reductant such as ferrous sulfate, the chromium is reduced to an insoluble Cr(III) species. Subsequent flotation of these precipitates using rhamnolipid demonstrated very high removal efficiencies [53,54].

4.3 Environmental benefits of glycolipid-based aqueous mining To meet the demands of increasing populations and sustain the proliferation of modern technologies, the extraction of metal resources increased 3.5 times from 1970 to 2017 with an average annual increase of 2.7%. Globally, mining accounts for 10% of climate change impacts, 3% of water stress, 1% of land-use biodiversity loss, and 12% of particulate matter health impacts [55]. These impacts arise intrinsically from the current paradigm of modern mining: enormous masses of stable, solid matrixes must be pulverized to free relatively miniscule amounts of metals into manageable forms. Aqueous mining, on the other hand, exploits resources that are already present in a free form that is easily handled. In lieu of a mine site that requires the disruption of tens of square kilometers of land, aqueous mining can be accomplished in the footprint of a wastewater treatment plant. Instead of contaminating water resources, the aqueous mining approach improves water resources during metals extraction. Energy expenditures and climate effects therefrom are drastically slashed because there is no requirement to move earth or crush and grind ores. Similarly, particulate matter health effects are also reduced as aqueous mining doesn’t disturb landscapes, crush or grind minerals, or create piles of waste rock or tailings from which particulate matter arises [17]. Because lithium is uniquely extracted from both ores and saline solutions (saline sources account for 66% of global supply) [56], it can be used to highlight the ways in which resource extraction can be less environmentally damaging when metals are sourced from aqueous resources. Life-cycle assessment (LCA) for the production of 1 kg of lithium carbonate showed that, in all 10 impact categories, rock-based lithium exceeds the impacts multifold over brine-based—these impact categories include measures of acidification potential, resource depletion, toxicity, and eutrophication potential. For example, rock-based lithium’s global warming potential (kg CO2 eq/kg) was 48 times higher than that of brine-based, and rock-based extraction utilized 30 times more water (kg/kg) than brine-based [57]. Due to increasing demands, decreasing ore grades, increasing orebody complexity, and failure of “economy-of-scale” efficiency advantages, the global environmental impacts of traditional mining are likely to worsen in the future [17]. Aqueous mining technologies are unlikely to ever supplant traditional mining supply; however, the side-by-side LCA impact comparison for sourcing aqueous resources versus traditional hardrock resources makes a compelling argument as to why aqueous resources are going to be increasingly integral as future supplements to material supplies. Furthermore, the remediation of waters through this approach will also address water scarcity issues by generating value-added water products from otherwise degraded resources. Glycolipid-based technologies offer an opportunity to accomplish this task in a manner that is not only environmentally compatible, but also specific and adaptable to various materials and process needs.

4.4 Rhamnolipid treatment of metal-contaminated groundwater: A case study of uranium in Arizona There is a substantial legacy of pollution from the development of the nuclear industry. During the uranium boom, which took place over 4 decades (1944–1986), approximately 4 million tons of uranium ore were extracted from mines in the Southwestern USA. This mining activity left a

4 Glycolipid-based mining of metals from aqueous sources

61

tremendous amount of uranium waste with, in some cases, dangerously high concentrations in the range of 10 mg kg1 to 10 g kg1 [58]. Many of these mine sites are located in arid states such as Arizona, New Mexico, and Utah [59]. In these areas, valuable water resources are impacted by uranium contamination either from legacy mining and milling activities or through continued contamination from unmitigated point sources such as tailings, unprocessed low-grade ores, and contaminated soils. The Navajo Nation alone encompasses over 500 abandoned uranium mining sites which present major human and environmental health risks due to radiological and physical hazards, ecological degradation, and water quality degradation. 518 sites occur within a mile of a perennial or intermittent surface water source; 58 sites, within a quarter mile of drinking water wells [60]. Of tested water sources on the Nation, 12.8% exceed national drinking water standards for uranium [61]. One such site on the Navajo Nation is the inactive Monument Valley uranium processing site in northeast Arizona. Now under the control of the U.S. Department of Energy due to the 1978 Uranium Mill Tailings Radiation Control Act, the Monument Valley site processed materials from 1955 until 1968, and uranium bearing materials were removed from the surface in 1994. Despite the removal of these source materials, some soil and groundwater remain contaminated with uranium. Using groundwater from this site, Hogan et al. [38] examined the utility of rhamnolipid-based ion flotation for the remediation of uranium in a lab-scale system. The aims of this study were to illustrate the effects of pH, monorhamnolipid source (biosynthetic or synthetic), and monorhamnolipid chain length on the efficacy of uranium flotation. A flotation apparatus as shown in Fig. 6 was utilized in this study. The column was borosilicate glass 50 cm tall with a diameter of 5.5 cm. Bubbles were generated using a 10–15 μm pore glass frit and nitrogen gas with a flow rate of 50 mL min1. 250 mL samples of groundwater were adjusted to various pH and combined with 0.5% (v/v) ethanol as frother and 250 μM monorhamnolipid as collector. Both biosynthetic and synthetic (GlycoSurf, Inc.) monorhamnolipid were studied. Sample anion and cation constituents were analyzed using a high-performance ion chromatograph and inductively coupled plasma mass spectrometer, respectively. Geochemical analysis (Table 3) of the groundwater showed the largest cation constituents were Ca, K, Mg, Na, and Si, each in the mg L1 range, and uranium was present at 442 μg L1. Sulfate, nitrate, chlorine, and fluorine were identified as major anions. Initial flotation tests at the native groundwater pH of 8 using (anionic) biosynthetic monorhamnolipid (bio-mRL) showed no uranium removal despite speciation modeling showing a majority (58.3%) of uranium as a cationic species. Decreasing pH was found to increase the efficacy of flotation by biomRL until the concentration was decreased to at or near the U.S. Environmental Protection Agency’s maximum contaminant level of 30 μg L1 at pH 5.5 (Fig. 8). At this pH, bio-mRL flotation was able to reduce the uranium concentration by 92.6% while only removing 6% of the column’s original volume as a uranium concentrate. Bio-mRL for this study was harvested from a P. aeruginosa bacterium that produces primarily rhamnolipid with C10 tails (75%–80% of the congener mixture). For the reasons noted above, biosynthetic materials can be difficult to use in large-scale applications. For this reason, synthetic monorhamnolipids with C10 (Rha-C10C10), C12 (Rha-C12C12), and C14 (Rha-C14C14) chain length tails were also examined in this study. Like the bio-mRL, all three of these monorhamnolipids produced foams at the groundwater’s pH of 8, but none of the molecules collected uranium during flotation. Decreasing the pH to 6.5 revealed the synthetic Rha-C10C10 matched the uranium removal of

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Chapter 3 Bioinspired glycolipids

Table 3 Major metal (>10 μg L21) and anion (>5 μmol L21) constituents present in Monument Valley groundwater. Constituent

μg L21

Al B Ba Ca K Li Mg Na Si Sr U V

44.8 56.0 38.2 52,200 1800 34.8 36,000 26,500 7070 516 442 28.5

Constituent

μmol L21

Cl F NO3 SO4

270 45.1 326 1580

Note: Concentrations are rounded to three significant figures. Table reproduced from Hogan DE, Stolley RM, Boxley C, Amistadi MK, Maier RM. Removal of uranium from contaminated groundwater using monorhamnolipids and ion flotation. J Environ Manage 2022;301:113835. https://doi.org/10.1016/J.JENVMAN. 2021.113835.

bio-mRL. However, neither the synthetic Rha-C12C12 nor Rha-C14C14 were able to generate foams capable of supporting the flotation process, and little to no uranium was removed from solution by these molecules. These results illustrate that: (1) the synthetic Rha-C10C10 performed similarly to the biosynthetic material; (2) monorhamnolipid lipid tail length was critical for performance—the long chain monorhamnolipid failed to support stable foams for effective flotation; (3) solution physiocochemical properties are important—speciation analysis of the groundwater system showed that while cationic uranium species were present at the native pH 8, these species were often relatively large and complex (e.g., (UO2)3(OH)+1 5 ). Decreasing pH led to the predominance of smaller cation species (e.g., UO2F+1, UO2OH+1, UO+2 2 ). The

5 Conclusion

63

450

Uranium Concentration in Column Solution (µg L-1)

400 350 300 250 200 150 100 50 0

5.5 (n=4)

5.75 (n=1)

6 (n=2)

6.25 (n=1)

6.5 (n=2)

pH FIG. 8 Effect of pH on uranium recovery from groundwater during ion flotation. Black bars show initial concentration, and white show final concentration. Number of replicates (n) given in parentheses. Error bars represent ranges for duplicated experiments (pH 6 and 6.5), and the standard deviation the experiment performed in quadruplicate (pH 5.5). Figure reproduced from Hogan DE, Stolley RM, Boxley C, Amistadi MK, Maier RM. Removal of uranium from contaminated groundwater using monorhamnolipids and ion flotation. J Environ Manage 2022;301:113835. https://doi.org/10.1016/J.JENVMAN.2021. 113835.

increase in flotation efficacy with decreasing pH suggests monorhamnolipid’s ability to bind uranium is dependent on the uranium species fitting within the defined binding pocket (Fig. 4).

5. Conclusion In summary, glycolipids preferentially bind toxic and critical metals like the rare earth elements. This interaction is the basis for several promising technologies such as ion flotation, MEUF, and precipitation-based technologies that have potential to meet demand for critical metals from both natural and contaminated water sources. Drawing from biosurfactant structures as inspiration, advances in green synthesis approaches for bioinspired glycolipids has potential to provide structural tunability that may provide a level of functional control over metal binding that is currently not available with natural glycolipids. Taken together, if implemented at commercial scale, these advances may allow recovery of critical metals from aqueous sources and reduce reliance on hardrock mining and the accompanying environmental impacts of this industry.

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References [1] Jarvis FG, Johnson MJ. A glyco-lipide produced by Pseudomonas aeruginosa. J Am Chem Soc 1949;71:4124–6. https://doi.org/10.1021/ja01180a073. [2] Franzetti A, Gandolfi I, Bestetti G, Smyth TJP, Banat IM. Production and applications of trehalose lipid biosurfactants. Eur J Lipid Sci Technol 2010;112:617–27. https://doi.org/10.1002/ejlt.200900162. [3] Ribeiro IA, Castro MF, Ribeiro MH, Gupta VK. Sophorolipids: production, characterization and biologic activity. In: Schmoll M, Tuohy M, Mazutti MA, editors. Applications of microbial engineering. 1st ed. Boca Raton: CRC Press; 2013. p. 367–407. https://doi.org/10.1201/b15250. [4] Saravanakumari P, Mani K. Structural characterization of a novel xylolipid biosurfactant from Lactococcus lactis and analysis of antibacterial activity against multi-drug resistant pathogens. Bioresour Technol 2010;101:8851–4. https://doi.org/10.1016/j.biortech.2010.06.104. [5] Bauer J, Brandenburg K, Z€ahringer U, Rademann J. Chemical synthesis of a glycolipid library by a solidphase strategy allows elucidation of the structural specificity of immunostimulation by rhamnolipids. Chem A Eur J 2006;12:7116–24. https://doi.org/10.1002/chem.200600482. [6] Coss CS, Carrocci T, Maier RM, Pemberton JE, Polt R. Minimally competent Lewis acid catalysts: indium(III) and bismuth(III) salts produce rhamnosides (¼6-deoxymannosides) in high yield and purity. Helv Chim Acta 2012;95:2652–9. https://doi.org/10.1002/hlca.201200528. [7] Pemberton JE, Polt RL, Maier RM, Coss CS. Synthesis of carbohydrate-based surfactants. US9499575B2; WO2014077960A1; 2016. [8] Abdel-Mawgoud AM, Lepine F, Deziel E. Rhamnolipids: diversity of structures, microbial origins and roles. Appl Microbiol Biotechnol 2010;86:1323–36. [9] Zhang L, Pemberton JE, Maier RM. Effect of fatty acid substrate chain length on Pseudomonas aeruginosa ATCC 9027 monorhamnolipid yield and congener distribution. Process Biochem 2014;49:989–95. https:// doi.org/10.1016/j.procbio.2014.03.003. [10] Tan H, Champion JT, Artiola JF, Brusseau ML, Miller RM. Complexation of cadmium by a rhamnolipid biosurfactant. Environ Sci Technol 1994;28:2402–6. [11] Hogan DE, Curry JE, Pemberton JE, Maier RM. Rhamnolipid biosurfactant complexation of rare earth elements. J Hazard Mater 2017;340:171–8. https://doi.org/10.1016/j.jhazmat.2017.06.056. [12] Ochoa-Loza FJ, Artiola JF, Maier RM. Stability constants for the complexation of various metals with a rhamnolipid biosurfactant. J Environ Qual 2001;30:479–85. [13] Schalnat TA. Metal complexation and interfacial behavior of the microbially-produced surfactant monorhamnolipid by Pseudomonas aeruginosa ATCC 9027. University of Arizona; 2012. [14] Petty TR. Final list of critical minerals 2018. Fed Regist 2018;83:23295–6. [15] European Commission. Directorate-General for Internal Market, Industry, Entrepreneurship and SMEs. In: Critical raw materials resilience: charting a path towards greater security and sustainability; 2020. https://eurlex.europa.eu/legal-content/EN/ALL/?uri¼CELEX:52020DC0474. [Document date: 03/09/2020, Report number: COM/2020/474 final]. [16] Blengini GA, Nuss P, Dewulf J, Nita V, Talens Peiro´ L, Vidal-Legaz B, et al. EU methodology for critical raw materials assessment: policy needs and proposed solutions for incremental improvements. Resour Policy 2017;53:12–9. https://doi.org/10.1016/J.RESOURPOL.2017.05.008. [17] Can Sener SE, Thomas VM, Hogan DE, Maier RM, Carbajales-Dale M, Barton MD, et al. Recovery of critical metals from aqueous sources. ACS Sustain Chem Eng 2021;9:11616–34. https://doi.org/10.1021/ acssuschemeng.1c03005. [18] Fortier SM, Nassar NT, Lederer GW, Brainard J, Gambogi J, McCullough EA. Draft critical mineral list—summary of methodology and background information—U.S. Geological Survey technical input document in response to Secretarial Order No. 3359. Reston, VA; 2018. https://doi.org/10.3133/ofr20181021.

References

65

[19] Critical Materials Institute. Critical materials periodic table 2020, https://www.ameslab.gov/cmi/researchhighlights/critical-materials-periodic-table. [Accessed 2 February 2022]. [20] Gammons CH, Wood SA, Jonas JP, Madison JP. Geochemistry of the rare-earth elements and uranium in the acidic Berkeley pit lake, Butte. Montana Chem Geol 2003;198:269–88. https://doi.org/10.1016/S0009-2541 (03)00034-2. [21] Hogan DE, Veres-Schalnat TA, Pemberton JE, Maier RM. Biosurfactant complexation of metals and applications for remediation. In: Mulligan CN, Mudhoo A, Sharma SK, editors. Biosurfactants: research trends and applications. Boca Raton, FL: CRC Press; 2014. p. 277–308. https://doi.org/10.1201/b16383. [22] Mulligan CN. Recent advances in the environmental applications of biosurfactants. Curr Opin Colloid Interface Sci 2009;14:372–8. https://doi.org/10.1016/J.COCIS.2009.06.005. [23] Pacwa-Płociniczak M, Płaza GA, Piotrowska-Seget Z, Cameotra SS. Environmental applications of biosurfactants: recent advances. Int J Mol Sci 2011;12:633–54. https://doi.org/10.3390/ijms12010633. [24] Sarubbo LA, Rocha Jr RB, Luna JM, Rufino RD, Santos VA, Banat IM. Some aspects of heavy metals contamination remediation and role of biosurfactants. Chem Ecol 2015;31:707–23. [25] Sebba F. Concentration by ion flotation. Nature 1959;184:1062–3. [26] Jorne J, Rubin E. Ion fractionation by foam. Sep Sci Technol 1969;4:313–24. [27] Pinfold TA. Ion flotation. In: Lemlich R, editor. Adsorptive bubble separation techniques. New York: Academic Press; 1972. p. 53–73. [28] Zouboulis AI, Matis KA. Ion flotation in environmental technology. Chemosphere 1987;16:623–31. https:// doi.org/10.1016/0045-6535(87)90275-X. [29] Banat IM, Makkar RS, Cameotra SS. Potential commercial applications of microbial surfactants. Appl Microbiol Biotechnol 2000;53:495–508. [30] Menezes CTB, Barros EC, Rufino RD, Luna JM, Sarubbo LA. Replacing synthetic with microbial surfactants as collectors in the treatment of aqueous effluent produced by acid mine drainage, using the dissolved air flotation technique. Appl Biochem Biotechnol 2011;163:540–6. https://doi.org/10.1007/s12010-010-9060-7. [31] Albuquerque CF, Luna-Finkler CL, Rufino RD, Luna JM, De Menezes CTB, Santos VA, et al. Evaluation of biosurfactants for removal of heavy metal ions from aqueous effluent using flotation techniques. Int Rev Chem Eng 2012;4:156–61. [32] Rangarajan V, Sen R. An inexpensive strategy for facilitated recovery of metals and fermentation products by foam fractionation process. Colloids Surf B Biointerfaces 2013;104:99–106. https://doi.org/10.1016/j. colsurfb.2012.12.007. [33] Chen H-R, Chen C-C, Reddy AS, Chen C-Y, Li WR, Tseng M-J, et al. Removal of mercury by foam fractionation using surfactin, a biosurfactant. Int J Mol Sci 2011;12:8245–58. https://doi.org/10.3390/ ijms12118245. [34] Yuan XZ, Meng YT, Zeng GM, Fang YY, Shi JG. Evaluation of tea-derived biosurfactant on removing heavy metal ions from dilute wastewater by ion flotation. Colloids Surf A Physicochem Eng Asp 2008;317: 256–61. https://doi.org/10.1016/j.colsurfa.2007.10.024. [35] Bodagh A, Khoshdast H, Sharafi H, Zahiri HS, Noghabi KA. Removal of cadmium(II) from aqueous solution by ion flotation using rhamnolipid biosurfactant as an ion collector. Ind Eng Chem Res 2013;52:3910–7. [36] Hogan DE, Curry JE, Maier RM. Ion flotation of La3+, Cd2+, and Cs+ using monorhamnolipid collector. Colloids Interfaces 2018;2:43. https://doi.org/10.3390/colloids2040043. [37] Shetty S, Chernyshova IV, Ponnurangam S. Foam flotation of rare earth elements by conventional and green surfactants. Miner Eng 2020;158. https://doi.org/10.1016/j.mineng.2020.106585, 106585. [38] Hogan DE, Stolley RM, Boxley C, Amistadi MK, Maier RM. Removal of uranium from contaminated groundwater using monorhamnolipids and ion flotation. J Environ Manage 2022;301. https://doi.org/ 10.1016/J.JENVMAN.2021.113835, 113835.

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[39] Malaeb L, Ayoub GM. Reverse osmosis technology for water treatment: state of the art review. Desalination 2011;267:1–8. https://doi.org/10.1016/j.desal.2010.09.001. [40] Fu F, Wang Q. Removal of heavy metal ions from wastewaters: a review. J Environ Manage 2011;92:407–18. [41] Mulligan CN. Enhancement of remediation technologies with biosurfactants. In: Mulligan CN, Mudhoo A, Sharma SK, editors. Biosurfactants: research trends and applications. Boca Raton, FL: CRC Press; 2014. p. 231–76. https://doi.org/10.1201/b16383. [42] Mulligan CN, Yong RN, Gibbs BF, James S, Bennett HPJ. Metal removal from contaminated soil and sediments by the biosurfactant surfactin. Environ Sci Technol 1999;33:3812–20. https://doi.org/10.1021/ es9813055. [43] El Zeftawy MAM, Mulligan CN. Use of rhamnolipid to remove heavy metals from wastewater by micellarenhanced ultrafiltration (MEUF). Sep Purif Technol 2011;77:120–7. https://doi.org/10.1016/ j.seppur.2010.11.030. [44] Abbasi-Garravand E, Mulligan CN. Using micellar enhanced ultrafiltration and reduction techniques for removal of Cr(VI) and Cr(III) from water. Sep Purif Technol 2014;132:505–12. https://doi.org/10.1016/ J.SEPPUR.2014.06.010. [45] Chai T, Yan H, Zhang Z, Xu M, Wu Y, Jin L, et al. Optimization of enhanced ultrafiltration conditions for cd with mixed biosurfactants using the box-Behnken response surface methodology. Water 2019;11:442. https://doi.org/10.3390/w11030442. [46] Dahrazma B, Mulligan CN, Nieh MP. Effects of additives on the structure of rhamnolipid (biosurfactant): a small-angle neutron scattering (SANS) study. J Colloid Interface Sci 2008;319:590–3. https://doi.org/ 10.1016/J.JCIS.2007.11.045. [47] Champion JT, Gilkey JC, Lamparski H, Retterer J, Miller RM. Electron-microscopy of rhamnolipid (biosurfactant) morphology—effects of pH, cadmium, and octadecane. J Colloid Interface Sci 1995;170: 569–74. https://doi.org/10.1006/jcis.1995.1136. [48] Verma SP, Sarkar B. Rhamnolipid based micellar-enhanced ultrafiltration for simultaneous removal of Cd(II) and phenolic compound from wastewater. Chem Eng J 2017;319:131–42. https://doi.org/10.1016/ J.CEJ.2017.03.009. [49] Ridha ZAM. Simultaneous removal of benzene and copper from water and wastewater using micellarenhanced ultrafiltration. Concordia Universiy; 2010. [50] Luna JM, Rufino RD, Sarubbo LA. Biosurfactant from Candida sphaerica UCP0995 exhibiting heavy metal remediation properties. Process Saf Environ Prot 2016;102:558–66. https://doi.org/10.1016/ J.PSEP.2016.05.010. [51] Santos DKF, Resende AHM, de Almeida DG, de Soares da Silva CF R, Rufino RD, Luna JM, et al. Candida lipolytica UCP0988 biosurfactant: potential as a bioremediation agent and in formulating a commercial related product. Front Microbiol 2017;8. https://doi.org/10.3389/fmicb.2017.00767. [52] Das P., Mukherjee S., Sen R.. Biosurfactant of marine origin exhibiting heavy metal remediation properties. Bioresour Technol 2009;100:4887–90. https://doi.org/The interaction. [53] Salmani Abyaneh A, Fazaelipoor MH. Evaluation of rhamnolipid (RL) as a biosurfactant for the removal of chromium from aqueous solutions by precipitate flotation. J Environ Manage 2016;165:184–7. https://doi. org/10.1016/j.jenvman.2015.09.034. [54] Shojaei V, Khoshdast H. Efficient chromium removal from aqueous solutions by precipitate flotation using rhamnolipid biosurfactants. Physicochem Probl Miner Process 2018;54:1014–25. [55] Oberle B, Bringezu S, Hatfeld-Dodds S, Hellweg S, Schandl H, Clement J, et al, Global Resources Outlook. Natural resources for the future we want 2019., 2019, https://www.resourcepanel.org/reports/globalresources-outlook. [56] Goonan TG. Lithium use in batteries: U.S. geological survey circular 1371 (2012)., 2012, https://pubs.usgs. gov/circ/1371/.

References

67

[57] Jiang S, Zhang L, Li F, Hua H, Liu X, Yuan Z, et al. Environmental impacts of lithium production showing the importance of primary data of upstream process in life-cycle assessment. J Environ Manage 2020;262. https://doi.org/10.1016/J.JENVMAN.2020.110253, 110253. [58] Abdelouas A. Uranium mill tailings: geochemistry, mineralogy, and environmental impact. Elements 2006;2:335–41. https://doi.org/10.2113/gselements.2.6.335. [59] Office of Legacy Management. Defense-related uranium mines FY 2017 annual report. Washington, DC: U.S. Department of Energy; 2018. [60] Ram NM, Moore C, McTiernan L. Cleanup options for Navajo abandoned uranium mines. Remediat J 2016;26:131–48. https://doi.org/10.1002/rem.21473. [61] Hoover J, Gonzales M, Shuey C, Barney Y, Lewis J. Elevated arsenic and uranium concentrations in unregulated water sources on the Navajo Nation, USA. Expo Health 2017;9:113–24. https://doi.org/10.1007/ s12403-016-0226-6.

CHAPTER

Rhamnolipids—Has the promise come true?

4

Holger Dittmanna, Eric D
ezielb, Marius Henkela, and Rudolf Hausmanna a

Department of Bioprocess Engineering, University of Hohenheim, Institute of Food Science and Biotechnology, e Biotechnologie, Institut National de la Recherche Scientifique Stuttgart, Germany, bCentre Armand-Frappier Sant
(INRS), Universit
e du Qu
ebec, Laval, QC, Canada

1. Introduction Rhamnolipids are very well studied bacterial glycolipid biosurfactants mostly known to be produced by Pseudomonas and Burkholderia species. They present excellent surface-activity, emulsifying and foaming as well as several other advantageous properties. An almost overwhelming number of scientific publications present their wide range of possible applications due to their surface-active and environmental properties, such as high biodegradability and low toxicity. Reported applications are mainly in the fields of hydrocarbon remediation, tertiary oil recovery, and personal care and food and household products. In addition, the antifungal and antimicrobial properties, low toxicity and immunostimulatory activities of rhamnolipids for applications in the agricultural, cosmetic, and pharmaceutical fields are largely described. As one of the few well-characterized microbial biosurfactants, they are easily produced by wild-type strains and are now commercially available from several manufacturers. Therefore, they represent a broad application interest in many industries. The market research and management consulting company Global Market Insights Inc. predicts that the rhamnolipid-based biosurfactants market will reach over USD 145 million by 2026 [1]. Several excellent and comprehensive reviews have been published over the years, including recent ones from Sobero´n-Cha´vez et al. (2021) and Kumar and Das (2018) [2,3].

2. Rhamnolipids prospects in retrospect view 2.1 Exploitation of antibiotic activity Research on rhamnolipid glycolipids and their applications dates back over seven decades. From the very beginning, antimicrobial properties were the focus of interest. Already in 1946 Bergstr€om et al. reported on a metabolic product (called pyolipic acid) of Pseudomonas pyocyanea, nowadays Pseudomonas aeruginosa, comprising a β-hydroxy-n-decanoic acid lipid moiety (Fig. 1) and that this compound inhibited Mycobacterium tuberculosis [4]. Several crystalline compounds and two partially purified oils were obtained from cultures of P. aeruginosa by Jarvis and Johnson (1949) [5]. These Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00004-1 Copyright # 2023 Elsevier Inc. All rights reserved.

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Chapter 4 Rhamnolipids—Has the promise come true?

FIG. 1 Most common congener structure of di-rhamnolipid (Rha-Rha-C10-C10) from P. aeruginosa.

substances were studied with respect to their antibacterial activity. Structural investigation of an acidic crystalline glycolipid, revealed the presence of L-rhamnose and β-hydroxy-n-decanoic acid in the structure, just as the pyolipic acid described previously. In contrast to pyolipic acid, the molecular weight was higher, as was the rhamnose-hydroxy acid ratio. A bacteriostatic effect was likewise observed against M. tuberculosis. The glycolipid from P. aeruginosa was further characterized by Edwards and Hayashi (1965) who assigned an alpha configuration to the rhamnosyl residues and reported the structure as 2-O-α-L-rhamnopyranosyl-α-L-rhamnopyranosyl-β-hydroxydecanoyl-βhydroxydecanoic acid [6–8]. Rhamnolipids are generally considered amphotheric molecules that promote the uptake and biodegradation of poorly water soluble substrates, such as vegetable oils and alkanes. Recent studies have attributed this role to a change in surface properties during interaction with microorganisms as well as on adherence [9,10]. Besides this, their function as immunomodulators, virulence factors, and antimicrobials, as well as their role in surface motility and biofilm development have been reported [11,12]. Nevertheless, their exact physiological function has not been comprehensively elucidated by now. Most of the identified bioactivities are derived from the surface activity, wettability, and other amphipathic properties of these molecules. As early as 1986 a first world patent for the continuous production of rhamnolipids which could be used as surfactants was granted [13]. In this patent, continuous cultures of Pseudomonas in submerged culture under aerobic conditions, are described for the production of rhamnolipids.

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To date, a moderate variety of rhamnolipid congeners and homologues have been described, which are confidently formed by only few bacterial species, essentially belonging to the Pseudomonas and Burkholderia genera. In addition, numerous reports of rhamnolipids from bacteria of other genera, families, classes, or even phyla exist, but require critical evaluation of the description in individual cases [14]. Indeed, the literature is contaminated with many studies lacking robust experimental evidence that rhamnolipids are actually produced by new bacterial species [15], thus one must remain critical. The excellent amphoteric properties of rhamnolipids and their easy bioproduction with available wild-type microorganisms have led to countless studies on the application of rhamnolipids. When comparing research in the field of biosurfactants in general to rhamnolipid-related research, as identified by keyword analysis in published research articles (Fig. 2), it is evident that there is continuous research interest in rhamnolipids. This becomes especially clear when investigating the fraction of rhamnolipid-related research compared to the overall included data. The potential uses of rhamnolipids have been widely investigated, for example as surfactants in detergents, personal care products, soil remediation, agriculture as biopesticides, especially against fungal infestation. The antimicrobial activities with an emphasis on agricultural and biomedical applications have been recently summarized [16–18]. In agricultural applications, at least one rhamnolipid-based product, Zonix Biofungicide commercialized by Jeneil Biosurfactant Company, LLC is successfully established. Jeneil claims that Zonix provides protection from plant pathogens and zoosporic contamination by oomycetes such Phytophthora, and Pythium, and that Zonix offers a safe, environmentally sound, and sustainable alternative to prevent disease outbreaks in crops [19].

FIG. 2 Articles per year from 2000 to 2021 taken from the peer-reviewed literature database Scopus (www.scopus.com) with respect to the keywords “biosurfactant,” “rhamnolipid,” and the quotient of “rhamnolipid”/“biosurfactant.” Results are limited to “article” and keyword mentioned in “article title, abstract, keywords.”

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Chapter 4 Rhamnolipids—Has the promise come true?

Rhamnolipids could also be used as an alternative to conventional synthetic antimicrobial preservatives for postharvest fruits and vegetables, or in cleaning solutions and disinfectants. In this context, biofilms are of particular interest as they can be a source of persistent contamination. For example, during food processing, they can contribute to food spoilage and disease transmission. To prevent the adhesion of bacteria and the formation of biofilms, there is a particular interest in treating surfaces with biosurfactants to modify the surface properties to make it more difficult for bacteria to adhere. Araujo et al. (2016), Hajfarajollah et al. (2015), Magalha˜es and Nitschke (2013), Meylheuc et al. (2006), Nitschke et al. (2009) as well as Arau´jo et al. (2011) analyzed the adhesion/antibiofilm and antimicrobial effects of rhamnolipid and other biosurfactants, against different microorganisms especially pathogens [20–25]. They all report on potential of tested rhamnolipids to decrease adhesion and biofilm formation on various surfaces. In addition to their broad applicability as antimicrobial agents, rhamnolipids can also find excellent applications due to their surface-active properties. The following paragraphs outline important areas of application based on examples of recent and older works.

2.2 Applications in environmental remediation, petroleum, and related areas Rhamnolipids use has been extensively investigated in the areas of biological environmental remediation where they can facilitate the removal of environmental pollutants such as hydrocarbons and heavy metals. Oil spills, which are unavoidable during extraction, processing, transportation and storage, pose a serious environmental threat to soil and groundwater. Biosurfactants such as rhamnolipids reduce the interfacial tension between oil and water and can thus facilitate the migration and dissolution of oil contaminants. There have been a number of detailed reviews on this topic over recent years [26–32]. Liu et al. (2021) summarize the characteristics of different types of surfactants such as nonionic, anionic, biological and mixed surfactants, their enhancements to the remediation of oilcontaminated soil and groundwater, and examine the factors influencing surfactant performance. In their review, Liu et al. (2021) compared various surfactant reports including the plant biosurfactant saponin and microbial biosurfactants such as surfactin and rhamnolipid for their remediation efficiency of oil-contaminated soils. Rhamnolipid showed the highest total petroleum hydrocarbons (TPH) removal efficiency in one study, which was higher than other biosurfactants and also significantly higher than select chemical surfactants (Tween 80 and Triton X-100); under other conditions, rhamnolipid displayed a better performance than four other biosurfactants (saponin, aescin, tannin, and lecithin) but slightly weaker than anionic synthetic surfactant sodium dodecyl sulfate (SDS). The summarized laboratory studies showed that rhamnolipids have the potential to improve the remediation of oil-contaminated groundwater. Liu et al. (2021) conclude that this demonstrates the excellent interfacial activity of rhamnolipid and its great potential for oil removal in aquifers [26]. Fenibo et al. (2019) reported that biosurfactants in general can compete with synthetic surfactants in petroleum industry applications and related environmental remediation, as they have proven advantages over synthetic surfactants, including environmental friendliness, biodegradability, low toxicity, and stability over a wide range of environmental conditions. Specifically, rhamnolipids have been used for various types of remediation of hydrocarbon spills more than any other biosurfactants. Fenibo et al. (2019) state that rhamnolipids have been proven to remove spills and also metals, however, at present

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synthetic surfactants remain preferred because, unlike biosurfactants, they are available in commercial quantities and prices [27]. Liu et al. (2018) provide a comprehensive review of the application of rhamnolipids in soil and groundwater remediation for the removal of hydrocarbon and heavy metal contaminants. The important properties of rhamnolipids related to pollutant removal are solubilization, emulsification, dispersion, foaming, wetting, complexation, and the ability to change the surface properties of bacterial cells. Especially, the remediation of hydrocarbons and their derivatives as well as metal contamination. Liu et al. (2018) conclude that the application of rhamnolipid is a good option to enhance the removal of these contaminants. Specifically, they addressed the use of rhamnolipids for bioremediation, soil washing, and other remediation technologies [29]. Rhamnolipid can either increase or decrease the availability of hydrocarbons (namely, hexadecane) for degradation by pseudo-solubilization dependent on the microorganism [33]. Also a rhamnolipid treatment may facilitate interfacial uptake of hexadecane [34]. Since efficient transport of bacterial cells to or through remediation sites is required for groundwater bioremediation the potential of rhamnolipids to enhance this factor may further improve the degradation process [35]. The ability of rhamnolipids to enhance the bioavailability of hydrocarbons through rhamnolipid solubilization has received considerable attention in soil bioremediation, and has been reported in numerous publications. Notable among these are reports of enhanced biodegradation of phenanthrene and of diesel oil in the presence of rhamnolipids. However, Liu et al. (2018) emphasize that the process of bioremediation with rhamnolipids is a complex phenomenon influenced by many factors, including soil and aqueous phase properties (e.g., rhamnolipid concentrations, pH, ionic strength, dissolved organic matter). Accordingly, bioavailability, hydrocarbon degradation activity, and bacterial transport may vary [29]. In the field of soil washing, the use of rhamnolipids succeeds in increasing the release of hydrocarbons from soils. The enhanced leaching of hydrocarbons from soils by rhamnolipids may be due to two mechanisms: desorption and pseudo-solubilization by formation of micelles. Bioavailability of micellar hydrophobic organic compounds is based on hemi-micelle formation on cell surfaces. Rhamnolipids-induced release of LPS and rhamnolipids adsorption can change cell surface hydrophobicity and therefore enhance bioavailability [36]. Liu et al. (2018) reported many successful examples that rhamnolipids can significantly enhance the removal of hydrocarbons from soil by washing techniques such as in the removal of TPH from low- and high-impact soils, in the removal of aromatic and paraffinic hydrocarbons, and in the removal of petroleum and diesel oil from contaminated soils. [29] According to Liu et al. (2018), these results indicate that rhamnolipids provide comparatively better performance than compared surfactants in the removal of petroleum hydrocarbons [29]. Olasanmi and Thring (2020) evaluated the rhamnolipid-enhanced washing of petroleumcontaminated soils and reached a maximum TPH reduction of 58.5% and even 76.8% for drill cuttings [37]. However, the efficiency of rhamnolipids to remove hydrocarbons when used for soil washing varies depending on the concentration and type of hydrocarbons. Other rhamnolipid-based applications reported include improved phytoremediation of soils contaminated with polycyclic aromatic hydrocarbons and improved oxidation of pollutants [29]. Heavy metals can be removed from soils by biosurfactant-enhanced washing and biosurfactantenhanced bioextraction. The mechanisms for removal of heavy metals from soils involves dissolution, surfactant-associated complexation, and ion-exchange [38]. Specifically, anionic biosurfactants such

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Chapter 4 Rhamnolipids—Has the promise come true?

as rhamnolipids with negative charge can form ionic bonds with cationic metals, resulting in stronger stabilization forces in the rhamnolipid-metal complex compared to those between the metal and the soil. Rhamnolipids can also compete for adsorption sites on the soil particles. As such, electrostatic repulsion between soil particles is enhanced and metal ions are mobilized. In addition, metal cations can also be enclosed by the rhamnolipid micelles. The use of rhamnolipids to enhance the recovery of heavy metal contaminants such as chromium, nickel, zinc, copper, lead, and cadmium from soils has attracted considerable attention over the years [29]. The summarized results lead Liu et al. (2018) to conclude that rhamnolipid was most effective in removing these metals from soil compared to other surfactants. Rhamnolipids have also been favorably evaluated for use in bioremediation of soils contaminated simultaneously by organic compounds and metals [29]. The physicochemical properties of rhamnolipid, i.e., solubilization, emulsification, and mobilization for hydrophobic organic compounds and complexation for metals, provide excellent performance in soil washing for abiotic removal of hydrocarbon and metal contaminants. Liu et al. (2018) state that most studies on rhamnolipid-assisted soil remediation show good results in the laboratory, but they encounter several drawbacks in actual remediation of contaminated sites in practice. Finally, significantly more field trials on the use of rhamnolipids for remediation should be conducted in the near future. Since field conditions are much more complex than the well-controlled conditions of laboratory experiments, the information obtained in the laboratory is not sufficient. For successful application of rhamnolipid in remediation, further accumulation of knowledge from field experiments is necessary [29]. The use of surfactants for enhanced oil recovery as a means of reducing interfacial tension has already been extensively investigated with the use of microbial surfactants. In addition to rhamnolipids, surfactin in particular has been tested. For example, recent works include that of Sakthipriya et al. (2021), Rocha et al. (2020), C^amara, J
essica Maria Damia˜o de Arruda et al. (2019), Sharma et al. (2018), and Cui et al. (2017) [39–43]. Sakthipriya et al. (2021), compared the effect of the biosurfactants, surfactin and rhamnolipids, on enhanced crude oil recovery with commercial conventional surfactants. They report that biosurfactants achieved higher oil recovery efficiency. With cetyltrimethylammonium bromide (CTAB) and SDS, at the same concentration, the recovery of crude oil improved by 7 and about 9%, respectively, while the efficiency of biosurfactants was about 15.5%. They attributed this to the reduction in interfacial tension between the crude oil and water systems in porous media, which was found to be the main cause of the improved oil recovery. They concluded that the enhanced oil recovery even at low concentrations and the stability at high salinity, pH, pressure, and temperature make rhamnolipids excellent candidates for their use in microbial enhanced oil recovery (MEOR) [39]. Rocha et al. (2020) compared the use of mono- and di-rhamnolipid mixtures for microbial MEOR applications. For this purpose, natural rhamnolipid mixtures with different proportions of mono- and di-rhamnolipids were used. All rhamnolipids showed robustness to high temperatures, salinity and pH variations, and high oil displacement ability, foam stability and wettability. Monorhamnolipids exhibited the lowest surface tension (26.40 mN/m), lowest interfacial tension (1.14 mN/m) and critical micelle concentration (CMC 27.04 mg/L), the highest emulsification index (EI24 100%) and the best wettability. Based on the results, it was concluded that mono-rhamnolipid has the best capacity for incorporating oil into micelles due to its most effective surface active physicochemical properties. Rocha et al. (2020) concluded that all rhamnolipids studied have potential for MEOR applications [40].

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C^amara, J
essica Maria Damia˜o de Arruda et al. (2019) evaluated the biosurfactant produced by P. aeruginosa in view of its ability to be used in MEOR and report that rhamnolipid at 2X the CMC showed a contribution to the total petroleum recovery factor by MEOR of around 12% [41].

2.3 Cosmetics, personal care, and household products For use in cosmetics, such as soaps, toothpastes, shampoos, detergents and creams and other consumer end-products, there is increasing demand for environment-friendly and sustainably produced ingredients. Rhamnolipids, which exhibit good to very good emulsification, wetting, foaming, and may at the same time be sustainably produced and are biodegradable, could be particularly suitable for this purpose. Thanks to these excellent properties, rhamnolipids are suitable for many different applications in these areas. Adu et al. (2020) and Lourith and Kanlayavattanakul (2009) reviewed the application of rhamnolipids and glycolipids in general in cosmetics and personal care products [44,45]. Rhamnolipids have been used in various healthcare products such as insect repellents, antacids, acne pads, contact lens solutions, and deodorants [46,47]. These products require surfactants which have high surface and emulsifying activities [48], the latter particularly key to the texture consistency of these products [49]. In addition to these special functions, the biological activities required for cosmetics expand the application of these surfactants and a delivery system has been achieved not only for emulsions but also for liposomes [50]. Cosmetics containing rhamnolipids have been patented having antiwrinkle and antiageing properties [51]. Because of their skin compatibility and extremely low skin irritation, several products have been launched in the past [49,52]. However, the pathogenicity status of P. aeruginosa producing rhamnolipids possibly represented an obstacle for their acceptance for use in food and cosmetic products and their production in heterologous hosts, such as Pseudomonas putida represents a big advance in the field. In this respect, only very recently rhamnolipid-based household products containing rhamnolipid produced in heterologous hosts, were launched on the market for the first time. Of particular note here are Unilever’s “Sunlight” or “Quix” dishwashing liquid, which were recently launched in Chile and Vietnam, respectively [53,54].

2.4 Food application There have been only few reports of potential applications of rhamnolipids in food, although the food industry may have a need for new emulsifiers and surfactants. One possible reason is that rhamnolipids are naturally produced by the opportunistically pathogenic bacterium P. aeruginosa, as already commented, and associated with its virulence. Thus, the main reason for the limited use of rhamnolipids in the food sector seems to be the lack of information on toxicity combined with the difficult commercial access in suitable grades. Reviews on biosurfactants applications in food are available [55–57]. The use of rhamnolipids in food formulations has been disclosed in two patents [58,59] to improve the shelf life and stability of bakery and dairy products. Improvement of dough stability, texture, volume and shelf life of bakery products has been presented by the addition of rhamnolipid surfactants, as well as the use of rhamnolipids to improve the properties of buttercream, croissants and frozen confectionery. It is in the nature of patents that the inventions disclosed herein cannot be reproduced with scientific accuracy. Furthermore, no food science publication by these authors on the subject has come to the public’s attention.

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In the review presented by Thakur et al. (2021), the authors mainly focus on the use of rhamnolipids against foodborne pathogens, such as Escherichia, Bacillus spp., Listeria, Campylobacter spp., Staphylococcus spp., Salmonella spp., and Clostridium spp., i.e., as preservative additives [60]. Rhamnolipids could be used as an alternative to prevent food contamination due to their antimicrobial properties against a wide range of microbes. In particular, they are capable of controlling the growth of Grampositive bacteria in acidic foods by enhancing their antimicrobial activity in acidic environments. Thakur et al. (2021) conclude that rhamnolipids could be used in the food industry to prevent food spoilage without compromising food quality [60].

3. Rhamnolipid bioproduction Bioproduction of rhamnolipids with P. aeruginosa is relatively simple and natural isolates can readily be found that produce rhamnolipids in useful titers. By mutagenesis and selective screening, Giani et al. (1997) generated high producer strains already in the 1990s that were used industrially for the production of rhamnolipids [61]. In a related patent, Giani et al. (1997) claimed they could achieve titers of 74–112 g/L of rhamnolipids with strains DSM 7107 and DSM 7108 [61]. This was achieved by batch and fed-batch cultures but could not be independently reproduced scientifically maybe because important methodological details are missing. The prototypic P. aeruginosa model strain PAO1 produces rhamnolipids in significant amounts and has also been used for studies of natural quorum sensing-regulated rhamnolipid biosynthesis. Rhamnolipid bioproduction has been studied bioprocess-wise in innumerable aspects using wildtype P. aeruginosa strains. However, complex aspects of molecular regulation in conjunction with process engineering frameworks that could lead to a deeper understanding of process progression remain few. Among these are the study of the complex quorum sensing regulatory network, monitoring of gene expression, modeling of rhamnolipid production, optimization of media composition and process control, and downstream processing techniques. For example, Schmidberger et al. (2013) studied the time course of gene expression in a batch process with nitrogen limitation [62]. Henkel et al. (2013) presented a kinetic model for the bioproduction of rhamnolipids in batch processes, including taking into account the cell density-dependent quorum sensing dynamics [63]. This model was used in a subsequent study to further develop a process model for rhamnolipid production in a bioreactor that could satisfactorily describe the time course for biomass, substrates, by-products, rhamnolipid, and extracellular quorum sensing signal molecules [64]. The basic complex problem of maintaining a consistently high product formation rate with wild-type strains has not been solved to date. Usually, vegetable oils are preferred as carbon source and nitrate salts as nitrogen source. The vegetable oil is usually used in large excess and also serves as a C-source and antifoaming agent whereby it is broken down extracellularly into glycerol and fatty acids. The released fatty acids, in turn, could exert an inhibitory effect if their concentration accumulates too high. Carbon control is therefore very useful and at the same time difficult to implement. The nitrogen source is usually added in limiting dosage from the beginning. To achieve a high induction of the biosynthetic rate, processes with dual simultaneous feed of nitrogen and carbon source would presumably have to be developed. A second approach to produce rhamnolipids at low cost is to reduce substrate costs by using agroindustrial wastes from different sources in a circular bioeconomy perspective [65]. One possibility is to use products from fast pyrolysis of lignocellulosic biomass. Arnold et al. (2019) suggest that

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comparable ranges of productivities and substrate-to-biomass yields could be achieved compared to glucose or acetate [66]. Continuous use of cells as biocatalysts and reuse of living biomass have also been addressed. For example, Lotfabad et al. (2017) investigated the immobilization of P. aeruginosa MR01 cells in both consecutive batch mode and flow-through mode in terms of their suitability as a system for rhamnolipid production [67]. In contrast to production with pathogenic wild-type strains, the use of heterologous hosts represents a modern approach that can also be used to overcome the complex regulatory mechanisms of rhamnolipid biosynthesis. At the same time, the use of sugars as a carbon source can eliminate the need for tropical palm oils, which are critical to the climate and environment. In P. aeruginosa, three genes rhlA, rhlB, and rhlC encode the final steps of rhamnolipid biosynthesis (reviewed in Refs. [68]). RhlA catalyzes the condensation of two 3-hydroxyalkanoic acid for the production of 3-(3-hydroxyalkanoyloxy)alkanoic acid (HAA), while RhlB and RhlC are rhamnosyltransferases adding successive L-rhamnose residues for the production of mono- and dirhamnolipids. These three biosynthetic genes are regulated in a cell density-dependant manner by the quorum sensing regulator RhlR, whose function requires a ligand produced by the acyl-homoserine lactone synthase RhlI. Constitutive expression of the rhamnolipid biosynthetic genes into a suitable and generally considered safe alternative industrial strain makes it easy to overcome the complex regulatory mechanisms of the natural P. aeruginosa host. Here, P. putida has been mainly used for this purpose, since the two precursors dTDP-L-rhamnose and 3-hydroxyalkanoic acids are endogenously present. For example, Cha et al. (2008) cloned the rhlABRI gene cluster from P. aeruginosa EMS1 into the host strain P. putida KT2440 using a pBBR1MCS2-based plasmid and relied on the rhlR/rhlI quorum sensing-based induction system of P. aeruginosa. In this way, a putative constitutive autoinduction uncoupled from the quorum sensing regulatory system was established. The highest titers were obtained when P. putida KT2440 (pNE2) was cultured on soybean oil, reaching levels of 7.3 g/L rhamnolipid after 72 h [69]. Another study used inducible promoters for the rhlAB operon from P. aeruginosa PAO1 [70]. They reported levels of 1.5 g/L with P. putida KT42C1 grown on LB medium supplemented with glucose [70]. Setoodeh et al. (2014) introduced the rhlAB genes of P. aeruginosa ATCC 9027 into P. putida KT2440 [71]. Here, however, only maximum concentrations of up to 0.57 g/L could be achieved [71]. Anaerobic cultivation of recombinant P. stutzeri DQ1 with 3 g/L NaNO3 as electron acceptor and yeast extract was performed by Zhao et al (2014), and a maximum concentration of 3.12 g/L was reported [72]. Noll et al. (2019) discussed the potential of a ROSE-like RNA-thermometer located in the 5’UTR of native rhlAB-genes for process control. An increase of specific rhamnolipid production rates of more than 60% between 37°C and 30°C was achieved, although the authors also suggest, that also multiple metabolic effects may have contributed to this [73]. Beuker at al. (2016) investigated in situ foam fractionation for rhamnolipid production in P. putida KT2440 using the genetic construct pSynPro8oT_rhlAB based on the plasmid pBBR1MCS-3. The authors studied the enrichment of bacteria and rhamnolipids in batch cultures in the foam fraction and obtained maximum concentrations of 2.15 g/L in the foam and enrichment factors for rhamnolipids of up to 200, while maintaining biomass in the bioreactor [74]. In another study, the authors used the same genetic construct for fed-batch cultivation with glucose as the carbon source. They reported a maximum value of 14.9 g/L mono-rhamnolipid representing the highest titer reported with heterologous production strains by now [75].

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Wigneswaran et al. (2016) also utilized P. putida KT2440 and the rhlAB genes of P. aeruginosa as the production strain. The maximum reported rhamnolipid concentration was 20 mg/L. They also successfully explored the potential of using a biofilm as an alternative production system [76]. Wittgens et al. (2017) reported that RhlA and RhlB are involved in rhamnolipid biosynthesis independently and not in the form of a heterodimer. In addition, there is evidence that mono-rhamnolipids as well as HAAs can be used by cells as extracellular precursors for the synthesis of di-rhamnolipids in both P. putida and P. aeruginosa [77]. Solaiman et al. (2015) used P. chlororaphis NRRL B-30761, a natural producer of monorhamnolipids, to express the rhamnosyltransferase gene rhlC from P. aeruginosa using an expression vector (pBS29-P2-gfp) containing a P. syringae promoter [78]. An uncommon host, Burkholderia kururiensis KP23T, was used by Tavares et al. (2013) [79]. The authors went on to compare the rhamnolipid production of the wild type with an enhanced rhamnolipid producer obtained by additional expression of rhlAB genes from P. aeruginosa PAO1. Actually, several Burkholderia species produce rhamnolipids, typically with longer fatty acid chain lengths [80,81]. One noteworthy example is B. glumae, which can produce rhamnolipids, without being an human pathogenic organism like P. aeruginosa [82]. However although some efforts have been invested to improve the culture conditions, titers of natural Burkholderia producers remain low [82]. A reduction of PHA-production of about 80% was achieved by Funston et al. (2017) along with improving the rhamnolipid-titer to 3.78 g/L [83]. Cabrera-Valladares et al. (2006) reported that the availability of dTDP-L-rhamnose, a substrate of RhlB, limits the production of mono-rhamnolipids in Escherichia coli. The authors’ data suggest that the production of mono-rhamnolipids in E. coli expressing the rhlAB operon is limited by the availability of dTDP-L-rhamnose. Expression of the rmlBDAC operon from P. aeruginosa in E. coli W3110 resulted in a mono-rhamnolipid titer of 0.121 g/L [84]. In an approach of interest to metabolic and protein engineering, Han et al. (2014) replaced single amino acids of the RhlB protein. They engineered all 19 possible mutants of RhlB at position 168 and concluded that mutation of a single amino acid at position 168 in RhlB alters the volume of the binding pocket for the substrate and thus affects the selectivity of rhamnolipid formation in E. coli. However, it should be noted that the ratio of Rha-C10-C8 (as well as Rha-C8-C10) to Rha-C10-C10 was reduced to 15%–30% from an initial value of approximately 40% in the wild-type strain. Since the precursor synthesis of HAA is catalyzed by RhlA, it can be assumed that the fatty acid chain length composition depends on the specificity of RhlA. No variants with longer fatty acid chain lengths were reported. Nevertheless, these results offer an interesting perspective for the possible development of specific rhamnolipid congeners [85]. In this regard, heterologous expression of biosynthetic genes [86] and semirational evolution of biosynthetic genes [87] offer promising avenues. Although the foaming properties are requested properties of the product, overwhelming foam during the cultivation can be a problem. To circumvent this problem Jiang et al. (2020) tested the use of stop valves to break the foam bubbles [88]. They reported that the stop valve can effective break the foam, but also mention that an upscaling of this method may be difficult [88]. Another approach was reached by differ the DO and pH. Sodagari et al. (2018) reported a foam reduction with P. aeruginosa at a pH of 5.5–5.7 and DO of 10% [89]. Xu et al. (2020) report decreasing bubble size during the cultivation and therefore decreasing foam reduction efficiency by using an ex situ foam control system [90]. In another study, the foam adsorption with hydrophobic adsorbent was used, where the rhamnolipid can

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bind and the cells can be recycled to the reactor media [91]. Bator et al. (2020) used ethanol as carbon source and antifoaming agent as well as a genetically modified P. putida KT2440 [92]. The product purification of rhamnolipids from fermentation broth has been underrepresented in the bioprocess engineering literature. Extraction and precipitation methods are common here. As an interesting example, foam fractionation for the continuous recovery of rhamnolipids is noteworthy in this regard. Sarachat et al. (2010) reported a biosurfactant recovery of 97% and an enrichment ratio of 4 in the foam compared to the medium in the bioreactor [93]. By using an external foam fractionation column, a rhamnolipid concentration of 7.5 g/L in the foam and a space-time yield of 0.21 g RL/(Lh) could be achieved [93].

4. Conclusions Has the promise come true? Yes and No. The answer depends on the point of view. Surfactants are among the most widely produced and used chemicals. In view of the three major planetary emergencies, climate change, biodiversity loss, and pollution from persistent chemicals, it is important to reduce greenhouse gas emissions, replace environmentally toxic or nondegradable chemicals with environmentally safe biochemicals, and do so while increasing the quantity and quality of available food. Can biosurfactants and rhamnolipids specifically contribute to this task? Yes, definitely. But there is still a long way to go and we are still at the very beginning of the S-curve of disrupting established chemical production. In our own opinion, the known biosurfactants rhamnolipid, sophorolipids, mannosylerythritol lipids, and surfactin will remain dominant in the foreseeable future. Why? Biosurfactants that are easy to produce with wild-type organisms, such as rhamnolipid, have been well studied for decades, have proven their application potential, and can be produced by small companies without large research capacities. Heterologous rhamnolipid production such as from EVONIK industries, will remain the exception. The next few years of biosurfactant and especially rhamnolipid development will be exciting. There is still much for science to discover and for investors and industry to exploit.

References [1] Global Market Insights Inc. Biosurfactants market size and share j industry statistics—2027., 2022, https:// www.gminsights.com/industry-analysis/biosurfactants-market-report. [Accessed 23 February 2022]. [2] Kumar R, Das AJ. Rhamnolipid biosurfactant. Springer; 2018. [3] Sobero´n-Cha´vez G, Gonza´lez-Valdez A, Soto-Aceves MP, Cocotl-Yan˜ez M. Rhamnolipids produced by Pseudomonas: from molecular genetics to the market. Microb Biotechnol 2021;14:136–46. [4] Bergstr€om S, Theorell H, Davide H. Pyolipic acid, a metabolic product of Pseudomonas pyocyanea, active against Mycobacterium tuberculosis. Arch Biochem 1946;10(1):165–6. [5] Jarvis FG, Johnson MJ. A glyco-lipide produced by Pseudomonas aeruginosa. J Am Chem Soc 1949;71:12. [6] Edwards JR, Hayashi JA. Structure of a rhamnolipid from Pseudomonas aeruginosa. Arch Biochem Biophys 1965;111(2):415–21. [7] Hauser G, Karnovsky ML. Studies on the production of glycolipide by Pseudomonas aeruginosa. J Bacteriol 1954;68:645–54.

80

Chapter 4 Rhamnolipids—Has the promise come true?

[8] Hauser G, Karnovsky ML. Rhamnose and rhamnolipide biosynthesis by Pseudomonas aeruginosa. J Biol Chem 1957;224:91–105. [9] Abdel-Mawgoud AM, L
epine F, D
eziel E. Rhamnolipids: diversity of structures, microbial origins and roles. Appl Microbiol Biotechnol 2010;86:1323–36.
The involvement of rhamnolipids in microbial cell adhesion and biofilm develop[10] Nickzad A, D
eziel E. ment—an approach for control? Lett Appl Microbiol 2014;58:447–53. [11] Thakur P, Saini NK, Thakur VK, Gupta VK, Saini RV, Saini AK. Rhamnolipid the glycolipid biosurfactant: emerging trends and promising strategies in the field of biotechnology and biomedicine. Microb Cell Factories 2021;20:1–15. [12] Laabei M, Jamieson WD, Lewis SE, Diggle SP, Jenkins ATA. A new assay for rhamnolipid detection— important virulence factors of Pseudomonas aeruginosa. Appl Microbiol Biotechnol 2014;98:7199–209. [13] Kappeli O, Guerra-Santos L. US 4628030A; 1986. [14] Abdel-Mawgoud AM, L
epine F, D
eziel E. Rhamnolipids: diversity of structures, microbial origins and roles. Appl Microbiol Biotechnol 2010;86:1323–36. https://doi.org/10.1007/s00253-010-2498-2. [15] Twigg MS, Baccile N, Banat IM, D
eziel E, Marchant R, Roelants S, Bogaert V, Inge NA. Microbial biosurfactant research: time to improve the rigour in the reporting of synthesis, functional characterization and process development. Microb Biotechnol 2021;14:147–70. https://doi.org/10.1111/1751-7915.13704. [16] Crouzet J, Arguelles-Arias A, Dhondt-Cordelier S, Cordelier S, Prsˇi
c J, Hoff G, et al. Biosurfactants in plant protection against diseases: rhamnolipids and lipopeptides case study. Front Bioeng Biotechnol 2020;1014. [17] Naughton PJ, Marchant R, Naughton V, Banat IM. Microbial biosurfactants: current trends and applications in agricultural and biomedical industries. J Appl Microbiol 2019;127:12–28. [18] Chen J, Wu Q, Hua Y, Chen J, Zhang H, Wang H. Potential applications of biosurfactant rhamnolipids in agriculture and biomedicine. Appl Microbiol Biotechnol 2017;101:8309–19. [19] Anon. Biosurfactants j Jeneil Biotech., 2022, https://www.jeneilbiotech.com/biosurfactants. [Accessed 24 February 2022]. [20] Arau´jo P, Lemos M, Mergulha˜o F, Melo L, Simo˜es M. Antimicrobial resistance to disinfectants in biofilms. Sci Against Microbial Pathogens: Commun Curr Res Technol Adv 2011;3:826–34. [21] de Araujo LV, Guimara˜es CR, Da Silva Marquita RL, Santiago VMJ, de Souza MP, Nitschke M, Freire DMG. Rhamnolipid and surfactin: anti-adhesion/antibiofilm and antimicrobial effects. Food Control 2016;63:171–8. [22] Hajfarajollah H, Mehvari S, Habibian M, Mokhtarani B, Noghabi KA. Rhamnolipid biosurfactant adsorption on a plasma-treated polypropylene surface to induce antimicrobial and antiadhesive properties. RSC Adv 2015;5:33089–97. [23] Magalha˜es L, Nitschke M. Antimicrobial activity of rhamnolipids against Listeria monocytogenes and their synergistic interaction with nisin. Food Control 2013;29:138–42. [24] Meylheuc T, Renault M, Bellon-Fontaine MN. Adsorption of a biosurfactant on surfaces to enhance the disinfection of surfaces contaminated with Listeria monocytogenes. Int J Food Microbiol 2006;109:71–8. [25] Nitschke M, Arau´jo LV, Costa S, Pires RC, Zeraik AE, Fernandes A, et al. Surfactin reduces the adhesion of food-borne pathogenic bacteria to solid surfaces. Lett Appl Microbiol 2009;49:241–7. [26] Liu J-W, Wei K-H, Xu S-W, Cui J, Ma J, Xiao X-L, et al. Surfactant-enhanced remediation of oilcontaminated soil and groundwater: a review. Sci Total Environ 2021;756, 144142. [27] Fenibo EO, Ijoma GN, Selvarajan R, Chikere CB. Microbial surfactants: the next generation multifunctional biomolecules for applications in the petroleum industry and its associated environmental remediation. Microorganisms 2019;7:581. [28] Patel S, Homaei A, Patil S, Daverey A. Microbial biosurfactants for oil spill remediation: pitfalls and potentials. Appl Microbiol Biotechnol 2019;103:27–37. [29] Liu G, Zhong H, Yang X, Liu Y, Shao B, Liu Z. Advances in applications of rhamnolipids biosurfactant in environmental remediation: a review. Biotechnol Bioeng 2018;115:796–814.

References

81

[30] Agarwal A, Liu Y. Remediation technologies for oil-contaminated sediments. Mar Pollut Bull 2015;101: 483–90. [31] De Silva R, Ca´ssia FS, Almeida DG, Rufino RD, Luna JM, Santos VA, Sarubbo LA. Applications of biosurfactants in the petroleum industry and the remediation of oil spills. Int J Mol Sci 2014;15:12523–42. [32] Pacwa-Płociniczak M, Płaza GA, Piotrowska-Seget Z, Cameotra SS. Environmental applications of biosurfactants: recent advances. Int J Mol Sci 2011;12:633–54. [33] Liu Y, Zeng G, Zhong H, Wang Z, Liu Z, Cheng M, et al. Effect of rhamnolipid solubilization on hexadecane bioavailability: enhancement or reduction? J Hazard Mater 2017;322:394–401. [34] Zhong H, Liu Y, Liu Z, Jiang Y, Tan F, Zeng G, et al. Degradation of pseudo-solubilized and mass hexadecane by a Pseudomonas aeruginosa with treatment of rhamnolipid biosurfactant. Int Biodeterior Biodegradation 2014;94:152–9. [35] Zhong H, Liu G, Jiang Y, Yang J, Liu Y, Yang X, et al. Transport of bacteria in porous media and its enhancement by surfactants for bioaugmentation: a review. Biotechnol Adv 2017;35:490–504. [36] Zeng Z, Liu Y, Zhong H, Xiao R, Zeng G, Liu Z, et al. Mechanisms for rhamnolipids-mediated biodegradation of hydrophobic organic compounds. Sci Total Environ 2018;634:1–11. [37] Olasanmi IO, Thring RW. Evaluating rhamnolipid-enhanced washing as a first step in remediation of drill cuttings and petroleum-contaminated soils. J Adv Res 2020;21:79–90. [38] Mao X, Jiang R, Xiao W, Yu J. Use of surfactants for the remediation of contaminated soils: a review. J Hazard Mater 2015;285:419–35. [39] Sakthipriya N, Kumar G, Agrawal A, Doble M, Sangwai JS. Impact of biosurfactants, surfactin, and rhamnolipid produced from Bacillus subtilis and Pseudomonas aeruginosa, on the enhanced recovery of crude oil and its comparison with commercial surfactants. Energy Fuel 2021;35:9883–93. [40] Rocha VAL, de Castilho LVA, de Castro RPV, Teixeira DB, Magalha˜es AV, Gomez JGC, Freire DMG. Comparison of mono-rhamnolipids and di-rhamnolipids on microbial enhanced oil recovery (MEOR) applications. Biotechnol Prog 2020;36, e2981. [41] C^amara JMDA, Sousa MASB, Barros Neto EL, Oliveira MCA. Application of rhamnolipid biosurfactant produced by Pseudomonas aeruginosa in microbial-enhanced oil recovery (MEOR). J Petrol Explorat Product Technol 2019;9:2333–41. [42] Sharma R, Singh J, Verma N. Optimization of rhamnolipid production from Pseudomonas aeruginosa PBS towards application for microbial enhanced oil recovery. 3 Biotech 2018;8:1–15. [43] Cui QF, Zheng WT, Yu L, Xiu JL, Zhang ZZ, Luo YJ, Sun SS. Emulsifying action of Pseudomonas aeruginosa L6-1 and its metabolite with crude oil for oil recovery enhancement. Pet Sci Technol 2017;35:1174–9. [44] Lourith N, Kanlayavattanakul M. Natural surfactants used in cosmetics: glycolipids. Int J Cosmet Sci 2009;31:255–61. [45] Adu SA, Naughton PJ, Marchant R, Banat IM. Microbial biosurfactants in cosmetic and personal skincare pharmaceutical formulations. Pharmaceutics 2020;12:1099. [46] Maier RM, Soberon-Chavez G. Pseudomonas aeruginosa rhamnolipids: biosynthesis and potential applications. Appl Microbiol Biotechnol 2000;54:625–33. [47] Kosaric N, Sukan FV. Biosurfactants. CRC Press; 1993. [48] Vasileva-Tonkova E, Galabova D, Karpenko E, Shulga A. Biosurfactant-rhamnolipid effects on yeast cells. Lett Appl Microbiol 2001;33:280–4. [49] Haba E, Pinazo A, Jauregui O, Espuny MJ, Infante MR, Manresa A. Physicochemical characterization and antimicrobial properties of rhamnolipids produced by Pseudomonas aeruginosa 47T2 NCBIM 40044. Biotechnol Bioeng 2003;81:316–22. [50] Ishigami Y, Gama Y, Nagahara H, Motomiya T, Yamaguchi M. Liposome containing rhamnolipids. Japanese Patent Kokai 1988;29:63–182. [51] Piljac T, Piljac G. Use of rhamnolipids in wound healing, treating burn shock, atherosclerosis, organ transplants, depression, schizophrenia and cosmetics (European Patent 1 889 623). New York: Paradigm Biomedical Inc; 1999.

82

[52] [53] [54] [55] [56] [57] [58] [59] [60]

[61] [62]

[63]

[64]

[65] [66] [67]

[68]

[69] [70]

[71]

[72]

Chapter 4 Rhamnolipids—Has the promise come true?

Desanto K. Rhamnolipid-based formulations. World Patent WO 2008013899 A3; 2008. Anon. Unilever and Evonik partner to launch green cleaning ingredient. Unilever PLC; 2019. Anon. We’re reimagining the future of cleaning. Unilever PLC; 2020. Nitschke M, Costa S. Biosurfactants in food industry. Trends Food Sci Technol 2007;18:252–9. Sharma D. Biosurfactants in food. Springer; 2016. Nitschke M, Silva SSE. Recent food applications of microbial surfactants. Crit Rev Food Sci Nutr 2018;58:631–8. Ingrid Paul Hilda Van Haesendonck, Mechelen (BE), Emmanuel Claude Albert Vanzeveren, Brussels (BE). US20060233935A1. Gandhi NR, Skebba VLP. Rhamnolipid compositions and related methods of use 2011: Google Patents. Thakur P, Saini NK, Thakur VK, Gupta VK, Saini RV, Saini AK. Rhamnolipid the glycolipid biosurfactant: emerging trends and promising strategies in the field of biotechnology and biomedicine. Microb Cell Factories 2021;20:1–15. Giani C, Wullbrandt D, Rothert R, Meiwes J. Pseudomonas aeruginosa and its use in a process for the biotechnological preparation of L-Rhamnose; 1997. Schmidberger A, Henkel M, Hausmann R, Schwartz T. Expression of genes involved in rhamnolipid synthesis in Pseudomonas aeruginosa PAO1 in a bioreactor cultivation. Appl Microbiol Biotechnol 2013;97:5779–91. Henkel M, Schmidberger A, K€uhnert C, Beuker J, Bernard T, Schwartz T, et al. Kinetic modeling of the time course of N-butyryl-homoserine lactone concentration during batch cultivations of Pseudomonas aeruginosa PAO1. Appl Microbiol Biotechnol 2013;97:7607–16. Henkel M, Schmidberger A, Vogelbacher M, K€ uhnert C, Beuker J, Bernard T, et al. Kinetic modeling of rhamnolipid production by Pseudomonas aeruginosa PAO1 including cell density-dependent regulation. Appl Microbiol Biotechnol 2014;98:7013–25. Gudin˜a EJ, Rodrigues AI, de Freitas V, Azevedo Z, Teixeira JA, Rodrigues LR. Valorization of agroindustrial wastes towards the production of rhamnolipids. Bioresour Technol 2016;212:144–50. Arnold S, Henkel M, Wanger J, Wittgens A, Rosenau F, Hausmann R. Heterologous rhamnolipid biosynthesis by P. putida KT2440 on bio-oil derived small organic acids and fractions. AMB Express 2019;9:1–7. Lotfabad TB, Ebadipour N, Roostaazad R, Partovi M, Bahmaei M. Two schemes for production of biosurfactant from Pseudomonas aeruginosa MR01: applying residues from soybean oil industry and silica sol-gel immobilized cells. Colloids Surf B: Biointerfaces 2017;152:159–68. Abdel-Mawgoud A, Hausmann R, L
epine F, M€uller M, D
eziel E. Rhamnolipids: detection, analysis, biosynthesis, genetic regulation, and bioengineering of production. Biosurfactants 2010;13–55. https://doi.org/ 10.1007/978-3-642-14490-5_2. Cha M, Lee N, Kim M, Kim M, Lee S. Heterologous production of Pseudomonas aeruginosa EMS1 biosurfactant in Pseudomonas putida. Bioresour Technol 2008;99:2192–9. Wittgens A, Tiso T, Arndt TT, Wenk P, Hemmerich J, M€ uller C, et al. Growth independent rhamnolipid production from glucose using the non-pathogenic Pseudomonas putida KT2440. Microb Cell Factories 2011;10:1–18. Setoodeh P, Jahanmiri A, Eslamloueyan R, Niazi A, Ayatollahi SS, Aram F, et al. Statistical screening of medium components for recombinant production of Pseudomonas aeruginosa ATCC 9027 rhamnolipids by nonpathogenic cell factory Pseudomonas putida KT2440. Mol Biotechnol 2014;56:175–91. Zhao F, Mandlaa M, Hao J, Liang X, Shi R, Han S, Zhang Y. Optimization of culture medium for anaerobic production of rhamnolipid by recombinant P. seudomonas stutzeri R hl for microbial enhanced oil recovery. Lett Appl Microbiol 2014;59:231–7.

References

83

[73] Noll P, Treinen C, M€uller S, Senkalla S, Lilge L, Hausmann R, Henkel M. Evaluating temperature-induced regulation of a ROSE-like RNA-thermometer for heterologous rhamnolipid production in Pseudomonas putida KT2440. AMB Express 2019;9:1–10. [74] Beuker J, Steier A, Wittgens A, Rosenau F, Henkel M, Hausmann R. Integrated foam fractionation for heterologous rhamnolipid production with recombinant Pseudomonas putida in a bioreactor. AMB Express 2016;6:1–10. [75] Beuker J, Barth T, Steier A, Wittgens A, Rosenau F, Henkel M, Hausmann R. High titer heterologous rhamnolipid production. AMB Express 2016;6:1–7. [76] Wigneswaran V, Nielsen KF, Sternberg C, Jensen PR, Folkesson A, Jelsbak L. Biofilm as a production platform for heterologous production of rhamnolipids by the non-pathogenic strain Pseudomonas putida KT2440. Microb Cell Factories 2016;15:1–13. [77] Wittgens A, Kovacic F, M€uller MM, Gerlitzki M, Santiago-Sch€ ubel B, Hofmann D, et al. Novel insights into biosynthesis and uptake of rhamnolipids and their precursors. Appl Microbiol Biotechnol 2017;101: 2865–78. [78] Solaiman DKY, Ashby RD, Gunther NW, Zerkowski JA. Dirhamnose-lipid production by recombinant nonpathogenic bacterium Pseudomonas chlororaphis. Appl Microbiol Biotechnol 2015;99:4333–42. [79] Tavares LFD, Silva PM, Junqueira M, Mariano DCO, Nogueira F, Domont GB, et al. Characterization of rhamnolipids produced by wild-type and engineered Burkholderia kururiensis. Appl Microbiol Biotechnol 2013;97:1909–21. [80] H€auβler S, Rohde M, von Neuhoff N, Nimtz M, Steinmetz I. Structural and functional cellular changes induced by Burkholderia pseudomallei rhamnolipid. Infect Immun 2003;71:2970–5. [81] Dubeau D, D
eziel E, Woods DE, L
epine F. Burkholderia thailandensis harbors two identical rhl gene clusters responsible for the biosynthesis of rhamnolipids. BMC Microbiol 2009;9:1–12. [82] Nickzad A, Guertin C, D
eziel E. Culture medium optimization for production of rhamnolipids by Burkholderia glumae. Colloids Interfaces 2018;2:49. [83] Funston SJ, Tsaousi K, Smyth TJ, Twigg MS, Marchant R, Banat IM. Enhanced rhamnolipid production in Burkholderia thailandensis transposon knockout strains deficient in polyhydroxyalkanoate (PHA) synthesis. Appl Microbiol Biotechnol 2017;101:8443–54. [84] Cabrera-Valladares N, Richardson A-P, Olvera C, Trevin˜o LG, D
eziel E, L
epine F, Sobero´n-Cha´vez G. Monorhamnolipids and 3-(3-hydroxyalkanoyloxy) alkanoic acids (HAAs) production using Escherichia coli as a heterologous host. Appl Microbiol Biotechnol 2006;73:187–94. [85] Han L, Liu P, Peng Y, Lin J, Wang Q, Ma Y. Engineering the biosynthesis of novel rhamnolipids in Escherichia coli for enhanced oil recovery. J Appl Microbiol 2014;117:139–50. https://doi.org/10.1111/jam.12515. [86] Wittgens A, Santiago-Schuebel B, Henkel M, Tiso T, Blank LM, Hausmann R, et al. Heterologous production of long-chain rhamnolipids from Burkholderia glumae in Pseudomonas putida—a step forward to tailormade rhamnolipids. Appl Microbiol Biotechnol 2018;102:1229–39. https://doi.org/10.1007/s00253-0178702-x. [87] Dulcey CE, de Los L, Santos Y, L
etourneau M, D
eziel E, Doucet N. Semi-rational evolution of the 3-(3hydroxyalkanoyloxy)alkanoate (HAA) synthase RhlA to improve rhamnolipid production in Pseudomonas aeruginosa and Burkholderia glumae. FEBS J 2019;286:4036–59. https://doi.org/10.1111/febs.14954. [88] Jiang J, Zu Y, Li X, Meng Q, Long X. Recent progress towards industrial rhamnolipids fermentation: process optimization and foam control. Bioresour Technol 2020;298, 122394. [89] Sodagari M, Invally K, Ju L-K. Maximize rhamnolipid production with low foaming and high yield. Enzym Microb Technol 2018;110:79–86.

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[90] Xu N, Liu S, Xu L, Zhou J, Xin F, Zhang W, et al. Enhanced rhamnolipids production using a novel bioreactor system based on integrated foam-control and repeated fed-batch fermentation strategy. Biotechnol Biofuels 2020;13:1–10. [91] Anic I, Apolonia I, Franco P, Wichmann R. Production of rhamnolipids by integrated foam adsorption in a bioreactor system. AMB Express 2018;8:1–12. [92] Bator I, Karmainski T, Tiso T, Blank LM. Killing two birds with one stone-strain engineering facilitates the development of a unique rhamnolipid production process. Front Bioeng Biotechnol 2020;899. [93] Sarachat T, Pornsunthorntawee O, Chavadej S, Rujiravanit R. Purification and concentration of a rhamnolipid biosurfactant produced by Pseudomonas aeruginosa SP4 using foam fractionation. Bioresour Technol 2010;101:324–30. https://doi.org/10.1016/j.biortech.2009.08.012.

CHAPTER

Biosurfactants as food additives: New trends and applications

5

Tathiane Ferroni Passosa, Paula de Camargo Bertusob, and Marcia Nitschkea,b a

Sa˜o Carlos Institute of Chemistry, University of Sa˜o Paulo, Sa˜o Carlos, SP, Brazil, bInterunits Graduate Program in Bioengineering, University of Sa˜o Paulo, Sa˜o Carlos, SP, Brazil

1. Biosurfactants in food formulation The development of industrialized products generates the need to introduce additives in food, and one of the main challenges when inserting a new element in food formulation is maintaining its organoleptic characteristics. However, beyond the sensory aspect, nutritional value, appearance, and safety must be considered when using an additive [1,2]. According to the European Food Safety Authority (EFSA), food additives are substances added to food formulations intentionally to perform a certain function. They can be characterized as preservatives, nutritional additives such as vitamins, fibers, and amino acids, flavoring agents, coloring agents, and texturing agents, among others [2]. The growing concern with conscious consumption, linking health and the environment, has led to the need of replacing synthetic additives with natural additives [3]. In this context, the food industry has explored the use of biosurfactants (BS) as food additives since the US Environment Protection Agency has approved the use of some types of BS in food products and other industrial applications [1]. A summary of the potential application of BS in food is illustrated in Fig. 1. BS such as lipopeptides, glycolipids, and lipoproteins can be isolated from plants or produced by certain species of microorganisms, such as bacteria, yeasts, or fungi [4,5]. In addition, the inherent biodegradability and sustainable nature of BS fulfill the current demand of the market [3]. Another advantage of using BS as food additives comprises their resistance to temperature variations, stability in acidic media and high salinity, typical conditions observed in the food environment [6]. For example, a glycolipid BS from Streptomyces sp. presented stability in a wide pH range (between 5.0 and 9.0) and NaCl concentration (1.5% w/v), which allows the maintenance of the molecular structure and the physicochemical properties, influencing the quality of the final product [7]. Furthermore, the diversity of BS structures allows an assertive selection of biomolecules according to the required application [8]. In addition to their surface-active properties, BS have been reported to improve texture and stability in doughs, to avoid separation of oil-based products, help in mixing ingredients, improve viscosity, and reduce energy value by replacing fats [1,8]. Rhamnolipids (RL) from Pseudomonas spp. are one of the most exploited BS in food. They have been included in formulations of flour-based doughs such as bread, pizzas and cakes, buttery creams, and in fresh or frozen products. More specifically, in ice cream and bakery products, RL can be used in consistency control, fat stabilization, and aging reduction [1]. The literature also reports the use of BS as food additives for the solubilization of aromatic oils for Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00005-3 Copyright # 2023 Elsevier Inc. All rights reserved.

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FIG. 1 General overview on prospective uses of BS in food.

flavor enhancement [8]. Most studies regarding the application of BS in food formulations involve their use as preservatives, where they act as antimicrobial or antioxidant agents, as well as emulsifiers. Some recent examples on such topic are further discussed.

1.1 BS as emulsifiers Texturizing agents, which act as emulsifiers and stabilizers, are applied to modify texture and represent an important class of food additives [2,9]. Furthermore, emulsions are key systems for solubilizing and dispersing hydrophobic components in water or hydrophilic components in nonpolar media, usually present in food formulations [2]. An emulsion is formed when two immiscible liquids are mixed and the most common type is a water-oil emulsion, which consists of microscopic drops of oil in water. Emulsions obtained through stirring are usually very unstable and can lead to phase separation even in short periods. Emulsifiers play an important role in the initial phase of the preparation of an emulsion, where they assist in the formation and stabilization of agglomerates during the homogenization of the mixture. Also, they tend to minimize phase separation due to their ability to control clusters, which favors the stability of heterogeneous systems [8,10]. The most common emulsifiers are surfactants, which reduce the interfacial tension, and consequently the free energy of the interface between the two liquids, increasing the stability of the emulsion by controlling the agglomeration of the different phases of the mixture [10]. Within this context, BS have drawn the attention of food industries aiming to reduce the use of synthetic emulsifiers or to avoid

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the use of soy emulsifiers derived from genetically modified plants [8]. Some BS have been described as good emulsifiers in systems composed of two or more immiscible phases, and in such cases, they are called bioemulsifiers [8,9]. Studies have reported improvements in doughs, concerning texture, volume, and conservation, after the addition of BS, especially in baked products. For example, a BS obtained from Candida bombicola was evaluated as an emulsifier in the formulation of cupcakes. The authors concluded that the emulsions formed by BS showed greater stability, and the cupcakes did not show physicochemical differences when compared to the standard formulation, suggesting its use as an alternative to vegetable fat [9]. Likewise, a lipopeptide obtained from the marine actinobacterium Nesterenkonia sp. incorporated into a muffin dough showed an improvement in organoleptic characteristics and an increase in softness when compared to the control [11]. Another BS isolated from Candida utilis, was applied in the formulation of cookies replacing animal fat. The final product was softer than the control without changing the physicochemical properties [12]. Supplementation of wheat flour by up to 30% with a BS from Bacillus subtilis improved the texture of sesame biscuit dough, leading to greater softness and ease of shaping product comparatively to the use of the commercial emulsifier glycerol monostearate [13]. In addition, an analogous BS, obtained from the same species of microorganism, also resulted in an improvement in bread dough, when incorporated at a concentration of 0.075% (w/w) [14]. The partial or complete replacement of egg yolks in cookie dough by a glycolipid BS, obtained from Saccharomyces cerevisiae, did not result in changes in texture or flavor after baking, maintaining its elasticity and firmness when compared to the standard formulation [15]. Some research described the use of microbial BS as emulsifiers for mayonnaise and salad dressings to improve consistency and texture. A bioemulsifier synthesized by C. utilis, improved the stability of salad dressing maintaining its emulsifying capacity, after one month of storage, even under extreme conditions of pH, temperature, and salinity [16]. These examples illustrate that BS can act in the same way as conventional emulsifiers, leading to a green alternative for additives in food formulation (Table 1). Moreover, BS have an additional advantage, since they can also inhibit the growth of several important foodborne pathogens [2].

1.2 Food preservatives: BS as antimicrobial agents Besides emulsifier and stabilizer properties, BS can increase the shelf life of foods as they significantly contribute to the prevention of microbiological contamination. Several reports in the literature have highlighted the antimicrobial properties of different types of BS against a broad spectrum of food pathogens [2]. RL have been exploited as alternatives to control the growth of microbes in foods. Ferreira et al. [17], reported that the antimicrobial efficacy of RL against food pathogens Listeria monocytogenes, Bacillus cereus, and Staphylococcus aureus was pH-dependent and improved at acidic conditions suggesting a good potential to avoid such pathogen contamination in low-acid or acid food. In addition, RL obtained from Pseudomonas plecoglossicida showed antimicrobial activity against S. aureus, Escherichia coli, and Aeromonas hydrophila. The authors reported a dose-dependent relationship, with the greatest inhibition effect being observed at 100 μg/mL of RL [18]. Rodrigues et al. [19], reported increasing antifungal activity of RL against Aspergillus niger and Aspergillus carbonarius in the presence of sodium chloride. Combination of RL-NaCl (3 g/L to 875 mM, for A. niger and 3 g/L to 375 mM

Table 1 Examples on recent food-related applications of microbial surfactants. BS application

Food/target/purpose

BS origin/type

Properties

Reference

Food formulation

Cookies Bread Cupcake Muffin Salad dressing Endospore-forming bacteria in milk B. cereus, S. aureus, A. niger L. monocytogenes, B. cereus, S. aureus S. aureus, E. coli, Aeromonas hydrophila E. coli, B. cereus, S. aureus S. aureus, P. aeruginosa, E. coli

C. utilis B. subtilis C. bombicola Nesterenkonia sp. C. utilis Dacryopinax spathularia Streptomyces sp. RL

Emulsifying antioxidant Emulsifying Emulsifying Emulsifying, antioxidant Emulsifying Antimicrobial Antimicrobial Antimicrobial

[12] [14] [9] [11] [16] [29] [7] [17]

Antimicrobial

[18]

Emulsifying, Antimicrobial Antimicrobial

[23] [28]

A. niger, A. carbonarius E. coli E. coli S. aureus S. aureus

Pseudomonas plecoglossicida RL Lactobacillus sp. Brachybacterium paraconglomeratum P. aeruginosa RL Lactobacillus sp. Bacillus licheniformis MEL BS RL

[19] [22] [47] [49] [50]

Orange Ultrafiltration membranes Ketchup, chocolate, soybean oil Cabbage, carrot, lettuce Vibrio cholerae S. aureus Curcumin Lutein L. monocytogenes, E. coli O157:H7 Nisin

RL-chitosan film RL Surfactin Bacillus sp. lipopeptide SL-gold NP RL-Chitosan NP RL-zein-alginate NP SL-zein NP Surfactin-Cinnamaldehyde NE RL-nisin liposomes

Antimicrobial Antibiofilm, antimicrobial Antibiofilm Antiadhesive, antibiofilm Disruption of milk-based biofilms Antioxidant, antimicrobial Washing antifouling Removal of food stain Heavy metals removal Antimicrobial Antibiofilm Nutraceutical delivery Encapsulation and delivery Antimicrobial Antimicrobial and active packaging

Control food pathogens

Edible coating Cleaning agent

Nanotechnology

NP, nanoparticles; NE, nanoemulsions; MEL, mannosylerythritol lipids; RL, rhamnolipids; SL, sophorolipids.

[20] [60] [63] [70] [99] [101] [104] [106] [111] [115,116]

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for A. carbonarius) resulted in 100% growth inhibition, suggesting a direct relationship between ionic strength and RL activity that can be exploited in salted food products. The combination of BS with natural biopolymers is an interesting approach to improve food safety. An edible coating formulated from a mixture of chitosan (2% w/v) and RL (2% w/v) was applied to extend the shelf life of sweet oranges stored at 25°C for 8 weeks. Authors demonstrated that chitosanRL films prevented the growth of spoilage microorganisms and maintained the physicochemical and sensory attributes of the fruits, suggesting they can replace chemical-based coatings for the prevention of postharvest loss in fruit quality [20]. BS obtained from opportunistic pathogenic bacteria, such as Pseudomonas aeruginosa, is still restricted to use in food. Thus, BS from Lactobacillus and yeasts, microorganisms often present in food, represent safer alternatives for industries [21]. A BS produced by Lactobacillus sp. showed antimicrobial activity against E. coli, and its efficacy in inhibiting microbial growth was similar to sodium dodecyl sulfate (SDS) a conventional synthetic surfactant [22]. Another BS obtained from Lactobacillus sp., isolated from Romanian traditional fermented food products, also showed inhibitory activity against E. coli [23]. Mandhu and Prapulla [24], demonstrated that a BS obtained from L. plantarum was effective against strains of E. coli, Yersinia enterocolitica, and S. aureus. Inhibition of microbial growth was dependent on the applied concentration, and the highest antimicrobial activity was observed at 25 mg/mL of the BS. In the same context, Mouafo et al. [25], and Hippolyte et al. [26], described antimicrobial efficacy of BS isolated from Lactobacillus sp. against food pathogens such as Bacillus sp., E. coli, Pseudomonas sp., Salmonella sp., and S. aureus. The antimicrobial potential of BS synthesized by unusual species of microorganisms are also described in the literature. Balan et al. [27], reported the production of a glycolipid obtained from the marine yeast Cyberlindnera saturnus able to completely inhibit the growth of B. cereus and E. coli at a concentration of 200 μg/mL. Similarly, a BS obtained from the marine actinobacterium Brachybacterium paraconglomeratum, showed bactericidal activity against a wide spectrum of microorganisms including S. aureus, P. aeruginosa, and E. coli [28]. The antimicrobial activity of a BS from Streptomyces sp. against B. cereus, S. aureus, and A. niger was comparable to nystatin and surfactin [7]. In a recent study, a glycolipid from the edible jelly fungus Dacryopinax spathularia was reported to be effective in controlling the growth and germination of spore-forming dairy-spoilage bacteria extending the shelf life of milk [29]. Although not yet fully elucidated, the antimicrobial action of BS has been associated with its interaction with the cell envelope. According to Sotirova et al. [30], RL can denature proteins and solubilize lipids present in the cell membrane, resulting in the formation of transmembrane pores, which facilitate the entry/leakage of molecules leading to cell lysis. Such hypothesis is supported by morphological changes observed in cell surface when treated with BS (Fig. 2) and is described in numerous reports in the literature.

1.3 Antioxidants Trends in healthy eating and the increasing progress in the search for functional natural foods represent an opportunity for the application of BS, especially those that exhibit antioxidant activity [5]. Autooxidation of lipids reduces food quality and safety, as it generates toxic compounds and leads to rancidity, resulting in unpleasant tastes. Antioxidant agents are used to avoid the process of oxidative

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FIG. 2 SEM images of Bacillus cereus cells before (left) and after (right) contact with RL solution (19.5 μg/mL) during 2 h. Cell damage is noticeable for the RL-treated sample compared to the control.

deterioration of lipids, increasing stability of foods, in addition to preventing degenerative and heart diseases [2]. The antioxidant activity of BS has been related to the presence of fatty acid molecules containing unsaturation [8]. A lipopeptide obtained from Nesterenkonia sp., which has unsaturation in its hydrophobic moiety, presented good antioxidant activity at a concentration of 6 mg/mL [11]. Other BS have also been reported as potential antioxidant agents, for example: C. utilis BS showed antioxidant capacity at concentrations below those considered toxic, and the effect was concentration-dependent [12]; a Lactobacillus casei BS showed antioxidant activity by scavenging of free radicals, reaching the best results at a concentration of 5 mg/mL [31]; Adetunji et al. [20], demonstrated that chitosan-RL coating films were also able to inhibit the production of superoxide free radicals. Although antioxidant properties of BS have not been widely explored, they represent a prospective area for the development of novel products based on extracellular enzymatic synthesis of tailor-made BS [32].

2. Use of BS in food processing 2.1 BS as antiadhesive and antibiofilm agents Biofilms are defined as a complex community of microorganisms adhered to each other or to a surface, that are surrounded by a self-produced extracellular polymeric material. When forming part of a biofilm, microbial cells show diverse physiological characteristics compared to their planktonic (freeliving) form [33–36]. One of the most important differences is the enhanced resistance of biofilms to sanitizers, antibiotics, host defenses, and shear forces since polymeric matrix act as a protective barrier to the adhered cells [34,37–40]. Biofilm-forming species may present genomic variations in key

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genes for biofilm formation and maturation, resulting in different characteristics (thickness, stability, and architecture) according to the environmental conditions and increasing the possibility of antimicrobial resistance [41,42]. Biofilm resistance also favors microbial persistence in the food industry and in food manipulation environments. If present in processing plants and factory equipment, biofilms from food pathogens can secrete toxins or liberate cells, which may contaminate the products and cause either intoxication or infection on consumers [41] thus, they represent a great concern to food industry and health authorities [40]. In addition, biofilms promote food spoilage, increase in the cost of the cleaning procedures, and may cause production line breaks [41,43]. This scenario reinforces the need for novel strategies for preventing biofilm formation and disrupt preexisting biofilms, especially considering the fact that most disinfectants used in food processing plants are reported not to be efficient against mature biofilms [43]. In this sense, BS have been described as potential alternatives to control adhesion and biofilms of microorganisms usually associated with contaminated food. Surfactin produced by B. subtilis was able to reduce S. aureus and L. monocytogenes adhesion to polystyrene samples up to 66% when applied under different temperatures (35°C, 25°C, and 4°C) [44]. Preconditioning of polystyrene and stainless steel with surfactin and RL inhibits significantly both the adhesion and biofilm formation for L. monocytogenes and Pseudomonas fluorescens strains [45]. The lipopeptide lichenysin obtained from Bacillus licheniformis was tested for its antiadhesive and biofilm disruption activity against several bacteria and yeast, including some food pathogens such as E. coli, L. monocytogenes, and S. aureus [46]. Lichenysin was able to prevent adhesion in a polystyrene surface up to 69% for S. aureus, 50% for E. coli, and 48% for L. monocytogenes; the biofilm disruption capacity of this BS was around 50% [46]. Giri et al. [47] have also reported the antibiofilm activity from a B. licheniformis derived BS against E. coli. The study also demonstrated that the BS was biocompatible and boosted immune responses against pathogens in fish, this feature could help improve the quality of food livestock and decrease the economic losses in industry. Kiran et al. [11], described a lipopeptide produced by a marine bacteria able to inhibit around 90% of S. aureus biofilm formation. Besides the antibiofilm activity, this BS showed no cytotoxicity when tested on shrimp and was successfully incorporated into a bakery recipe, illustrating its potential applicability in the food industry. Sophorolipids (SL) were successfully applied to disrupt mature biofilms of a mixed culture of B. subtilis and S. aureus [48]. The BS mannosylerythritol lipids (MEL) have also shown activity against S. aureus, acting as an antiadhesive preventing the attachment of cells on tested surfaces and impairing biofilm survival on glass, polystyrene, and stainless-steel surfaces [49]. Most studies regarding antibiofilm/antiadhesive activity of BS are performed in culture medium; however, it is important to evaluate their efficiency using food models as growing substrates. In this view, Silva et al. [50] reported that RL are effective on removing S. aureus biofilms established on polystyrene surfaces in the presence of skim milk. The milk-based biofilms were removed up to 89% after 2 h contact with BS solution at different concentrations and temperatures, suggesting good potential as cleaning agents in dairy processing plants. The mechanisms involved in antiadhesive and biofilm disruption activity (Fig. 3B) demonstrated by BS has been studied. Araujo et al. [45], reported that when coated with RL, polystyrene surfaces showed a decrease in hydrophobicity and an increase on acid character. Similar results were observed when surfactin was used to coat stainless steel surfaces [45]. The authors state that those physicochemical changes promoted by BS are partially responsible for the antiadhesive effect observed. Other

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FIG. 3 Conceptual image of the effects promoted by BS in (A) Emulsion formation ; (B) cleaning of food processing pipelines; (C) removal of heavy metals from vegetables.

factors likely involved in the antiadhesive activity is the quorum sensing signaling molecules and the stress caused by the BS, as well as the nature of the surface and medium and presence of cell appendages, among others [51]. When considering preexisting biofilms, on the other hand, BS may act by changing the physical properties of the biofilm surface and reducing the bacteria-surface interactions or interfering with key proteins present on the extracellular matrix [52]. To disrupt a biofilm, the surfactants need to penetrate the matrix, adsorb, and reduce interfacial tension between surface and biofilm matrix; decrease attractive forces involved in microbial adhesion [53] and/or improve the solubility of matrix favoring their removal to aqueous solution [2]. Among key proteins present in biofilms, TasA is responsible for producing amyloid fibers able to link cells inside the biofilm and when deleted, the development of strong biofilm of B. subtilis, B. cereus, and E. coli is impaired [54,55]. Some antibiofilm drugs were reported to act by interfering with the correct assembly of TasA and thus destroying preformed biofilms [55]. Gene regulation is an important factor affecting BS activity on biofilms. Lactic acid bacteriaderived BS were capable of inhibiting S. aureus adhesion to polystyrene and also showed an antibiofilm effect by reducing significantly cell growth [56]. The authors reported that several genes related to biofilm formation in S. aureus such as dltB, sarA, sortaseA, agrA, icaA, and cidA were down-regulated after exposure to these BS [56] suggesting their potential to control S. aureus biofilms. Cramton et al. [57] first demonstrated that deleting the ica locus eliminates biofilm formation by impairing the

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production of adhesins thus, cells are not able to adhere to one another to form the layers of the biofilm [57,58]. Mutations in the sarA gene were also reported to impair biofilm formation in S. aureus, in some conditions inhibiting its formation by 80% [59]. Knowledge about gene regulation involved on adhesion and biofilm formation when cells are treated with BS, can help to elucidate their mechanism of action at molecular level.

2.2 BS as cleaning agents As described in the previous section, several BS are reported to act in preventing formation and disrupting preexisting biofilms, especially on plastic, glass and stainless-steel surfaces, that are commonly found on food processing facilities. This suggests that those BS can also be utilized as cleaning agents. For the majority of food processing plants, sterilization is not possible nor cost-effective so different sanitation and disinfection protocols are applied to reduce the presence of harmful fungi and/or bacteria and prevent them to acquire resistance [43]. One of the major problems in dairy industry is the membrane fouling caused by the large amount of proteins found on whey, especially in processes involving ultra-filtration [60]. Long et al. [61], evaluated the potential of RL in cleaning different types of membranes (polysulfone-g-poly-ethylene glycol, polysulfone, polyacrylonitrile, and polyethersulfone). The BS washing showed no damage to the membrane structure and was capable to restore the membrane flux to 95.5%, a greater value when compared to chemical surfactants and a commercial membrane cleaner (from 51% to 76%). Aghajani et al. [60], also evaluated the potential of RL as an antifouling agent. According to the authors, RL were chosen for this study because they are biodegradable and eco-friendly and showed high membrane cleaning efficiency with a less concentrated solution when compared to other chemicals (such as NaOH and SDS). Similarly to a previous report [61], the authors stated that the use of RL did not interfere with the membrane structure and obtained a flux recovery of 100% [60]. Surfactin has also been reported to efficiently clean polysthersulfonate membranes clogged with bovine serum albumin achieving a 97% of flux recovery when compared to new membranes [62]. Research on BS have also reported their use as detergents, especially for laundry purposes [63–66] nevertheless, some results can be extended to food applications. A BS produced by Ochrabactrum intermedium was capable of removing 60% of an olive oil stain when tested alone and 82% when combined with a commercial detergent [67]. Surfactin produced by B. subtilis completely remove various stains derived from food products, such as ketchup, chocolate, and yogurt, and showed high removal values for motor oil, corn oil and soybean oil, comparable to the commercial laundry detergent [63]. The same authors reported that surfactin improved the performance of commercially available detergents, produced no skin irritation, showed high biodegradability and was classified as a low toxicity level comparatively to synthetic surfactants [63]. These features make BS desirable for food industry applications, especially on surfaces that have direct contact with food products, since in the circumstance of incomplete removal, the BS won’t represent any harm to the final consumers. RL produced by P. aeruginosa was incorporated in a liquid detergent formulation and its performance was compared to commercially available detergents. RL-based detergent showed slightly smaller detergency values than commercial ones, but it produced less foam, which provides efficient cleaning with a low water usage [68]. In addition, this BS was reported as a potential additive to enhance the cleaning power of detergents [68].

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Besides cleaning equipment surfaces, BS have also been reported to sanitize food itself. A glycolipid from Bacillus sp. was applied to remove cadmium contamination from potato, garlic, radish, and onion achieving a maximum removal of 73% [69]. The vegetables were washed thoroughly with water, sliced, and then exposed to cadmium chloride solutions (0.4 and 0.5 mg/mL) for 30 min. Further, samples were treated with BS solution and change in cadmium concentration was measured by absorbance [69]. Similarly, a BS produced by B. licheniformis showed 61% removal of cadmium from carrots, radishes, ginger, and potatoes [47]. Another lipopeptide produced by Bacillus sp. was also used to treat different vegetables samples (cabbage, carrot, and lettuce) and it was capable of reducing the concentrations of copper, lead, cadmium, zinc, and nickel from 59% up to 87% [70]. Pieces of the tested vegetables were placed on either BS solution or water for 20 min and, after sample preparation, the heavy metal content was analyzed by atomic absorption spectroscopy [70]. These authors suggested that the presence of the lipopeptide can reduce the surface tension and produce aggregates with the heavy metal forming a precipitate. The use of such lipopeptide could improve the quality of the aforementioned vegetables by removing the excess of heavy metals, which sometimes is above the values permitted by legislation. The use of BS to remove heavy metals is not a novel issue; however, most studies emphasized environmental applications. Literature reports that BS can form complexes with different types of metals and thus, they can help removing heavy metals from industrial effluents and contaminated sites [71]. Anionic BS are capable of forming strong ionic bonds with cationic metals, such as cadmium, favoring their removal from contaminated places [71,72]. Metal-BS complexes can desorb from the soil matrix and further metal can be incorporated into micelles by either electrostatic attraction between negatively charged surfaces and metals or chemical bonding between hydroxide groups and metals [71,73]. These metal-BS interactions can also occur on the charged polar head of BS on micelle surface [71]. Using transmission electron microscopy assays, Das et al. [74], showed that the anionic peptide head groups of a BS produced by a marine bacterium were the binding site for the positively charged metals, lead and cadmium. Qi et al. [75], also showed that the free carboxylic end of the fatty acid in SL formed complexes with cadmium. The removal of metal contaminants using BS is a prospective field to develop innovative cleaning agents for food (Fig. 3C).

3. Nanotechnology, food, and BS Nanotechnology (NT) offers a wide range of opportunities for the development and application of nanosized materials with unique properties that are useful in diverse industrial fields including the food sector. The manufacturing of nanomaterials usually involves physical or chemical methods that utilize high temperatures, are energy consuming or use harsh reducing agents and solvents generating toxic wastes [76]. Surfactants play an essential function in NT as stabilizers, growth control agents, templates, and modifiers [77]. The amphiphilic and self-assembly nature of surfactants, their surface/interfacial tension reduction, particle stabilization ability, and control of physicochemical properties through aggregation, render surfactants as key molecules in nanoscience. Another important feature of surfactants is the possibility for structural variations, allowing the design of molecules for specific target applications [78]. The claims for a more sustainable society, demanding for “green and clean” technologies does not exclude nanomaterials synthesis. Therefore, the exploitation of novel and environmental-friendly

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methods based on biological synthesis emerges as potential candidates to replace the traditional ones. Synthesis of nanoparticles (NP) even using plants and microorganisms as “nanofactories” or by using their metabolites have been described [79]. BS present physicochemical properties similar to chemical surfactants [80] thus, they can be utilized in NT with several advantages. Their status of “natural” molecules showing low eco-toxicity and biodegradability together with their biocompatibility, structural diversity, biological activity, and production from renewable resources are some benefits over the synthetics [2,80]. Moreover, the production of BS satisfies most principles of green chemistry representing important tools for sustainable innovation [3]. The use of overproducing strains, metabolic engineering techniques, more effective downstream processes and heterologous expression methods are also strategies that have been described to turn BS production by traditional fermentation processes more competitive [81,82]. Actually, RL production costs have been significantly reduced and they are becoming suitable to large-scale exploitation, showing that the efforts to turn BS economic are, in fact, effective [83]. The growing demand for green surfactants boosts BS production and global BS market is expected to reach USD 1645.8 million by the end of 2027 [84]. However, it is also important to point out the main gaps or disadvantages involving the use of BS. Their microbial origin, although gives them the status of natural, may also hampered their use in food especially if the producing strain is not considered safe. Within this context, BS from yeasts or GRAS bacteria are more prospective candidates to furnish food-grade BS. Most BS are biosynthesized as mixtures of congeners and the composition of the final product may be difficult to control. A recent trend to solve these problems is the use of bio-inspired surfactants obtained by enzymatic synthesis [85]. This approach allows the development of tailor-made BS glycolipids with single composition, high purity and specific length/type of hydrophobic and hydrophilic moieties. Additionally, such BS are synthesized in a cell-free environment avoiding safety concerns. Owing to the great diversity of BS structures, only some enzymes are available to explore their cell-free synthesis. However, efforts on elucidate microbial biosynthetic pathways and enzymatic engineering techniques can lead to the design of novel bio-inspired compounds. Similar to other industrial sectors, food industry is looking for the new properties and functions provided by the nanoscale to develop innovative products. In this sense, food NT represents an emerging field for the application of BS, not only due to their inherent surface-active properties and green status, but also by their well-known bioactivity. Furthermore, the incorporation of BS in nanostructures can provide novel physicochemical and biological properties suitable for a variety of food-related applications [86]. Besides their intrinsic nutritional value, food products can also have health-promotion benefits; however, the low bioavailability and stability of most health-promoting compounds must be improved to sustain their benefits [87]. A promising alternative to overcome such barrier is the exploitation of nanosized structures. In general, food NT is categorized into three main groups: ingredients for food processing; sensing; and packaging [88]. The encapsulation of active compounds (vitamins, antioxidants, nutraceuticals) in nanocarriers to increase their bioavailability and protection during processing and storage; preparation of nanoemulsions with enhanced functional properties; active nanocoated surfaces to increase safety and preservation; smart sensing packaging to improve quality/safety are some examples of potential applications of NT in food chain [89].

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4. BS in food nanotechnology Although BS have been applied in several industrial segments, ranging from petroleum to cosmetics, their use in NT is still restricted. Most studies emphasize medical/pharmaceutical applications of BS-nanosized materials owing to their biological properties [86,90]. The unique features presented by BS-based nanomaterials are drawing attention of food/agriculture field and some attempts on their exploitation have been described in literature (Table 1). Nanoparticles: In food, metal and organic NP, are essentially applied in packaging materials or in biosensors. Silver NP, gold NP, liposomes, gold nanorods, MnO2 nanosheets and graphene quantum dots, among others, have been designed to sense microbes or chemicals/spoiling molecules [88,89,91]. Most NP are incorporated in analytical devices in combination with biological receptors for rapid detection of important food pathogens like Salmonella sp. and E. coli O157:H7 [89]. Apart from sensing devices, NP can also be included into materials to develop active or smart packing. Active packaging contains specific molecules with ability to absorb or release compounds into or from food aiming to improve food quality [89]. Zinc, silver, zinc oxide, silver-zinc, titanium oxide, and chitosan NP have been incorporated in active packaging to act as antimicrobials, antioxidants, oxygen and ethylene scavengers or moisture absorbents to prevent spoilage, improve shelf life and enhance the safety of meats, fruits, and vegetables [88,91]. By contrast, smart packaging provides information on the quality characteristics and can be developed by using sensing molecules to measure pH, moisture, and gases inside the food product [88,89]. Most sensors are based on organic pigments to detect pH changes and few examples on nanomaterials for such application are available. Nano-sized titanium dioxide (TiO2) was utilized as oxygen detector [92]. Ammonium molybdate was adsorbed in zeolite nanopores to prepare zeolite-molybdate tablets used to scavenge and measure ethylene in avocados packages [93]. Chemical and physical methods are traditionally utilized for synthesis of NP; however, these approaches are expensive and release toxic/hazardous materials into the environment [76]. In addition, the use of toxic chemical reagents makes the obtained NP less suitable for medical or food applications. Chemical reduction and microemulsion-based techniques are the most utilized methods for the synthesis of metal NP and in both cases, surfactants play an essential role. They can act as capping agents decreasing the tendency of the NP to agglomerate, by steric or electrostatic stabilization [94]. Furthermore, the capping agents serve as a diffusion barrier to increase the NP growth and are key elements to stabilize NP, favoring their uniform dispersion in solution [95]. A greener approach for the synthesis of NP is the use of intra or extracellular microbial-mediated synthesis [79], or by using microbial-derived molecules to assist the nanomaterial synthesis [96]. The former has some limitations associated with less control of NP size and shape and it is difficult to separate the NP from culture medium [79]. The use of self-assembly molecules derived from microorganisms or plants such as proteins, fatty acids, or surfactants is advantageous since NP synthesis can occur without interference of cells and media metabolites; in addition, they can also act as templates [97]. Considering that one of the most important issues to green synthesis of NP is the use of nonhazardous materials, replacing chemical surfactants by BS is an interesting option. Some attempts to include BS in NP synthesis even as reagents, carriers or actives are discussed below. Acidic SL diacetate synthesized by Cryptoccocus sp. was utilized to develop bio-functionalized zinc oxide NP able to inhibit the growth of the Salmonella enterica. Enhanced antimicrobial activity promoted by the inclusion of SL surfactant was associated to increasing cell permeability and protein

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leakage [98]. Gold NP functionalized with SL were prepared, and their antimicrobial activity was determined. The presence of BS improved significantly bacterial inhibition and efficacy against Gramnegative bacteria such as Vibrio cholerae [99] showing potential for food and water disinfection. B. subtilis BS increased the stability of biogenic silver NP and enhanced their antimicrobial activity against several phytopathogenic fungi and Gram-positive bacteria. No inhibition effect was observed for most fungal strains with chemical synthesized silver NP (without BS) comparatively to biogenic NP, suggesting BS have an important contribution to antifungal action of the NP [100]. Chitosan-RL NP prepared by ionic gelation method presented spherical shape, average size of 288 nm and positive charge. The addition of RL reduced the size and polydispersity index of chitosan NP and improved antimicrobial activity against S. aureus both planktonic and biofilms. Hybrid chitosan-RL NP showed low cytotoxicity and high antimicrobial activity [101] suggesting good potential to develop active packaging to control S. aureus in food products. Curcumin is a polyphenolic compound that exhibit beneficial properties to human health [102], however its use as a nutraceutical ingredient in functional food and beverages is limited by its low water solubility and stability [103]. A protein/polysaccharide/surfactant NP prepared with zein, propyleneglycol alginate, and RL were designed as delivery system of curcumin. The presence of the BS increased the encapsulation efficiency of curcumin to 92% and improved its photo-stability and bioaccessibility. Authors conclude that such hybrid NP are promising systems to the delivery of hydrophobic nutraceuticals in food and supplements [104]. In a similar work, zein/propyleneglycol alginate/RL complex NP were fabricated for co-delivery of resveratrol and coenzyme Q10. The incorporation of RL enhanced the physicochemical stability and modulated the microstructure of the complex NP. An in vitro gastrointestinal digestion model demonstrated that RL and resveratrol act synergistically to sustain the release of nutraceuticals from the NP. The complex NP showed the potential for designing delivery vehicles of multiple hydrophobic nutraceuticals with different polarities [105]. The bioactive carotenoid lutein was encapsulated in zein NP coated with SL. The resulting spherical NP have around 200 nm, was negatively charged, have an encapsulation efficiency of 90% and increased the water solubility of lutein about 80 times. The lutein carrying NP had an excellent biocompatibility and bioaccessibility, indicating that the core/shell NP can be suitable for encapsulating water-insoluble bioactive compounds in food systems [106]. Nanoemulsions: Nanoemulsions (NE) can be defined as kinetically stable colloidal systems formed by two immiscible liquids stabilized by a surfactant, with droplets size lower than 200 nm [107]. The unique characteristics of such small size emulsions, as high relative surface area and long-term stability make NE ideal for the encapsulation and effective delivery of bioactive lipophilic compounds [107,108]. The delivery in NE formulations of nutraceuticals, coloring and flavoring agents, antimicrobials, and other active ingredients, can enhance the quality, functional properties, nutritional value, and shelf life of foods [108]. Microbial surfactants are good candidates to the preparation of food-grade NE, however, reports in literature are yet limited. One of the earliest studies in this emerging field was reported by Joe et al. [109] using surfactin from B. subtilis. Several cooking oils were initially screened for preparation of surfactin-based NE and sunflower oil, which showed the least droplets particle size (72.5 nm), was selected for further studies. Antimicrobial activity of surfactin-sunflower oil NE was evidenced against food pathogenic bacteria such as L. monocytogenes, B. cereus, Salmonella sp., and S. aureus, as well as,

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against Rhizopus nigricans, A. niger, and Penicillium sp. Indigenous microbial population was also reduced in apple juice, milk, chicken, and vegetables treated with the surfactin-oil NE demonstrating its potential as a natural preservative for food applications [109]. RL were utilized for preparing oil-in-water NE with medium chain triglycerides and food-grade oils from fish, corn, and lemon. The RL-coated droplets were stable in a wide range of temperatures (30°C– 90°C), salt concentrations (60 Lipopeptide

[19]

Phenanthrene

47

57

Glycolipid

[20]

Glycerol

32

42

Rhamnolipid

[21]

Sucrose PAH

32 33

44

Glycoprotein Lipidic surfactant

[22] [23]

Capsular EPS

[24]

Glycolipopeptide

[25]

Lipopeptide

[26]

Motor oil; Peanut oil Kerosene

60

30

>60 EPS

[27]

Diesel oil

32

Lipopeptide

[28]

Glucose

67

>60 Glycoprotein EPS

[29]

Paraffins, crude oil Glucose, PAH

–

[30] [31] [32] [33] [32]

Tetradecane 29 Paraffins, kerosene 30 Tetradecane 63

55

Sulfated heteropolysaccharide Glycolipid Glycolipid Glycolipid EPS

Alkanes, Veg oils

24

100

Rhamnolipid

[34]

Veg oil

27–29

90– 100

Rhamnolipid

[35]

EPS

[36]

Glucose

Empty fields mean that these parameters were not determined by the source.

78

Rhamnolipid

30

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have been derived from terrestrial habitats, but members of this genus are common in marine environments [37,38]. The most studied biosurfactant producer is Pseudomonas aeruginosa. It grows well on various hydrocarbon and nonhydrocarbon substrates and produces rhamnolipids that can form stable emulsions with crude oil and kerosene [39]. Another study observed that P. aeruginosa had a high affinity for crude oil (93% cell adhesion to crude oil) which is also an indication of biosurfactant production [40]. P. aeruginosa DQ8 strain was shown to decrease the ST from 63 to 38 mN/m of culture broth in the presence of various crude oil fractions including polycyclic aromatic hydrocarbons (PAHs) [41]. In addition, nonhydrocarbon substrates such as soybean oil, fish oil, mannitol, and glycerol, can be utilized by Pseudomonas aeruginosa to produce nontoxic biosurfactant which could be useful in oil spill bioremediation as an alternative to chemical dispersants or as substitute of synthetic surfactants in commercial dispersant formulations [42–44]. Strains of P. aeruginosa grown on glycerol produced rhamnolipids (3.8 g/L; CMC 50 mg/L) which reduced the ST to 29 mN/m and emulsified petrol (EI24 70%) and diesel (EI24 80%), further indicating its potential application in oil recovery and bioremediation [44]. A species of Pseudomonas putida, stain BD2, was isolated from Arctic soil that was able to grow on glucose and produce rhamnolipid and sophorolipid simultaneously; the rhamnolipid reduced ST to 31 mN/m and emulsified vegetable oil at 70% efficiency [45].

3.2 Bacillus Members of the genus Bacillus have been predominantly isolated from oil reservoirs or oil contaminated soils and shown to be particularly efficient biosurfactant producers with applications in microbial enhanced oil recovery (MEOR). Bacillus methylotrophicus USTBa, for example, was isolated from a petroleum reservoir and grew well on crude oil in aqueous medium. After 12 days of incubation, B. methylotrophicus removed more than 90% of the crude oil. The ST of the culture medium was measured at 28 mN/m indicating that the organism produced a strong biosurfactant [45]. In addition, the biosurfactant was stable under various pH values and high temperatures (up to 100°C) suggesting its potential application as an oil spill treatment agent in marine environments and in MEOR processes where high salinity and temperatures are common [18]. B. subtilis strain A1 was able to achieve 78% emulsification activity by the production of lipopeptide biosurfactant when grown on crude oil as a sole source of carbon [46]. This strain completely degraded a range of the low-molecular weight alkanes (C10–C14) and up to 97% of the high-molecular weight alkanes (C15–C19) after 7 days of incubation at 40°C. These results suggest that B. subtilis A1 strain could be used in oil spill remediation where light crude oils (high proportion of alkanes) have been spilled. A nonpathogenic Bacillus licheniformis R2 was studied for its potential use in MEOR in laboratory conditions. It produced a low-yield lipopeptide biosurfactant (1 g/L) that lowered the ST to 28 mN/m and the IFT between heavy crude oil and formation water-brine used in core flooding to 0.53 mN/m [47]. When incubated at 85°C, the R2 biosurfactant recovered 37% heavy crude oil recovery over residual oil saturation and retained 88% activity for 90 days [47].

3.3 Acinetobacter Acinetobacter is a genus of gram-negative strictly aerobic Gamma-proteobacteria belonging to the order of Pseudomonadales. Acinetobacter is ubiquitous in nature and commonly found in marine environments. Many species from this genus are known hydrocarbon-degraders that produce extracellular

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EPS [12,48,49]. A. calcoaceticus and A. radioresitensis synthesize emulsan and alasan, respectively, well-known HMW bioemulsifiers [50,51]. A. calcoaceticus was able to synthesize rhamnolipids with CMC of 15 mg/L [52]. A. radioresistens can produce a yield of 4.6 g/L of EPS when grown on ethanol as the sole carbon and energy source [51]. Strains of A. calcoaceticus, and A. oleivorans isolated from the Canadian North Atlantic have been shown to produce bioemulsifiers when grown on petroleum hydrocarbons as the sole carbon source (E24% > 50%) [53]. Given that the strains were isolated and hence adapted to the cold marine environment of the Northern Atlantic, the bioemulsifiers they produce could be effective under low temperature and harsh conditions in offshore oil spills remediation.

3.4 Antarctobacter Antarctobacter is a genus of gram-negative bacteria (order Rhodobacterales), that are strictly aerobic bacteria. Only one species, Antarctobacter heliothermus, has been validly taxonomically described which was isolated from Antarctica [54]. Antarctobacter sp. strain TG22 was isolated from seawater and was found to produce an extracellular water-soluble glycoprotein-type polymer (designated AE22) which formed stable emulsions with different vegetable oils at concentration as low as 0.02% [55]. The strain was grown on marine broth supplemented with 1% glucose and was able to produce an average dry-weight yield of 21 mg/L. The carbohydrate content (total of 15%) of AE22 was dominated by glucosamine, glucuronic acid, fucose and mannose. The protein content represented 5% of the polymer and lipids were not detected, leaving the rest of the polymer content (80%) unidentified [55]. The emulsifying activity of Antarctobacter TG22 polymer was comparable to that of xanthan gum which could be considerably useful in applications for healthcare and food additives.

3.5 Rhodococcus The genus Rhodococcus includes metabolically diverse species that are capable to thrive in different habitats [56]. Members of the genus have been studied mainly for their ability to degrade hydrocarbons and pollutants from different environments [57–59]. Rhodococcus erythropolis, Rhodococcus aurantiacus, and Rhodococcus ruber are among the best known biosurfactant producers of the genus [60,61]. Rhodococcus erythropolis 3C-9 has been shown to grow and produce biosurfactant (CMC of 50 mg/L) only on n-alkanes as the sole carbon source, whereas glucose could not enhance its productivity. The 3C-9 biosurfactant contained fatty acids with lengths from C10 to C22 (docosenoic acid being the most prevalent followed by hexadecenoic acid) and two glycolipids (each dominated by glucose and trehalose monosaccharides). In addition, the 3C-9 biosurfactant significantly enhanced the solubility of PAH substrates [61]. Rhodococcus ruber stain AC 239 produced a small amount of cell-bound glycolipid-type biosurfactant when grown on 1% diesel (v/v). The AC 239 biosurfactant did not reduce the ST as observed for other glycolipid biosurfactants but it emulsified different hydrocarbons with better success (20%–50% greater EI24) when free cells were present in the culture [60]. Rhodococcus fascians extracted from Antarctic soil produced a glycolipid with rhamnose sugars which is not typical for Rhodococcus which usually produces trehalose biosurfactants [62].

3.6 Halomonas Halomonas is a wide-spread genus of the order Alteromonadales. There organisms are found in diverse habitats of both marine [12,53] and terrestrial environments, including hypersaline lakes [63], soils [64–67], and hot springs (45). Members of Halomonas are known to respond to hydrocarbon

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enrichment [31,53,68] and produce EPS [69–71] with versatile properties. A thermophilic H. nitroreducens strain WB1 isolated from a hot spring produced an EPS that was effective at emulsifying different vegetable oils (68%–85%) and aliphatic hydrocarbons (56%–65%) in addition to binding metals. The monosaccharide composition of WB1’s EPS was predominantly composed of glucose, mannose and galactose, and traces of uronic acids [72]. The EPS from Halomonas eurihalina strain H96, isolated from saline soil in Spain, has been characterized to contain high amounts of uronic acids [73] similarly to some marine-derived strains [71]. In addition to emulsifying activity, several species of halophilic Halomonas have been shown to produce highly sulphated exopolysaccharides [31,67] with anticancer activity. For example, halophilic H. stenophila strain B100 exerted a selective proapoptotic effect in T cells from acute lymphoblastic leukemia [74]. EPS form H. halocynthiae KMM 1376 had inhibitory effect on human cancer cell line MDA-MB-231 at concentrations of 50–100 μg/mL [75].

3.7 Alcanivorax Alcanirorax is a gram-negative genus of the Gamma-proteobacteria (order Oceanospirillales) of strictly aerobic marine obligate hydrocarbonoclasic bacteria (OHCB) utilizing predominantly alkanes up to C32 and branched aliphatics [76–78]. The best known species of the genera is Alcanivorax borkumensis which produces a low molecular weight anionic glycolipid biosurfactant when grown on hydrocarbons [79]. This particular glycolipid consists of a glucose sugar linked to a tetrameric chain of fatty acids of C6–C10 length and can be either cell-bound or extracellular [11]. Marine isolate Alcanivorax borkumensis SK2 grown on crude oil produced twice more biosurfactant than in the absence of hydrocarbons. When heavy hydrocarbon fractions were used as the sole carbon source, biosurfactant production was the highest (70 mg/L), only slightly higher than its production when crude oil was used as carbon source (50  20 mg/L). However, the purification of the biosurfactant was easier when the culture was fed with heavy oil fraction as it remained on the surface at all times and consequently there were no substrate impurities [80]. Another species, Alcanivorax dieselolei strain B-5, is the second strain in the genus that has been reported to produce biosurfactant with good surface-active properties (can lower ST to 32 mN/m and emulsify n-hexadecane at 75%). The chemical analysis of the biosurfactant produced revealed that it is a linear lipopeptide with CMC value of 40 mg/L which is comparable to that of rhamnolipids and surfactin [81]. These characteristics make the B-5 lipopeptide an attractive alternative for enhanced oil recovery and bioremediation applications.

3.8 Pseudoalteromonas Pseudoalteromonas is a genus of the order Alteromonadales, that are commonly found in sea ice and cold waters and are well-known producers of glycolipid-type EPS with a wide-range of biological activities and chemical composition [82]. Pseudoalteromonas sp. strain SM20310 isolated from Arctic Sea ice produced EPS (yield of 567 mg/L) with mannose and glucose being the predominant carbohydrates. The ecological role of the EPS was determined to be its improvement of the high-salinity and low-temperature tolerance of the strain [83]. Another study found that marine Pseudoalteromonas (isolated from Antarctica) produced EPS that contained 40% protein with mannose, glucose and galacturonic acid representing the main monosaccharides [84]. The carbohydrate content of EPS from Pseudoalteromonas agarivorans stain Hao 2018 (yield 4.5 g/L) isolated from Yellow Sea of China contained 90% glucose and 6% mannose. The main biological activity of this EPS was moisture retention and absorption of free radicals (i.e. antioxidant) with potential applications in food and

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cosmetics industries [85]. Pseudoalteromonas sp. strain MD12–642 (isolated from Madeira) produced EPS with a particularly high content of uronic acids (up to 68%), so it might find potential applications in the biomedical industry as active ingredients for antithrombotic and antiarthritic drugs [86]. Pseudoalteromonas sp. strain TG12 (isolated from West Scotland) produced EPS that was able to effectively emulsify n-hexadecane (EI24 of 60%) and some vegetable oils. This strain also contained high levels of uronic acids (29%) in addition to xylose (27%), glucosamine (25%) and was effective in desorption of sediment-adsorbed metals (e.g. Al3+, Fe2+/3+, K+, Mg2+, Na+, and Si4+) [87].

3.9 Marinobacter Marinobacter is a genus that is also within the order Alteromonadales. Members of the genus, such as M. hydrocarbonoclasticus and M. algicola are commonly isolated from oil-enriched marine environments [68,88]. Although Marinobacter can use hydrocarbons as a carbon source, various studies demonstrated that it can also grow and produce EPS on other carbon sources such as glucose. Marinobacter species have been shown to produce exopolysaccharide polymers with excellent emulsifying activity against hydrocarbons that were superior to commercial synthetic surfactants like Tween 80 [89]. Marinobacter sp. W1–16 from Antarctic surface seawater produced EPS (molecular weight of 260 kDa) with varying yields, strongly depending on the sugar substrate used to grow the strain and the incubation temperature. The highest yield was obtained when the culture was grown at 15°C and in the presence of 2% glucose. However, the strain was able to synthesize EPS, even at 4°C, albeit in lower quantities, suggesting that the EPS might have cryoprotective functions [89]. In addition, Marinobacter sp. MCTG107b was able to produce a glycolipid-type biosurfactant (grown on glucose) with di-rhamnolipid congeners present that was able to lower the ST to 30 mN/m [90]. Marine sediment isolates of the genus Marinobacter produced powerful emulsifiers when grown on glucose or soybean oil, with activity against hexane and toluene in the range of 45%–64% and 33%–75%, respectively, with some strains producing stable emulsions at 4°C and after high-temperature treatment for up to 18 months [91].

4. Current exploitation of biosurfactants in the oil industry 4.1 Soil bioremediation Hydrocarbon soil contamination, produced by drilling, leaking pipelines, storage tanks, transportation, etc., is a widespread problem with long lasting environmental impacts. Being highly hydrophobic, particularly when adsorbed onto soil particles, hydrocarbons, and heavy metals are very resistant to removal. Typically, a variety of physical and chemical treatments, such as removal, incineration, soil washing, and solvent extraction have been used successfully in the past. However, such techniques are deeply damaging to the soil structure and the autochthonous biodiversity, as well as cost prohibitive. As such, bioremediation is the preferred soil treatment due to its efficiency, lower environmental impact (e.g., low or no toxicity), and cost-effectiveness [92]. Bioremediation involves naturally occurring soil microorganisms which convert petroleum hydrocarbons into carbon dioxide, water, and cell biomass. There are many factors that influence the rate and extent of hydrocarbon degradation in soils, such as moisture content, aeration, pH, temperature, the biological condition of the soil (aged vs fertile soils; nutrient content, and bioavailability), and the concentration, molecular structure and bioavailability of the hydrocarbon

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contaminants [93,94]. The optimization of these environmental factors is critical for the bioremediation success. Soil bioremediation can be conducted either in place (i.e., in situ), or the contaminated soil is upended and, transported to be subsequently treated elsewhere (ex situ). In situ bioremediation involves, generally, the treating of only the top 30-cm layer of the soil with fertilizers to stimulate indigenous soil microorganisms to break down the hydrocarbons [95]. This treatment is the preferred method of choice, but the risk of contaminating underlying aquifers with dissolved hydrocarbons must be considered. Partially purified biosurfactants have been used in situ to increase the solubility and bioavailability of hydrocarbons, and other hydrophobic contaminants, by increasing their surface area [96,97]. A field trial on LaTouche Island in Alaska demonstrated that a biologically derived surfactant, PES-51, could remove 30% of semivolatile petroleum hydrocarbons from a subsurface beach material [98]. A study from Argentina demonstrated that surfactin from B. subtilis strain O9 contributed to significantly more removal of crude oil from sandy loam soil within a period of 300 days [99]. The addition of rhamnolipid from Pseudomonas aeruginosa strain SSC2 to crude oil-contaminated soil sludge resulted in 98% degradation after 4 weeks compared to the nonrhamnolipid control treatment (67%). The effect was enhanced by adding nutrients to the treatments [100]. An in situ experiment of soil bioremediation conducted near an oil production facility in Pakistan demonstrated that higher crude oil degradation (up to 77%) was achieved in soil treated with a combination of a specialized bacterial consortium, rhamnolipids, and nutrients [101]. The efficiency of MELs produced by Candida antarctica SY16 to degrade crude oil in soil was investigated by [102]. The authors compared different bioremediation techniques (i.e., natural attenuation, biostimulation, bioaugmentation, biosurfactant addition, and a combination of all) and concluded that the combined treatment of biostimulation, bioaugmentation with oil degrading Nocardia sp. H17–1 and with MELs caused the highest total petroleum hydrocarbon degradation rate during the first 4 weeks of treatment. However, at the end of the experiment (100 days) the amount of residual hydrocarbons was similar for all treatments [102].

4.2 Microbial enhanced oil recovery (MEOR) MEOR is a process in which microorganism and/or their metabolic by-products are injected into mature oil reservoirs for the recovery of residual crude oil that was not extracted during the initial and secondary extraction processes. The idea behind MEOR is that when favorable conditions are present in the reservoir, the introduced microbes grow exponentially and their metabolic products would mobilize the residual oil [103]. MEOR has its advantages and limitations, and the various processes of its application have been described extensively in the literature and recently summarized by [104]. There are some promising results reported from different research groups that investigate biosurfactants for MEOR applications. Of all known biosurfactants, lipopeptides were mostly used in laboratorybased MEOR studies due to their ability to reduce the IFT to below 0.1 mN/m [105–107]. Both benchscale and in situ lipopeptide production by stains of Bacillus spp. have proven successful in improving the oil recovery, including from wells close to their production limits [108,109]. Surfactins have been shown to maintain activities under a wide range of temperature, pH, and salinity while able to recover sand trapped oil. For example, B. subtilis produced surfactin at high temperature which could emulsify diesel with 90% efficiency and recover over 60% of oil entrapped in sand core [110]. Surfactin was recently shown to alter the wettability of CO2 injected in a subsurface rock formation demonstrating its potential suitability in carbon capture and storage application [111]. Lichenysin was reported to reduce the IFT to

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values of less than 10 2 mN/m (even at low concentrations of 10–60 mg/L) and to have exceptional stability under temperatures as high as 140°C, a pH range from 6 to 10, at salinities up to 10% NaCl, and at calcium (as CaCl2) concentrations up to 340 mg/L. In core flooding experiments, partially purified lichenysin recovered up to 40% of residual oil from sandstone cores compared to 10% recovery when chemical surfactants were applied [112]. Addition of biosurfactants during chemical surfactant flooding can improve the flooding performance in general. In the presence of rhamnolipids, the adsorption to sandstone of the surfactant alkylbenzene sulfonate was reduced by 25%–30% and the quality of oil recovery increased by 7%. It has been suggested that rhamnolipids act as sacrificial agents by preferably adsorbing to the oil sands, making the surfactant more available for displacement activity and resulting in altering the wettability of porous media [113]. Macromolecular biopolymers such as emulsan were shown to remove up to 98% of preadsorbed crude oil to limestone core samples, even at low concentration of 0.5 mg/mL [114]. Recently, another biopolymer produced by Rhizobium viscosum CECT908 showed better efficiency than xanthan gum in the recovery of heavy oil [115].

4.3 Marine oil spill response Crude oil is highly hydrophobic and hence has a very low water solubility, although it is composed of thousands of hydrocarbons and nonhydrocarbon species and metals, each with their respective aqueous solubilities. When an oil is introduced into a water phase, it will float on the surface due to its lower density relative to water. Together with viscosity, ST is an indication of how rapidly and to what extent an oil spreads over the surface and, when dispersed, within the subsurface. The lower the IFT with water, the greater is the extend of spreading [116]. To increase the solubility of oil in water (i.e., to decrease the ST between oil and water), chemicals are applied to an oil slick [3]. Dispersed oil is usually in the form of fine neutrally buoyant droplets with higher surface area-to-volume ratio (diameter size 1–70 μm) compared to nondispersed oil, thus making the oil available for biodegradation by hydrocarbon-degrading bacteria [117]. The natural fate of crude oil biodegradation (biological oxidation by microorganisms) in the marine environment has been extensively described [95,118–124]. Marine bioremediation research has largely been limited to application of fertilizers and/or seed cultures of highly efficient oil-degrading microorganisms, though with conflicting results [125]. The limitation of marine oil remediation when relying solely on indigenous microorganisms is that the concentrations of cells in oil-polluted open water systems is never high enough to effectively emulsify oil [96]. The addition of surfactants (biogenic or synthetically produced) aims to disperse/emulsify the oil and, in turn, speed up the biodegradation process. Biosurfactants have been shown to be effective in dispersing crude oil and enhancing the biodegradation process only under laboratory conditions due to logistical, financial and regulatory limitations of conducting large-scale field trials. In a laboratory study, a bacterial consortium containing oil degrading strains of Ochrobactrum and Brevibacillus with/without rhamnolipids were added to crude oil in 1 L water to stimulate marine oil spill bioremediation. The results showed that the removal efficiency of oil by the bacterial consortium alone was 6% lower compared to the combination of the consortium and biosurfactant [126]. The authors also noted that the presence of rhamnolipid enhanced the biodegradation of alkanes of chain length > nC15, but, interestingly, had the opposite effect on shorter chain alkanes (nC13–C15) by reducing their solubility and increasing their stability. However, the overall removal efficiency of n-alkanes by the bacterial consortium and rhamnolipid was higher than the control. Similar trend was observed for PAH and biomarkers [126]. These results were more or less consistent with another study which also used rhamnolipids in combination with a preadapted bacterial consortium [127], and which were more

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pronounced when nutrients were added to the treatments. The average specific degradation rate was reported to be 23, 20, and 10 times higher than the control for nC15, nC20, and nC25, respectively. Rhamnolipid also stimulated the growth of hydrocarbon degraders within 5 days, which were able to utilize 50% of the crude oil saturated fraction. In addition, LMW PAHs and, notably, the biomarkers pristane and phytane were also significantly degraded in the presence of rhamnolipid [127]. The origin of nutrients (i.e., organic lipophilic or water-soluble) that are added together with rhamnolipids to the treatments can further enhance the degradation of crude oil in seawater and sediment environments [128–130]. Bioemulsifiers can also be used in oil spill response with promising results. A bioemulsifier exopolysaccharide produced by Acinetobacter calcoaceticus, called EPS2003, was shown to be effective in enhancing crude oil biodegradation in natural seawater microcosms [131]. The addition of EPS2003 to the microcosms not only enhanced hydrocarbon-degrading bacteria including Alcanivorax, Marinobacter, Oceanospirillum, and Pseudomonas, but also caused a 2-fold faster biodegradation of the total oil compared to microcosms without the EPS [131]. All these studies, however, focused entirely on the degradation rate of crude oil on a handful of selected oil-degrading bacteria without investigating the indigenous marine microbial community response as a whole. Understanding how natural microbial communities are affected by crude oil with/without the presence of biosurfactants is crucial for advancing the rationale for further research into their suitability for oil spill response. However, due to varying reasons, including current high costs of biosurfactants available on the market and logistics around field studies, there is a marked lack of reported field studies investigating the effectiveness of biosurfactants as oil spill treating agents. A laboratory study compared a synthetic chemical dispersant, called Ultrasperse II, and a surfactin produced by Bacillus sp. strain H2O-1, using a natural marine microbial community [132]. The surfactin enriched for hydrocarbonoclastic bacteria more so than the synthetic dispersant, but no difference in oil biodegradation between the two was observed. Similar results were found in a more recent study in which rhamnolipid, trehalolipid and sophorolipid biosurfactants were compared to three commercial dispersants in oiled microcosms with marine coastal water. Although the biosurfactants caused differential microbial responses, the rate of alkane biodegradation was similar to the microcosms amended with dispersants [133]. In our previous work, rhamnolipid from P. aeruginosa and the synthetic chemical dispersant Finasol OSR52 (stockpiled worldwide) were compared on the response of a natural bacterial community from the Faroe-Shetland Channel, a subarctic region located in the Northeast Atlantic, to crude oil [134]. The rhamnolipid promoted higher diversity in oil-degrading bacteria than the synthetic dispersant, however, the crude oil was ultimately more biodegraded when synthetic dispersant was added to the oil, except for its aromatic fraction. Notably, the synthetic dispersant resulted in a clear inhibition of Cycloclasticus—a genus comprising species of obligate oil-degrading bacteria which are recognized for using aromatic hydrocarbons as a preferred source of carbon and energy [76]. Although biosurfactants may have a positive effect on the oil-degrading microbial community, it is necessary to advance their performance in dispersing crude oil. Developing novel types of biogenic surface-active compounds and more environmentally friendly technologies to combat large offshore oil spills is fertile ground for ongoing and future exploration. In this respect, research and development of biosurfactants for treating oil spills at sea has significantly intensified, particularly over the past 10 years due mainly to concerns over the enormous quantities of the synthetic chemical dispersant Corexit that were used during the Deepwater Horizon oil spill in the Gulf of Mexico. Much of this anew activity ensued through the Gulf of Mexico Research Initiative (GoMRI) and is discussed in the following section.

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5. Recent trends in the development of bio-based dispersants to combat marine oils spills The development of a new generation of dispersants that are as, or more, effective than commercial synthetic dispersants, cost efficient, and have minimal side effects when they come in contact with, or are ingested by, marine organisms and humans is a research area that has gained attention since the Deepwater Horizon disaster. Through GoMRI in the USA as part of the Consortium for the Molecular Engineering of Dispersant Systems (C-MEDS), several projects have been underway aiming to develop bio-based dispersants, either from microorganisms or other natural sources, or using foodgrade ingredients (e.g., silica, polyethylene glycol) that are common additives in food and medicine, and that can be obtained relatively cheaply by the ton (some of the materials from biological sources are described below). Led by scientists at Texas A&M Galveston as part of the Aggregation and Degradation of Dispersants and Oil by Microbial Exopolymers (ADDOMEx) group have shown that EPS produced by microorganisms (microalgae and bacteria) is more efficient at oil dispersal than the synthetic chemical dispersant Corexit [135]. In particular, EPS with a higher protein-to-polysaccharide ratio resulted in higher enzymatic action and marine-oil-snow sedimentation efficiency, higher microbial diversity and cell abundance, and in more extensive biodegradation compared to oil treatments with Corexit, although the latter maintained a more stable emulsion of the oil droplets. Their findings also showed that the microbes in natural samples of seawater were more stressed when exposed to crude oil or to oil together with Corexit, and in response they release more EPS that is higher in protein and carbohydrate/ sugar content, but the EPS aggregates that form do not grow large enough to eventually sediment down. In the absence of Corexit, however, the protein-rich EPS which is formed is significantly more efficient in purging the water column from the oil. Following this, the researchers are exploring ways to trigger the production of protein-rich EPS by natural communities of microorganisms in seawater in the event of another large oil spill. Scientists from the University of Maryland and Tulane University have investigated the use of foodgrade emulsifiers as substitutes for synthetic chemical dispersants. By examining the stability of emulsions of crude oil in seawater in the presence of various food-grade emulsifiers, they found that lecithin (a cell membrane component) from soybean in combination with Tween 80 (emulsifier used in ice cream and other foods) effectively disperse and produce more stable emulsions of crude oil than Corexit [136,137]. In another C-MEDS project, researchers are working on new classes of ‘green’ dispersants made from naturally occurring inorganic and biomolecular materials. Using combinations of natural clay minerals and new carbon materials with synthetic polymer-based materials, new products are being evaluated to test for adhering strongly to the oil-water interface and stabilize oil droplets, which could prevent the formation of large slicks. C-MEDS researchers from the University of South Florida published research showing cactus mucilage dispersed crude oil more efficiently than synthetic dispersants, and notably requiring lower concentrations [138]. In another work, biopolymers derived from cactus mucilage and chitosan show promise in synergistically working together with chemical dispersants, potentially helping to reduce the use of solvents that are typically intrinsic in synthetic chemical dispersant formulations. Taking a different approach, C-MEDS researchers are also exploring new natural gelation agents to prevent oil slicks from spreading and, consequentially, reaching coastlines. Work led by Tulane University and collaborators used a gel-like matrix incorporated with Tween 80 and lecithin which resulted

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in improved stabilization of crude oil in seawater emulsions – more so over longer periods compared to traditional liquid dispersants [139]. The gel-like formulation was designed to largely replace DOSS, the key surfactant component of synthetic chemical dispersants, with lecithin. The application of foodgrade surfactants into a gel-like mesophase acts as a compact buoyant pod for improved delivery of the surfactants to sea surface oil slicks. It does this by way of remaining afloat where the oil is largely confined and avoiding the use of polypropylene glycol and the generation of volatile solvents in the atmosphere through aerial or ship-based spraying. Researchers from the University of Texas at Austin investigated the potential use of nanoparticles as nonconventional dispersants and as tools to improve existing surfactant-based dispersants. Their work led to the development of nanoparticles that are less toxic, and that are more efficient oil spill treatments compared to synthetic chemical dispersants. By mixing hydrophilic nanoparticles (i.e., bare colloidal silica) with a weakly interacting zwitterionic surfactant (caprylamidopropyl betaine or CAPB) to generate a high hydrophilic-lipophilic balance, the nanoparticles and surfactant acted synergistically in forming finer emulsions with enhanced stability, particularly so in a seawater aqueous environment [140]. CAPB is a surfactant that is formed using fatty acids from coconut or palm kernel oil and used in personal care products. Scientists from Brown University and the University of Rhode Island studied the interactions of the obligate hydrocarbon-degrading bacterium Alcanivorax borkumensis with oil across oil-water interfaces that had varying amounts of the following different surfactants: cetylytrimethylammonium bromide (CTAB), lecithin (from soybean), sodium dodecyl sulfate (SDS), dioctyl sulfosuccinate sodium salt (DOSS), and Tween 20, and they compared this to Corexit as the ‘gold’ reference standard. The researchers recorded changes in the growth rate, lag time, and cell density of A. borkumensis at the oil-water interface containing low to high levels of these surfactants and found that not all of them aided this organism’s degradation of oil [141]. The food-grade surfactant, Tween 20, was found to work best by synergistically working with the organism, increased the surface area of oil droplets, and resulting in higher bacterial growth and oil degradation. Conversely, the other surfactants inhibited the adherence of bacterial cells to oil, limiting their biodegradation capacity. In conclusion, the authors recommended further investigation into the use of different surfactants, in particular Tween 20, to replace the current stockpile of synthetic chemical dispersants to treat future oil spills. In a similar study, researchers from the University of Houston compared the food-grade surfactant Tween 20 with several synthetic chemical dispersants to determine how they affect the adhesion of the hydrocarbondegrading species Marinobacter hydrocarbonoclasticus to oil droplets (20–60 μm), which is for some hydrocarbon-degrading bacteria an initial key step for biodegradation [142]. They found that increasing concentrations of all surfactants tested resulted in reduced adhesion of the cells to oil droplets, though electrostatic charge associated with some of the surfactants tested appeared to influence adhesion. Their results suggest that the choice of surfactant(s) in dispersant formulations should be accounted for with respect to how it affects bacterial adhesion to oil droplets and, hence, the biodegradation process. In a study led by Tulane University in collaboration with Lappeenranta University of Technology, Finland, the stability of carboxymethylated chitosan nanoparticles cross-linked with either magnesium, calcium or strontium ions were studied under different pH and salinity in an effort to determine which could be used in oil spill treatment [143]. The nanoparticles cross-linked with calcium ions, as well as when cross-linked with dodecane, were found to be most stable, showing potential for oil-spill treatment. In another study, scientists from different institutions in Canada conducted work on the design of

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a lipopeptide biosurfactant produced by Bacillus subtilis N3d1P from fish waste-based peptone as a primary nutrient substrate for this bacterium [144]. The produced lipopeptide was evaluated as an ingredient together with DOSS, which is the key surfactant ingredient found in Corexit 9500. At a biodispersant ratio of 80/20 (v/v) of lipopeptide/DOSS, a high dispersion efficiency was achieved of 76.8% for Alaskan North Slope crude oil.

6. Conclusion and perspectives Biosurfactants and bioemulsifiers have gained high interest in recent years, due largely to consumer demand for natural ingredients and by companies in search of chemical ingredients conferring improved functional properties and that can be derived from sustainable sources. The marine environment, which is recognized to harbor the largest biosphere and microbial diversity on Earth, offers great potential in the discovery of novel types of these biomolecules for a wide range of industrial applications. With respect to the Oil & Gas industry, the application of biosurfactants for MEOR and in dispersant formulations to treat oil spills are areas of significant interest, but not yet applied on an industrial scale. In the case of MEOR, even though this technology has been around for around 70 years, it has not been widely used by the industry, mainly because of a lack of multidisciplinary research to resolve many of the limitations or knowledge gaps that hinder its advancement. In the case of treating oil spills, chemical dispersants have been used for over 50 years and today they are the preferable treatment for marine oil spills. The toxicity of dispersants to marine life has come a long way since the first studies back in the 1960s [1], but their use does not fail to raise controversy and debate, even today. Numerous studies considering dispersants have reported conflicting results about their toxicological effects, even when the same type of dispersant was used across different studies. This is an area that needs more attention if dispersants are to gain confidence from the scientific community and wider public for their use. In the end, their benefits should outweigh any acute or lasting damage they could pose to marine life and human health. Therefore, to realize the development of bio-based dispersants, such as those produced by microorganisms, there is an urgent need to overcome some major limitations surrounding the microbial production of biosurfactants, principally the high costs involved and the relatively low yields that are often achieved from microorganisms, as addressed in other reviews [145,146]. Even though it may be many years before any are approved and become part of the stockpiled inventory of dispersant agents that stand at the ready for use in the event of an oil spill, the future may hold promise for battling spills with a more environmentally friendly breed of dispersants. The recent emergence of heightened research on this front seems promising. While prevention of oil spills in the first place is paramount, more reliance on bio-based dispersants to treat oil spills at sea should help reduce the potential detrimental environmental impacts that synthetic chemical dispersants can cause. It will be important to select dispersants that speed up the rate and extent that spilled oil is biodegraded by oil-degrading populations of microorganisms. To this end, funding and collaborations between the oil industry and academic researchers need to be expanded. There will be many challenges along the way, but at the very start of this journey there will need to be a requirement for close engagement, respect and a building of trust, between all concerned stakeholders.

References

121

References [1] Portmann JE, Connor PM. The toxicity of several oil-spill removers to some species of fish and shellfish. Mar Biol 1968;1(4):322–9. [2] National Commission on the BP Deepwater Horizon Oil Spill and Offshore Drilling. Deep water: the Gulf oil disaster and future of offshore drilling; 2011. [3] Brakstad OG, Nordtug T, Throne-Holst M. Biodegradation of dispersed Macondo oil in seawater at low temperature and different oil droplet sizes. Mar Pollut Bull 2015;93(1–2):144–52. [4] Rahsepar S, Smit MPJ, Murk AJ, Rijnaarts HHM, Langenhoff AAM. Chemical dispersants: oil biodegradation friend or foe? Mar Pollut Bull 2016;113–9. [5] Hamdan LJ, Fulmer PA. Effects of COREXIT EC9500A on bacteria from a beach oiled by the deepwater horizon spill. Aquat Microb Ecol 2011;63(2):101–9. [6] Kleindienst S, Seidel M, Ziervogel K, Grim S, Loftis K, Harrison S, et al. Chemical dispersants can suppress the activity of natural oil-degrading microorganisms. Proc Natl Acad Sci 2015;112(48):14900–5. [7] McFarlin KM, Prince RC, Perkins R, Leigh MB. Biodegradation of dispersed oil in Arctic seawater at 1°C. PLoS One 2014;9(1):1–8. [8] Brakstad OG, Ribicic D, Winkler A, Netzer R. Biodegradation of dispersed oil in seawater is not inhibited by a commercial oil spill dispersant. Mar Pollut Bull 2018;129(2):555–61. [9] Mulligan CN. Environmental applications for biosurfactants. Environ Pollut 2005;133(2):183–98. [10] Mulligan CN. Recent advances in the environmental applications of biosurfactants. Curr Opin Colloid Interface Sci 2009;14(5):372–8. [11] Abraham W-R, Yakimov MM, Golyshin PN, LUnsdorf H, Lang S, Timmis KN, et al. Alcanivorax borkurnensis gen. Now, sp. nov., a new, hydrocarbon-degrading and surfactant-producing marine bacterium. Int J Syst Bacteriol 1998;48(1998):339–48. [12] Hassanshahian M, Emtiazi G, Cappello S. Isolation and characterization of crude-oil-degrading bacteria from the Persian Gulf and the Caspian Sea. Mar Pollut Bull 2012;64(1):7–12. [13] Ragu
ene`s GHC, Peres A, Ruimy R, Pignet P, Christen R, Loaec M, et al. Alteromonas infernus sp. Nov., a new polysaccharideproducing bacterium isolated from a deep-sea hydrothermal vent. J Appl Microbiol 1997;82(4):422–30. [14] Ragu
ene`s G, Pignet P, Gauthier G, Peres A, Christen R, Rougeaux H, et al. Description of a new polymersecreting bacterium from a deep-sea hydrothermal vent, Alteromonas macleodii subsp. fijiensis, and preliminary characterization of the polymer. Appl Environ Microbiol 1996;62(1):67–73. [15] Rougeaux H, Talaga P, Carlson RW, Guezennec J. Structural studies of an exopolysaccharide produced by Alteromonas macleodii subsp. fijiensis originating from a deep-sea hydrothermal vent. Carbohydr Res 1998;312(1–2):53–9. [16] Passeri A, Lang S, Wagner F, Wray V. Marine biosurfactants II. Production and characterisation of an anionic trehalose tetraester from the marine bacterium Arthrobacter sp. EK 1. Naturforsch 1991;46c:204–9. [17] Cooper DG, Goldenberg BG. Surface-active agents from two Bacillus species. Appl Environ Microbiol 1987;53(2):224–9. [18] Chandankere R, Yao J, Cai M, Masakorala K, Jain AK, Choi MMF. Properties and characterization of biosurfactant in crude oil biodegradation by bacterium Bacillus methylotrophicus USTBa. Fuel 2014;122:140–8. [19] Fooladi T, Moazami N, Abdeshahian P, Kadier A, Ghojavand H, Wan Yusoff WM, et al. Characterization, production and optimization of lipopeptide biosurfactant by new strain Bacillus pumilus 2IR isolated from an Iranian oil field. J Petrol Sci Eng 2016;145:510–9. [20] Reddy MS, Naresh B, Leela T, Prashanthi M, Madhusudhan NC, Dhanasri G, et al. Biodegradation of phenanthrene with biosurfactant production by a new strain of Brevibacillus sp. Bioresour Technol 2010;101 (20):7980–3.

122

Chapter 6 Use of biosurfactants in oil industry and environmental remediation

[21] Dı´az De Rienzo MA, Kamalanathan ID, Martin PJ. Comparative study of the production of rhamnolipid biosurfactants by B. thailandensis E264 and P. aeruginosa ATCC 9027 using foam fractionation. Process Biochem 2016;51(7):820–7. [22] Arora P, Kshirsagar PR, Rana DP, Dhakephalkar PK. Hyperthermophilic Clostridium sp. N-4 produced a glycoprotein biosurfactant that enhanced recovery of residual oil at 96 °C in lab studies. Colloids Surf B Biointerfaces 2019;182(July). [23] Ibacache-Quiroga C, Ojeda J, Espinoza-Vergara G, Olivero P, Cuellar M, Dinamarca MA. The hydrocarbon-degrading marine bacterium Cobetia sp. strain MM1IDA2H-1 produces a biosurfactant that interferes with quorum sensing of fish pathogens by signal hijacking. J Microbial Biotechnol 2013;6 (4):394–405. [24] Casillo A, Sta˚hle J, Parrilli E, Sannino F, Mitchell DE, Pieretti G, et al. Structural characterization of an allaminosugar-containing capsular polysaccharide from Colwellia psychrerythraea 34H. Antonie van Leeuwenhoek. Int J Gen Mol Microbiol 2017;110(11):1377–87. [25] Thavasi R, Jayalakshmi S, Balasubramanian T, Banat IM. Biosurfactant production by Corynebacterium kutscheri from waste motor lubricant oil and peanut oil cake. Lett Appl Microbiol 2007;45(6):686–91. [26] Cooper DG, Zajic JE, Gerson DF. Production of surface-active lipids by Corynebacterium lepus. Appl Environ Microbiol 1979;37(1):4–10. [27] Iyer A, Mody K, Jha B. Emulsifying properties of a marine bacterial exopolysaccharide. Enzyme Microb Technol 2006;38(1–2):220–2. [28] Sriram MI, Gayathiri S, Gnanaselvi U, Jenifer PS, Mohan Raj S, Gurunathan S. Novel lipopeptide biosurfactant produced by hydrocarbon degrading and heavy metal tolerant bacterium Escherichia fergusonii KLU01 as a potential tool for bioremediation. Bioresour Technol 2011;102(19):9291–5. [29] Guti
errez T, Leo VV, Walker GM, Green DH. Emulsifying properties of a glycoprotein extract produced by a marine Flexibacter species strain TG382. Enzyme Microb Technol 2009;45(1):53–7. [30] Hao DH, Lin JQ, Song X, Lin JQ, Su YJ, Qu YB. Isolation, identification, and performance studies of a novel paraffin-degrading bacterium of Gordonia amicalis LH3. Biotechnol Bioprocess Eng 2008;13 (1):61–8. [31] Calvo C, Martı´nez-Checa F, Toledo F, Porcel J, Quesada E. Characteristics of bioemulsifiers synthesised in crude oil media by Halomonas eurihalina and their effectiveness in the isolation of bacteria able to grow in the presence of hydrocarbons. Appl Microbiol Biotechnol 2002;60(3):347–51. [32] Malavenda R, Rizzo C, Michaud L, Gerc¸e B, Bruni V, Syldatk C, et al. Biosurfactant production by Arctic and Antarctic bacteria growing on hydrocarbons. Polar Biol 2015;38(10):1565–74. [33] Vasileva-Tonkova E, Gesheva V. Biosurfactant production by antarctic facultative anaerobe Pantoea sp. during growth on hydrocarbons. Curr Microbiol 2007;54(2):136–41. [34] Xia WJ, Luo ZB, Dong HP, Yu L. Studies of biosurfactant for microbial enhanced oil recovery by using Bacteria isolated from the formation water of a petroleum reservoir. Pet Sci Technol 2013;31(21):2311–7. [35] da Silva RdCFS, de Almeida DG, PPF B, Rufino RD, de Luna JM, Sarubbo LA. Production, formulation and cost estimation of a commercial biosurfactant. Biodegradation 2019;30(4):191–201. [36] Ragu
ene`s G, Christen R, Guezennec J, Pignet P, Barbier G. Vibrio diabolicus sp. nov., a new polysaccharide-secreting organism isolated from a deep-sea hydrothermal vent polychaete annelid, Alvinella pompejana. Int J Syst Bacteriol 1997;47(4):989–95. [37] Baumann P, Bowditch ROND, Baumann L, Beaman B. Taxonomy of marine Pseudomonas species: P. stanieri sp. nov.; P. perfectomarina sp. nov., nom. Rev.; P. nautical; and P. doudoroffii. Int J Syst Bacteriol 1983;33(4):857–65. [38] Bollinger A, Thies S, Katzke N, Jaeger KE. The biotechnological potential of marine bacteria in the novel lineage of Pseudomonas pertucinogena. J Microbial Biotechnol 2020;13(1):19–31. [39] Shahaliyan F, Safahieh A, Abyar H. Evaluation of emulsification index in marine Bacteria Pseudomonas sp. and Bacillus sp. Arab J Sci Eng 2015;40(7):1849–54. [40] Thavasi R, Jayalakshmi S, Banat IM. Effect of biosurfactant and fertilizer on biodegradation of crude oil by marine isolates of Bacillus megaterium, Corynebacterium kutscheri and Pseudomonas aeruginosa. Bioresour Technol 2011;102(2):772–8.

References

123

[41] Zhang Z, Hou Z, Yang C, Ma C, Tao F, Xu P. Degradation of n-alkanes and polycyclic aromatic hydrocarbons in petroleum by a newly isolated Pseudomonas aeruginosa DQ8. Bioresour Technol 2011;102(5):4111–6. [42] Coelho J, Rivonkar CU, Bhavesh NS, Jothi M, Sangodkar UMX. Biosurfactanat production by the quinoline degrading marine bacterium Pseudomonas sp. strain GU 104, and its effect on the metabolism of green mussel Perna viridis L. Indian J Mar Sci 2003;32(3):202–7. [43] Prieto LM, Michelon M, Burkert JFM, Kalil SJ, Burkert CAV. The production of rhamnolipid by a Pseudomonas aeruginosa strain isolated from a southern coastal zone in Brazil. Chemosphere 2008;71(9):1781–5. [44] Das P, Yang XP, Ma LZ. Analysis of biosurfactants from industrially viable Pseudomonas strain isolated from crude oil suggests how rhamnolipids congeners affect emulsification property and antimicrobial activity. Front Microbiol 2014;5(DEC):1–8. [45] Janek T, Łukaszewicz M, Krasowska A. Identification and characterization of biosurfactants produced by the Arctic bacterium Pseudomonas putida BD2. Colloids Surf B Biointerfaces 2013;110:379–86. [46] Parthipan P, Preetham E, Machuca LL, Rahman PKSM, Murugan K, Rajasekar A. Biosurfactant and degradative enzymes mediated crude oil degradation by bacterium Bacillus subtilis A1. Front Microbiol 2017;8 (FEB):1–14. [47] Joshi SJ, Geetha SJ, Desai AJ. Characterization and application of biosurfactant produced by Bacillus licheniformis R2. Appl Biochem Biotechnol 2015;177(2):346–61. [48] Pines O, Gutnick D. Role for emulsan in growth of Acinetobacter calcoaceticus RAG-1 on crude oil. Appl Environ Microbiol 1986;51(3):661–3. [49] Barkay T, Navon-Venezia S, Ron EZ, Rosenberg E. Enhancement of solubilization and biodegradation of polyaromatic hydrocarbons by the bioemulsifier alasan. Appl Environ Microbiol 1999;65(6):2697–702. [50] Kaplan N, Rosenberg E, Jann B, Jann K. Structural studies of the capsular polysaccharide of Acinetobacter calcoaceticus BD4. Eur J Biochem 1985;152(2):453–8. [51] Navon-Venezia S, Zosim Z, Gottlieb A, Legmann R, Carmeli S, Ron EZ, et al. Alasan, a new bioemulsifier from Acinetobacter radioresistens. Appl Environ Microbiol 1995;61(9):3240–4.  ´ A, et al. Structural and physiochemical [52] Hosˇkova´ M, Jezˇdı´k R, Schreiberova´ O, Chudoba J, Sˇ´ır M, Cejkova characterization of rhamnolipids produced by Acinetobacter calcoaceticus, Enterobacter asburiae and Pseudomonas aeruginosa in single strain and mixed cultures. J Biotechnol 2015;193:45–51. [53] Cai Q, Zhang B, Chen B, Zhu Z, Lin W, Cao T. Screening of biosurfactant producers from petroleum hydrocarbon contaminated sources in cold marine environments. Mar Pollut Bull 2014;86(1–2):402–10. [54] Labrenz M, Collins MD, Lawson PA, Tindall BJ, Braker G, Hirsch P. Antarctobacter heliothermus gen. nov., sp. nov., a budding bacterium from hypersaline and heliothermal Ekho Lake. Int J Syst Bacteriol 1998;48(4):1363–72. [55] Guti
errez T, Mulloy B, Bavington C, Black K, Green DH. Partial purification and chemical characterization of a glycoprotein (putative hydrocolloid) emulsifier produced by a marine bacterium Antarctobacter. Appl Microbiol Biotechnol 2007;76(5):1017–26. [56] Finnerty WR. The genus Rhodococcus identification and classification. Annu Rev Microbiol 1992;46:193–218. [57] Whyte LG, Smits THM, Labb
e D, Witholt B, Greer CW, Van Beilen JB. Gene cloning and characterization of multiple alkane hydroxylase systems in Rhodococcus strains Q15 and NRRL B-16531. Appl Environ Microbiol 2002;68(12):5933–42. [58] Kuhn E, Bellicanta GS, Pellizari VH. New alk genes detected in Antarctic marine sediments. Environ Microbiol 2009;11(3):669–73. [59] Wang W, Zhong R, Shan D, Shao Z. Indigenous oil-degrading bacteria in crude oil-contaminated seawater of the Yellow Sea, China. Appl Microbiol Biotechnol 2014;98(16):7253–69. [60] Bicca FC, Fleck LC, Ayub MAZ. Production of biosurfactant by hydrocarbon degrading Phodococcus ruber and Rhodococcus erythropolis. Rev Microbiol 1999;30:231–6. [61] Peng F, Liu Z, Wang L, Shao Z. An oil-degrading bacterium: rhodococcus erythropolis strain 3C-9 and its biosurfactants. J Appl Microbiol 2007;102(6):1603–11.

124

Chapter 6 Use of biosurfactants in oil industry and environmental remediation

[62] Gesheva V, Stackebrandt E, Vasileva-Tonkova E. Biosurfactant production by halotolerant rhodococcus fascians from Casey Station, Wilkes Land, Antarctica. Curr Microbiol 2010;61(2):112–7. [63] Poli A, Moriello VS, Esposito E, Lama L, Gambacorta A, Nicolaus B. Exopolysaccharide production by a new Halomonas strain CRSS isolated from saline lake Cape Russell in Antarctica growing on complex and defined media. Biotechnol Lett 2004;26(21):1635–8. [64] Arias S, del Moral A, Ferrer MR, Tallon R, Quesada E, B
ejar V. Mauran, an exopolysaccharide produced by the halophilic bacterium Halomonas maura, with a novel composition and interesting properties for biotechnology. Extremophiles 2003;7(4):319–26. [65] Mata JA, B
ejar V, Llamas I, Arias S, Bressollier P, Tallon R, et al. Exopolysaccharides produced by the recently described halophilic bacteria Halomonas ventosae and Halomonas anticariensis. Res Microbiol 2006;157(9):827–35. [66] Llamas I, Amjres H, Mata JA, Quesada E, B
ejar V. The potential biotechnological applications of the exopolysaccharide produced by the halophilic bacterium Halomonas almeriensis. Molecules 2012;17 (6):7103–20. [67] Amjres H, B
ejar V, Quesada E, Carranza D, Abrini J, Sinquin C, et al. Characterization of haloglycan, an exopolysaccharide produced by Halomonas stenophila HK30. Int J Biol Macromol 2015;72:117–24. [68] Gutierrez T, Singleton DR, Berry D, Yang T, Aitken MD, Teske A. Hydrocarbon-degrading bacteria enriched by the deepwater horizon oil spill identified by cultivation and DNA-SIP. ISME J 2013;7 (11):2091–104. [69] Guti
errez T, Mulloy B, Black K, Green DH. Glycoprotein emulsifiers from two marine Halomonas species: chemical and physical characterization. J Appl Microbiol 2007 Nov;103(5):1716–27. [70] Gutierrez T, Morris G, Green DH. Yield and physicochemical properties of EPS from Halomonas sp. strain TG39 identifies a role for protein and anionic residues (sulfate and phosphate) in emulsification of n-hexadecane. Biotechnol Bioeng 2009;103(1):207–16. [71] Gutierrez T, Morris G, Ellis D, Mulloy B, Aitken MD. Production and characterisation of a marine Halomonas surface-active exopolymer. Appl Microbiol Biotechnol 2020;104(3):1063–76. [72] Chikkanna A, Ghosh D, Kishore A. Expression and characterization of a potential exopolysaccharide from a newly isolated halophilic thermotolerant bacteria Halomonas nitroreducens strain WB1. PeerJ 2018;2018 (4):1–18. [73] B
ejar V, Llamas IMU, Calvo C, Quesada E. Characterization of exopolysaccharides produced by 19 halophilic strains of the species Halomonas eurihalina. J Biotechnol 1998;61(2):135–41. [74] Ruiz-Ruiz C, Srivastava GK, Carranza D, Mata JA, Llamas I, Santamarı´a M, et al. An exopolysaccharide produced by the novel halophilic bacterium Halomonas stenophila strain B100 selectively induces apoptosis in human T leukemia cells. Appl Microbiol Biotechnol 2011;89(2):345–55. [75] Kokoulin MS, Filshtein AP, Romanenko LA, Chikalovets IV, Chernikov OV. Structure and bioactivity of sulfated α-D-mannan from marine bacterium Halomonas halocynthiae KMM 1376T. Carbohydr Polym 2020;1:229. [76] Head IM, Jones DM, R€oling WFM. Marine microorganisms make a meal of oil. Nat Rev Microbiol 2006;4 (3):173–82. [77] Yakimov MM, Timmis KN, Golyshin PN. Obligate oil-degrading marine bacteria. Curr Opin Biotechnol 2007;18(3):257–66. [78] Olivera NL, Nievas ML, Lozada M, del Prado G, Dionisi HM, Sin˜eriz F. Isolation and characterization of biosurfactant-producing Alcanivorax strains: hydrocarbon accession strategies and alkane hydroxylase gene analysis. Res Microbiol 2009;160(1):19–26. [79] Schneiker S, Martins dos Santos VAP, Bartels D, Bekel T, Brecht M, Buhrmester J, et al. Genome sequence of the ubiquitous hydrocarbon-degrading marine bacterium Alcanivorax borkumensis. Nat Biotechnol 2006;24(8):997–1004. [80] Antoniou E, Fodelianakis S, Korkakaki E, Kalogerakis N. Biosurfactant production from marine hydrocarbon-degrading consortia and pure bacterial strains using crude oil as carbon source. Front Microbiol 2015;6(APR):1–14.

References

125

[81] Qiao N, Shao Z. Isolation and characterization of a novel biosurfactant produced by hydrocarbon-degrading bacterium Alcanivorax dieselolei B-5. J Appl Microbiol 2010;108(4):1207–16. [82] Holmstr€om C, Kjelleberg S. Marine Pseudoalteromonas species are associated with higher organisms and produce biologically active extracellular agents. FEMS Microbiol Ecol 1999;30(4):285–93. [83] Liu SB, Chen XL, He HL, Zhang XY, Bin XB, Yu Y, et al. Structure and ecological roles of a novel exopolysaccharide from the Arctic Sea ice bacterium Pseudoalteromonas sp. strain SM20310. Appl Environ Microbiol 2013;79(1):224–30. [84] Nichols CM, Lardie`re SG, Bowman JP, Nichols PD, Gibson JAE, Gu
ezennec J. Chemical characterization of exopolysaccharides from Antarctic marine bacteria. Microb Ecol 2005;49(4):578–89. [85] Hao L, Liu W, Liu K, Shan K, Wang C, Xi C, et al. Isolation, optimization of fermentation conditions, and characterization of an exopolysaccharide from Pseudoalteromonas agarivorans Hao 2018. Mar Drugs 2019;17(12). [86] Roca C, Lehmann M, Torres CAV, Baptista S, Gaud^encio SP, Freitas F, et al. Exopolysaccharide production by a marine Pseudoalteromonas sp. strain isolated from Madeira Archipelago Ocean sediments. N Biotechnol 2016;33(4):460–6. [87] Gutierrez T, Shimmield T, Haidon C, Black K, Green DH. Emulsifying and metal ion binding activity of a glycoprotein exopolymer produced by Pseudoalteromonas sp. strain TG12. Appl Environ Microbiol 2008;74(15):4867–76. [88] Gauthier MJ, Lafay B, Christen R, Fernandez L, Acquaviva M, Bonin P, et al. Marinobacter hydrocarbonoclasticus gen. nov., sp. nov., a new, extremely halotolerant, hydrocarbon-degrading marine bacterium. Int J Syst Bacteriol 1992;42(4):568–76. [89] Caruso C, Rizzo C, Mangano S, Poli A, Di Donato P, Nicolaus B, et al. Isolation, characterization and optimization of EPSs produced by a cold-adapted Marinobacter isolate from Antarctic seawater. Antarct Sci 2019;31(2):69–79. [90] Tripathi L, Twigg MS, Zompra A, Salek K, Irorere VU, Gutierrez T, et al. Biosynthesis of rhamnolipid by a Marinobacter species expands the paradigm of biosurfactant synthesis to a new genus of the marine microflora. Microb Cell Fact 2019;18(1):1–12. [91] Raddadi N, Giacomucci L, Totaro G, Fava F. Marinobacter sp. from marine sediments produce highly stable surface-active agents for combatting marine oil spills. Microb Cell Fact 2017;16(1):1–13. [92] Mulligan CN. Sustainable remediation of contaminated soil using biosurfactants. Front Bioeng Biotechnol 2021;9:195. [93] Venosa AD, Zhu X. Biodegradation of crude oil contaminating marine shorelines and freshwater wetlands. Spill Sci Technol Bull 2003;8(2):163–78. [94] Huesemann MH. Biodegradation and bioremediation of petroleum pollutants in soil. In: Singh A, Ward OP, editors. Applied bioremediation and phytoremediation soil biology. Springer-Verlag: Berlin Heidelberg; 2004. p. 13–34. [95] Atlas RM, Hazen TC. Oil biodegradation and bioremediation: a tale of the two worst spills in U.S. history. Environ Sci Technol 2011;45(16):6709–15. [96] Ron EZ, Rosenberg E. Biosurfactants and oil bioremediation. Curr Opin Biotechnol 2002;13(3):249–52. [97] Bustamante M, Dura´n N, Diez MC. Biosurfactants are useful tools for the bioremediation of contaminated soil: a review. J Soil Sci Plant Nutr 2012;12(4):667–87. [98] Tumeo M, Braddock J, Venator T, Rog S, Owens D. Effectiveness of a biosurfactant in removing weathered crude oil from subsurface beach material. Spill Sci Technol Bull 1994;1(1):53–9. [99] Cubitto MA, Mora´n AC, Commendatore M, Chiarello MN, Baldini MD, Sin˜eriz F. Effects of Bacillus subtilis O9 biosurfactant on the bioremediation of crude oil-polluted soils. Biodegradation 2004;15(5):281–7. [100] Cameotra SS, Singh P. Bioremediation of oil sludge using crude biosurfactants. Int Biodeter Biodegr 2008;62(3):274–80.

126

Chapter 6 Use of biosurfactants in oil industry and environmental remediation

[101] Tahseen R, Afzal M, Iqbal S, Shabir G, Khan QM, Khalid ZM, et al. Rhamnolipids and nutrients boost remediation of crude oil-contaminated soil by enhancing bacterial colonization and metabolic activities. Int Biodeter Biodegr 2016;115:192–8. [102] Baek KH, Yoon BD, Kim BH, Cho DH, Lee IS, Oh HM, et al. Monitoring of microbial diversity and activity during bioremediation of crude oil-contaminated soil with different treatments. J Microbiol Biotechnol 2007;17(1):67–73. [103] Gao CH, Zekri A. Applications of microbial-enhanced oil recovery Technology in the Past Decade. Energy Sources, Part A Recover Util Environ Eff 2011;33(10):972–89. [104] Nikolova C, Gutierrez T. Use of microorganisms in the recovery of oil from recalcitrant oil reservoirs: current state of knowledge, technological advances and future perspectives. Front Microbiol 2020;10. [105] Youssef N, Simpson DR, Duncan KE, McInerney MJ, Folmsbee M, Fincher T, et al. In situ biosurfactant production by Bacillus strains injected into a limestone petroleum reservoir. Appl Environ Microbiol 2007;73(4):1239–47. [106] Gudin˜a EJ, Pereira JFB, Rodrigues LR, Coutinho JAP, Teixeira JA. Isolation and study of microorganisms from oil samples for application in microbial enhanced oil recovery. Int Biodeter Biodegr 2012;68:56–64. [107] Pereira JFB, Gudin˜a EJ, Costa R, Vitorino R, Teixeira JA, Coutinho JAP, et al. Optimization and characterization of biosurfactant production by Bacillus subtilis isolates towards microbial enhanced oil recovery applications. Fuel 2013;111:259–68. [108] Al-Wahaibi Y, Joshi S, Al-Bahry S, Elshafie A, Al-Bemani A, Shibulal B. Biosurfactant production by Bacillus subtilis B30 and its application in enhancing oil recovery. Colloids Surf B Biointerfaces 2014;114:324–33. [109] Al-Sayegh A, Al-Wahaibi Y, Al-Bahry S, Elshafie A, Al-Bemani A, Joshi S. Microbial enhanced heavy crude oil recovery through biodegradation using bacterial isolates from an Omani oil field. Microb Cell Fact 2015;14(141):1–11. [110] Makkar RS, Cameotra SS. Biosurfactant production by a thermophilic Bacillus subtilis strain. J Ind Microbiol Biotechnol 1997;18(1):37–42. [111] Park T, Joo HW, Kim GY, Kim S, Yoon S, Kwon TH. Biosurfactant as an enhancer of geologic carbon storage: microbial modification of interfacial tension and contact angle in carbon dioxide/water/quartz systems. Front Microbiol 2017;8(JUL):1–12. [112] McInerney MJ, Javaheri M, Nagle DP. Properties of the biosurfactant produced by Bacillus licheniformis strain JF-2. J Ind Microbiol 1990;5(2–3):95–101. [113] Perfumo A, Rancich I, Banat IM. Possibilities and challanges for biosurfactants use in pertroleum industry. In: Sen R, editor. Biosurfactants. Landes Bioscience and Springer Science +Business Media, LLC; 2010. p. 136–45. [114] Gutnik DL, Aviv R, Rosenberg E, Belsky I, Zinaida Z, Sava K. Apoemulsans. USA; Patent 4380504; 1983. p. 1–18. [115] Couto MR, Gudin˜a EJ, Ferreira D, Teixeira JA, Rodrigues LR. The biopolymer produced by Rhizobium viscosum CECT 908 is a promising agent for application in microbial enhanced oil recovery. N Biotechnol 2019;49:144–50. [116] Fingas M. Introduction to oil chemistry and properties. Oil spill science and technology. Elsevier Inc; 2011. p. 51–9. [117] The International Tanker Owners Pollution Federation Limited I. Use of dispersants to treat oil spills. Impact PR Des Ltd; 2011. p. 4–12. [118] Kostka JE, Prakash O, Overholt WA, Green SJ, Freyer G, Canion A, et al. Hydrocarbon-degrading bacteria and the bacterial community response in Gulf of Mexico beach sands impacted by the Deepwater horizon oil spill. Appl Environ Microbiol 2011;77(22):7962–74. [119] Campo P, Venosa AD, Suidan MT. Biodegradability of Corexit 5900 and dispersed South Louisiana crude oil at 5C and 25C. Environ Sci Technol 2013;47:1960–7.

References

127

[120] Prince RC, McFarlin KM, Butler JD, Febbo EJ, Wang FCY, Nedwed TJ. The primary biodegradation of dispersed crude oil in the sea. Chemosphere 2013;90(2):521–6. [121] Wade TL, Sweet ST, Sericano JL, Guinasso NL, Diercks A-R, Highsmith RC, et al. analyses of water samples from the deepwater horizon oil spill: documentation of the subsurface plume. In: Monitoring and modeling the deepwater horizon oil spill: a record breaking enterprise. American Geophysical Union; 2011. p. 77–82. [122] Seidel M, Kleindienst S, Dittmar T, Joye SB, Medeiros PM. Biodegradation of crude oil and dispersants in deep seawater from the Gulf of Mexico: insights from ultra-high resolution mass spectrometry. Deep Res Part II Top Stud Oceanogr 2016;129(MAY):108–18. [123] Joye SB, Kleindienst S, Gilbert J, Handley K, Weisenhorn P, Overholt W, et al. Responses of Microbial Communities to Hydrocarbon Exposures. Oceanography 2016;29(3):136–49. [124] Hazen TC, Prince RC, Mahmoudi N. Marine Oil Biodegradation. Environ Sci Technol 2016;50(5):2121–9. [125] Prince RC. Bioremediation of marine oil spills. In: Timmis KN, editor. Handbook of hydrocarbon and lipid microbiology. Berlin Heidelberg: Springer; 2010. p. 2618–26. [126] Chen Q, Bao M, Fan X, Liang S, Sun P. Rhamnolipids enhance marine oil spill bioremediation in laboratory system. Mar Pollut Bull 2013;71(1–2):269–75. [127] Nikolopoulou M, Eickenbusch P, Pasadakis N, Venieri D, Kalogerakis N. Microcosm evaluation of autochthonous bioaugmentation to combat marine oil spills. N Biotechnol 2013;30(6):734–42. [128] McKew BA, Coulon F, Yakimov MM, Denaro R, Genovese M, Smith CJ, et al. Efficacy of intervention strategies for bioremediation of crude oil in marine systems and effects on indigenous hydrocarbonoclastic bacteria. Environ Microbiol 2007;9(6):1562–71. [129] Nikolopoulou M, Kalogerakis N. Enhanced bioremediation of crude oil utilizing lipophilic fertilizers combined with biosurfactants and molasses. Mar Pollut Bull 2008;56(11):1855–61. [130] Nikolopoulou M, Pasadakis N, Kalogerakis N. Evaluation of autochthonous bioaugmentation and biostimulation during microcosm-simulated oil spills. Mar Pollut Bull 2013;72(1):165–73. [131] Cappello S, Genovese M, Della Torre C, Crisari A, Hassanshahian M, Santisi S, et al. Effect of bioemulsificant exopolysaccharide (EPS2003) on microbial community dynamics during assays of oil spill bioremediation: a microcosm study. Mar Pollut Bull 2012;64(12):2820–8. [132] CRDA C, DDA J, Alvarez VM, van Elsas JD, Seldin L. Response of the bacterial community in oilcontaminated marine water to the addition of chemical and biological dispersants. J Environ Manage 2016;184:473–9. [133] Thomas GE, Brant JL, Campo P, Clark DR, Coulon F, Gregson BH, et al. Effects of dispersants and biosurfactants on crude-oil biodegradation and bacterial community succession. Microorganisms 2021;9 (6):1200. [134] Nikolova CN, Ijaz UZ, Magill C, Kleindienst S, Joye SB, Gutierrez T. Response and oil degradation activities of a Northeast Atlantic bacterial community to biogenic and synthetic surfactants. Microbiome 2021;9 (1):191. [135] Schwehr KA, Xu C, Chiu MH, Zhang S, Sun L, Lin P, et al. Protein: polysaccharide ratio in exopolymeric substances controlling the surface tension of seawater in the presence or absence of surrogate Macondo oil with and without Corexit. Mar Chem 2018;206:84–92. [136] Athas JC, Jun K, McCafferty C, Owoseni O, John VT, Raghavan SR. An effective dispersant for oil spills based on food-grade amphiphiles. Langmuir 2014;30(31):9285–94. [137] Riehm DA, Neilsen JE, Bothun GD, John VT, Raghavan SR, McCormick AV. Efficient dispersion of crude oil by blends of food-grade surfactants: toward greener oil-spill treatments. Mar Pollut Bull 2015;101(1):92–7. [138] Alcantar NA, Fox DI, Thomas S, Toomey RG. Use of cactus mucilage as a dispersant and absorbant for oil in oil-water mixtures. USA: USF Patents 65; 9163374; 2015. p. 65. [139] Owoseni O, Zhang Y, Omarova M, Li X, Lal J, McPherson GL, et al. Microstructural characteristics of surfactant assembly into a gel-like mesophase for application as an oil spill dispersant. J Colloid Interface Sci 2018;524:279–88.

128

Chapter 6 Use of biosurfactants in oil industry and environmental remediation

[140] Worthen AJ, Foster LM, Dong J, Bollinger JA, Peterman AH, Pastora LE, et al. Synergistic formation and stabilization of oil-in-water emulsions by a weakly interacting mixture of zwitterionic surfactant and silica nanoparticles. Langmuir 2014;30(4):984–94. [141] Bookstaver M, Bose A, Tripathi A. Interaction of Alcanivorax borkumensis with a surfactant decorated oilwater interface. Langmuir 2015;31(21):5875–81. [142] Dewangan NK, Conrad JC. Adhesion of Marinobacter hydrocarbonoclasticus to surfactant-decorated Dodecane droplets. Langmuir 2018;34(46):14012–21. [143] Kalliola S, Repo E, Sillanp€a€a M, Singh Arora J, He J, John VT. The stability of green nanoparticles in increased pH and salinity for applications in oil spill-treatment. Colloids Surf A Physicochem Eng Asp 2016;493:99–107. [144] Zhu Z, Zhang B, Cai Q, Ling J, Lee K, Chen B. Fish waste based Lipopeptide production and the potential application as a bio-dispersant for oil spill control. Front Bioeng Biotechnol 2020;8(July):1–16. [145] Banat IM, Satpute SK, Cameotra SS, Patil R, Nyayanit NV. Cost effective technologies and renewable substrates for biosurfactants’ production. Front Microbiol 2014;5(DEC):1–18. [146] Tripathi L, Irorere VU, Marchant R, Banat IM. Marine derived biosurfactants: a vast potential future resource. Biotechnol Lett 2018;40(11 12):1441–57.

CHAPTER

Biosurfactants produced from corn steep liquor and other nonconventional sources: Their application in different industries

7

X. Vecinoa, A.B. Moldesa, A. Martı´nez-Arcosa, B. Cid-P
ereza,b, A. Lo´pez-Prietoa, and J.M. Cruza a

Chemical Engineering Department, CINTECX, University of Vigo, Vigo, Spain, bAnalytical and Food Chemistry Department, Faculty of Chemistry, University of Vigo, Vigo, Spain

1. Introduction Biosurfactants possess an extensive and expansive market, including cosmetic, pharmaceutical, agrochemical, cleaning, and environmental industries with a market size of USD 5.52 billion by 2022, at a compound annual growth rate (CAGR) of 5.6% [1]. However, the market ceiling size could be even higher, taking into account the market size of surfactants (USD 52.4 billion by 2025 from USD 42.1 billion in 2020, at a CAGR of 4.5%) [2]. Biosurfactants are surface-active compounds produced by different microorganisms, including, bacteria, yeasts and fungi [3]. These can be linked to the cell membrane of microorganisms, known in this case as cell-bound biosurfactants, or excreted to the culture medium [4]. Biosurfactants possess several advantages over chemical synthetic homologues, such as low toxicity and high biodegradability [5,6] and being moreover stable at extreme pH and salinity [7]. The composition and yield of biosurfactants during their biotechnological production are linked to the culture medium conditions like pH, nutrient composition, agitation, oxygen availability, and temperature [8]. It is important to remark that the same microorganisms can produce different biosurfactant extracts depending on the culture conditions and even depending on the fermentation stage: lag, stationary or exponential phase [9]. Several works have been published about the capacity of Bacillus subtilis to produce a group of biosurfactants that includes surfactin, fengycin, or iturin. These biosurfactants are lipopeptides with different molecular weights between 1044 and 1486 Da, and diverse fatty acid and amino acid compositions [10]. Numerous publications exist in the literature where it has been reported that biosurfactants are produced as a mixture of congeners [11]. At this point, an important question is whether biosurfactants are more biocompatible and more stable than chemical surfactants, and so, what is wrong with biosurfactants? Why does the commercialization of biosurfactants not finish clearing in comparison with chemically synthesized surfactants? The reason could be found in the total capital cost investment as well as in the production Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00007-7 Copyright # 2023 Elsevier Inc. All rights reserved.

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cost of biosurfactants in comparison with their chemically synthesized homologues. Some authors have suggested that the use of secondary raw materials as carbon or nitrogen sources for obtaining more industrial cost-competitive biosurfactants in a sustainable way [12]. Therefore, Patria et al. [13] have proposed the use of food waste for obtaining rhamnolipids in controlled fermentations using Acinetobacter calcoaceticus, which was isolated from petroleum-contaminated soil due to the improved of the apparent solubility of polycyclic aromatic hydrocarbons (PAHs) [14]. Authors stimated to obtain 548,226–662,256 kg/year of 50% pure rhamnolipids, depending on the number and size of bioreactors used during the fermentative processes, although all the proposed sceneries used the same total working volume of bioreactors (350 m3). In this simulation tool, after a fermentation process, the culture medium was subjected to centrifugation and ultrafiltration steps. Afterwards, rhamnolipids from the liquid phase in this set-up were extracted under differential extraction with hexane, followed by a distillation process. Finally, rhamnolipids were freeze-dried. The total equipment cost (TEC) for this process is an average of USD 9,366,566.77 from the three scenarios. According to Patria et al. [13], the annual rhamnolipid production cost was around USD 24,840,987.94–21,888,147.01, with the minimum selling price of 50% purity rhamnolipids being about USD 36.03–37.51 per kg. That is a competitive selling price as the market selling price of 50% purity rhamnolipids syrup is USD 225.00 per kg [13]. Additionally, Table 1 shows the economic analysis carried by Patria et al. [13], taking into account the mean values from the three proposed scenarios. The sceneries proposed by Patria et al. [13] are quite favorable from an economical point of view for the biotechnological production of biosurfactants, but these economic figures could be improved if biosurfactants were produced as a natural by-product in a fermented stream avoiding the controlled fermentation process, as is the case of the biosurfactants produced from corn steep liquor (CSL). There exist several patents and works where it has been demonstrated that the corn wet-milling industry generates a residual stream spontaneously fermented by lactic acid bacteria as well by Bacillus sporulated strains [15,16] that produce biosurfactants, which can be extracted from CSL by liquid-liquid extraction using chloroform or ethyl acetate [17,18]. The processes to obtain biosurfactants from CSL [15] are similar to those proposed by Patria et al. [13] but avoid the investment and cost related to the biotechnological production of biosurfactants. Both biosurfactants are recovered from the fermented media using organic solvents. Thus, the investment and production cost could be reduced by at least threefold in the case of using CSL as a source of biosurfactants in comparison with the sceneries proposed by Patria et al. [13] for obtaining rhamnolipids from food waste streams under controlled fermentations. Table 2 includes the characteristics of the biosurfactant extracts obtained from CSL. Depending on the extractive process, biosurfactant extracts with slightly different characteristics can be obtained [17,19]. It has been demonstrated that the biosurfactant extract obtained from CSL with chloroform (named BS1) can reduce the surface tension of water about 40 mN/m and possesses a critical micellar concentration (CMC) of about 146 mg/L, whereas the biosurfactant extract obtained from CSL using ethyl acetate (named BS2) reduces the surface tension of water up to 38 mN/m with a CMC of 176 mg/L. Table 2 also includes the fatty acid composition and amino acids that are included in these biosurfactant extracts, which are composed of lipopeptides, free fatty acids, phospholipids, and antioxidants [17,19]. Authors have demonstrated that BS1 possesses a higher content of phospholipids in comparison with BS2, although BS1 possesses a lower EC50. Among the antioxidant compounds, BS1

1 Introduction

131

Table 1 Economic evaluation for rhamnolipid biosurfactant production. Category

Costs (€)a

Raw material cost Utility Operating labor Direct supervisory and clerical labor Maintenance and repairs Operating supplies Laboratory charges Patent and royalties Depreciation Local taxes and insurance Plant overhead costs Administrative costs Research and Development Costs Distribution and marketing costs Contingency Annual rhamnolipid production cost (RLPC) Annual production capacity of 50% purity rhamnolipids (kg/year) Selling price of 50% purity rhamnolipids syrup (per kg) Annual revenue Gross profit Net profit Minimum selling price of 50% purity rhamnolipids (per kg) Cumulative net present value (NPV) Internal rate of return (%)

1,973,231.07 1,358,728.28 604,200.87 90,630.13 1,848,949.99 277,342.50 90,630.13 584,394.47 3,022,523.87 924,474.99 1,526,268.59 381,567.15 3,485,305.09 2,727,174.21 584,394.47 19,479,815.8 612,381.01 189.71 116,176,836.31 96,697,020.51 81,077,880.19 31.89 529,383,178.13 58.53

a Dollar to euro conversion carried out on September 6, 2021. Adapted from Patria RD, Wong JWC, Johnravindar D, Uisan K, Kumar R, Kaur G. Food waste digestatebased biorefinery approach for Rhamnolipids production: a techno-economic analysis. Sustain Chem 2021;2:237–253. https://doi.org/10.3390/suschem2020014.

is composed of vanillic acid, p-coumaric acid, ferulic acid, sinapic acid, and quercetin; whereas BS2 is composed of protocatechuic acid, vanillic acid, caffeic acid, epicatechin, p-coumaric acid, ferulic acid, sinapic acid, and quercetin. The EC50 values of BS1 and BS2 were of 8.51 and 9.65 mg/mL, respectively [19]. CSL is the first residual stream proposed in the literature as a direct source of biosurfactants and in the following sections we present some possible uses and applications that have been demonstrated until now in the environmental, nanotechnology, cosmetic, pharmaceutical, and personal care fields as well as for agrochemical and food applications. Additionally, it is important to highlight that the microorganism producer of this biosurfactant extract from CSL is a Bacillus strain that was recently isolated and identified as Aneurinibacillus aneurinilyticus [4,20] as shown in Fig. 1.

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Chapter 7 Their application in different industries

Table 2 Chemical characteristics of the biosurfactants obtained from CSL after liquid-liquid extraction with organic solvents. Properties

BS1

BS2

Reference

Surface tension reduction of water (mN/m) CMC (mg/L) Composition C (%) N (%) H (%) S (%) Amino acids (%) Fatty acids (%) Antioxidants

40.1

37.7

[16]

145.77

176.02

[16] [16]

75.20 1.32 11.72 < 0.30 8.25 49.1 Vanillic acid (14.7%), p-coumaric acid (4.0%), sinapic acid (37.8%), quercetin (32.8%), and ferulic acid (10.7%)

68.83 1.08 11.08 < 0.30 6.75 NA* Protocatechuic acid (32.8%), vanillic acid (17.9%), p-coumaric acid (13.4%), sinapic acid (13.5%), quercetin (12.9%), and ferulic acid (9.6%) NA*

Phospholipids (%)

21

[18]

[16]

NA* Not analyzed.

FIG. 1 Microscopical images of the Aneurinibacillus aneurinilyticus strain isolated from corn steep liquor obtained with a stereoscopic zoom microscope.

2 Use of naturally produced biosurfactants from CSL in different industries

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2. Use of naturally produced biosurfactants from CSL in different industries 2.1 Environmental applications Biosurfactants are widely used in the environmental field, especially in bioremediation, oil spill cleanup operations, soil remediation, and flushing, among others [5]. The properties of a biosurfactant, like its toxicity, are important for use as a nonhazardous substance in this field. In this regard, it has been demonstrated that biosurfactants extracted from CSL are less toxic and less irritating [21] than chemically synthesized surfactants with a half-life (t1/2) lower than 30 days [22]. The t1/2 indicates the time required to reduce the concentration of biosurfactant by 50%, using a concentration of biosurfactant of 1 g/L. In the literature, few works exist related to the biodegradability of biosurfactants. Rodrı´guez-Lo´pez et al. [22] have studied the spontaneous biodegradation of biosurfactant extract from CSL under different pH (5–7), temperature (5–45 °C) and biodegradation time (15–55 days) scenarios, without adding any inoculum at the beginning of the assay, corroborating the biodegradable character of the biosurfactant extract obtained from CSL. Moreover, Rodrı´guez-Lo´pez et al. [23] studied the cytotoxic effect of biosurfactants obtained from CSL using fibroblast cell lines and observed that biosurfactant extracts obtained from CSL were not cytotoxic at the highest concentration assayed (1 g/L). Regarding the irritant character of the biosurfactant extract from CSL, Rodrı´guez-Lo´pez et al. [21] published that the biosurfactant extract obtained with chloroform (named BS1) was not an irritant at the highest concentration proposed in the Hen’s egg-chorioallantoic membrane (HET-CAM) test (1 g/L), whereas sodium dodecyl sulfate (SDS) at the same concentration produced hemorrhage in small vessels of the Hen’s egg choriaollantoic membrane. In the HET-CAM test, chemicals are placed in direct contact with the chorioallantoic membrane of the Hen’s egg, observing if vascular injury or coagulation occurs in response to the assayed substance. Table 3 includes the hazardous characteristics that are evaluated in a substance and define the hazardous characteristics of the biosurfactant extracts obtained from CSL regarding their composition and the information reordered in several publications [16,17,19]. Based on their nonhazardous characteristics, biosurfactants extracted from CSL could be proposed as potential cleaning agents or as solubilizing agents in the bioremediation of contaminated sites. Therefore, Vecino et al. [24] demonstrated the potential use of a biosurfactant from CSL in the elimination of PAHs from soil, observing that only 1.7% of naphthalene remained in the soil after decontamination, whereas anthracene and pyrene were reduced up to 51.7% and 69.4%, respectively. On the other hand, Bustos-Va´zquez et al. [25] proposed a biosurfactant from CSL for the removal of burnet oil from sand. After performing discontinuous extraction experiments with several formulated cleaner solutions with the biosurfactant, the authors concluded that a 1/20 solid/liquid ratio (w:v) and 6 h of contact were enough to achieve nearly 67% of oil extraction. Instead, other works have included a biosurfactant extract obtained from CSL in a formulation of lignocellulosic biocomposites to treat wastewater. The obtained results showed an increase of around 10% and 62% in the elimination of dye compounds and removal of sulfates, respectively [26].

2.2 Nanotechnology applications In general, nanotechnology term is used to describe the design, production, and application of structures at the nanoscale level. This level can help for the availability and distribution of the principles active ingredients, for example, through the skin suggesting to be a good technology for cosmetic and

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Table 3 Hazardous characteristics of the biosurfactants obtained from CSL (BS1 and BS2) after liquid-liquid extraction with organic solvents. Hazardous properties

Definition

Biosurfactants from CSL

HP1

Explosive

HP2

Oxidizing

HP3

Flammable

HP4

Irritant

HP5

Specific target organ toxicity/Aspiration toxicity

HP6

Acute toxicity

HP7

Carcinogenic

HP8

Corrosive

HP9

Infectious

HP10

Toxic for reproduction

HP11

Mutagenic

HP12

Release of an acute toxic gas

HP13

Sensitizing

HP14

Ecotoxic

HP15

Waste capable of exhibiting a hazardous property listed above is not directly displayed by the original waste

( ) No explosive substances were detected ( ) No oxidizing substances were detected ( ) No flammable substances were detected ( ) A HET CAM test was carried out using 1 g/L of BS ( ) No toxic substances were detected ( ) No toxic substances were detected ( ) No carcinogenic substances were detected ( ) No carcinogenic substances were detected ( ) No microorganisms were detected in BS ( ) No toxic for reproduction substances were detected ( ) No mutagenic substances were detected ( ) Biosurfactant contains substances with a high boiling point ( ) No sensitizing substances were detected ( ) No ecotoxic substances were detected ( )

dermatological applications [27] The application of biosurfactants in the field of nanotechnology has had spectacular growth in recent years, with India (23%) the country with the most publications related to the subject, followed by Canada (9%), Brazil (8%), the United States (7%), China (5%), and the United Kingdom (5%) (see Fig. 2). Additionally, Fig. 2B shows the percentage of publications by year from 2006 to 2021. In the year 2010 and from 2016 up to nowadays, there was an increase in the number of publications on the application of biosurfactants in nanotechnology. This could be because nanotechnology has potential applications in almost all disciplines of science and technology, from electronics, sensing and catalysis to cosmetics and drug delivery [28–31], but

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FIG. 2 Percentage of publications about the use of biosurfactants in the field of nanotechnology (A) per country and (B) per year.

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almost all of the methods used to synthesize nanoparticles (NPs) nowadays usually involve the use of hazardous reactants with a high environmental and human health impact. This is the reason why recent studies have pointed out the use of biosurfactants as an environmentally friendly alternative for the synthesis of NPs [27,32,33]. Additionally, biosurfactants impart bioactivity to the NPs, making biosurfactant-mediated synthesis a more attractive option [27,34]. In NP synthesis, biosurfactants are employed as capping agents providing uniform NP dispersion in the liquid medium. Moreover, they can be used as stabilizing and dispersing agents, since biosurfactants are adsorbed onto the metal NPs to stabilize and prevent their agglomeration [27,35,36]. Among these, glycolipid biosurfactants like rhamnolipids are the most used in nanotechnology, followed by lipopeptide biosurfactants. Rhamnolipids allow the synthesis of NPs with small size and good stability on their own or combined with other compounds. Moreover, they can act as coating and capping agents and functionalize the surface of NPs providing them more biocompatibility. Regarding lipopeptide biosurfactants, they are mainly used in nanomedicine as nanocarriers of different drugs since they act as coating agents of NPs of lower sizes than other types. However, the application of glycolipopeptides, glycopeptides and glycoproteins in the synthesis of NPs is not an active field of application yet [32]. There are many different nanomaterials, with gold and silver NPs (Au-NPs, Ag-NPs) very attractive due to their unique optical properties. Some of the biosurfactants proposed in the literature for NPs synthesis are produced by pathogenic microorganisms like Pseudomonas aeruginosa (P. aeruginosa), which reduces their applicability. This is the reason why other authors have started to report the use of biosurfactants from nonpathogenic strains, for example produced by lactic acid bacteria, a microorganism listed as “generally recognized as safe” (GRAS) [37], in Ag and Au NPs synthesis. In this case, the NPs could be considered as biocompatible material since biosurfactants are obtained from a GRAS microorganism [34].

2.3 Cosmetic, pharmaceutical, and personal care applications The potential applications of biosurfactants in different fields have been demonstrated by many studies, including cosmetic, pharmaceutical, and personal care applications [38–42]. However, limited research exists regarding the uses of these compounds in products of the pharmaceutical and cosmetic industries. Nowadays, chemical surfactants are included, mainly in cosmetics and personal care products, as stabilizing agents [43]. However, these chemical compounds are increasingly being replaced by biosurfactants due to the goal of developing more natural and biocompatible formulations [44]. An example of this is the use of natural surfactants, like mannosylerythritol lipids (MELs), in the formulation of hair care products [45]. Another example of a biosurfactant in hair care products is the lipopeptide biosurfactant extract from CSL. In this regard, Rinco´n-Fonta´n et al. [46,47] demonstrated the positive effect of this biosurfactant on damaged and dyed hair. Authors corroborated that a chemical surfactant (e.g., hexadecyltrimethylammonium bromide (CTAB), which is a cationic synthetic surfactant used in cosmetic formulation) causes more hair damage than the biosurfactant extract from CSL, which maintains hair with better smoothness and flexibility and obtains more defined cuticles. Additionally, biosurfactants have been used to improve the solubilization of antidandruff ingredients in shampoos. Despite there being many agents that exhibit an antimicrobial action against the growth of the organisms responsible for dandruff, Zn pyrithione is preferred in terms of overall performance. In this sense, Rinco´n-Fonta´n et al. [48] demonstrated the potential of using a combination of biosurfactant extract

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from CSL and Tween 80 as good additives in antidandruff formulations because they can improve the solubility of Zn pyrithione in aqueous formulations up to 147 times in comparison with its solubility in water. Furthermore, the authors proved the inclusion of the biosurfactant extract from CSL with another antiseptic agent, tea tree oil, which could improve the final oil-in-water emulsion developed. This was the first work where a biosurfactant is considered to be a stabilizing agent in antidandruff formulations [48]. On the other hand, biosurfactants are taking a growing interest in the formulations of dermal products, where synthetic surfactants are known for producing adverse effects [49]. Therefore, the use of biosurfactants is a promising strategy for the formulation of more environmentally friendly and sustainable dermal products. For instance, Knoth et al. [50], in a preliminary study to evaluate the suitability of biosurfactant extract from CSL as an excipient in dermal formulations, determined different physicochemical properties, such as wettability, foaming capacity, hydrophilic-lipophilic balance (HLB)-value, stabilizing properties, and antioxidant capacity. The authors showed that the biosurfactant extract from CSL could be a suitable co-stabilizer for nanoemulsions and nanocrystals for dermal application due to its good surface-active properties. In addition, as the biosurfactant extract possesses antioxidant capacity, authors found skin protective properties and that the biosurfactant increased the dermal penetration efficacy for lipophilic actives. Overall, the biosurfactant extract from CSL could be used as an excipient in different dermal formulations, showing that this compound could be used as a co-stabilizer with antioxidative and penetration enhancing properties at the same time. In the field of dermal products, inorganic compounds are usually less irritating than organic compounds. Therefore, recent works have focused on mica, a silicate mineral obtained from the mining industry, to develop sun creams that are safer for the skin, as demonstrated by Rinco´n-Fonta´n et al. [51]. The sunscreen protection factor (SPF) is a method to quantify the level of photoprotection provided by materials. Measuring the amount of UV light absorbed by a material Rinco´n-Fonta´n et al. [51], determined the SPF of different formulations based on different mica minerals mixed or not with a biosurfactant extract from CSL, concluding that the biosurfactant could increase the SPF values of the micas by more than 2000%. This is because the addition of biosurfactant extract produced a synergistic effect on the protective properties of mica since biosurfactant can protect against the UV radiation by itself and enhances the formation of a Pickering emulsion (named the emulsion composed of an oil and inorganic material). Moreover, another dermal application of the biosurfactant extract from CSL was carried out by Rinco´n-Fonta´n et al. [52]. In this case, the authors studied the synergistic effect between mica and biosurfactant to stabilize Pickering emulsions containing vitamin E using a triangular design. vitamin E is a fat-soluble antioxidant that can act as a peroxyl radical scavenger, and, therefore, it is a sun protector with poor solubility in water. The results showed that the presence of biosurfactant extract improved the emulsion volume up to 70% after 22 days, for an emulsion composed of vitamin E and biosurfactant, while the presence of a biosurfactant extract with 10% of mica improved the vitamin E emulsion volume up to 80% after 30 days of the assay. For emulsions formulations, Rinco´n-Fonta´n et al. [53] proposed the biosurfactant extract from CSL as a stabilizing agent of vitamin C in aqueous cosmetic formulations, and, therefore, to improve the stability of this vitamin included in cosmetic and pharmaceutical formulations. In this work, a factorial design was established to obtain a theoretical equation to predict the degradation of vitamin C in the presence and absence of a biosurfactant extract. The results showed that the biosurfactant application could reduce the degradation of vitamin C to 58% after 14 days in comparison with the same formulations in absence of

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biosurfactant extract. This finding opens the door to the use of this lipopeptide biosurfactant as a stabilizing agent of formulations containing unstable active principles like vitamin C. On the other hand, Rodrı´guez-Lo´pez et al. [21] evaluated the preservative and irritant capacity of the biosurfactant extract from CSL, since there are important challenges for the pharmaceutical and cosmetic industries. The authors observed that the biosurfactant extract obtained from CSL had important preservative capacity against Staphylococcus aureus (S. aureus), Escherichia coli (E. coli), and P. aeruginosa, killing all these germs in 7–14 days. In addition, it had an inhibitory effect on Aspergillus brasiliensis (A. brasiliensis), reducing the growth of this microorganism up to 95% after 14 days. In regard to irritant capacity, the biosurfactant extract did not show any irritant effect on HET-CAM test, contrary to SDS, used as control, which produced intravascular coagulation. Additionally, Rodrı´guez-Lo´pez et al. [54] studied the use of a biosurfactant extract from CSL as a potential ingredient in antiacne formulations, specifically Acne vulgaris. The cosmetic formulations evaluated were composed by nonnano zinc oxide (ZnO) (0%–2%), biosurfactant extract from CSL (0%–5%) and salicylic acid (0%–2%), studying the inhibitory effect on Cutibacterium acnes (also named Propionibacterium acnes). An inhibitory effect on C. acnes of the biosurfactant extract was shown in formulations when this was combined with an intermediate concentration of ZnO (1%), and it was significantly higher than ZnO alone and comparable to ZnO (1%) with salicylic acid (1%). Recently, Rodrı´guez-Lo´pez et al. [55] evaluated the effect of the presence of biosurfactant extract from CSL on the permeation of pharmaceutical compounds (such as benzocaine, benzoic acid, benzotriazole, caffeine, lidocaine, salicylic acid, ibuprofen, indomethacin, procaine, and tetracaine) through a silicone membrane simulating skin membrane. The results showed that compounds with a relative molecular mass below 200 had an increase in permeation in the presence of the biosurfactant extract, whereas those above 200 had a decrease in the permeation capacity. Some of the applications of the biosurfactant extract from CSL in the cosmetic, pharmaceutical and personal care industries are summarized in Table 4.

2.4 Agrochemical applications In recent years, environmental regulations, and the growing interest in replacing persistent substances with other more biodegradable ingredients have promoted a reduction in the use of pesticides and chemical surfactants in numerous areas of agriculture, including crop protection and agro-chemical formulations. Biosurfactants are nontoxic compounds that play an important role in sustainable agriculture due to their biological nature and easy degradability [56]. Biosurfactants have a promising future to remove plant pathogens due to their antimicrobial activity [57,58] and they can even replace chemical surfactants in pesticide formulations, mitigating their harmful effects and permanence on crop surfaces, water, and groundwater [59] as well as agricultural soils [60]. In this way, Lo´pez-Prieto et al. [61] studied the antifungal activity of a biosurfactant extract obtained from CSL against bacterial pathogens like P. aeruginosa and E. coli. The inhibition percentages of these two pathogens were measured after 24 and 48 h of incubation at 37°C and the biosurfactant extract obtained from CSL inhibited 100% of both P. aeruginosa and E. coli at concentrations of 0.9 and 1.0 g/L, respectively. Moreover, the filtration of the biosurfactant extract provided a significant increase of its antimicrobial capacity, particularly in the case of P. aeruginosa, which can be attributed to the removal of impurities from the biosurfactant extract that could protect the pathogen microorganism. Additionally, the same biosurfactant extract from CSL was evaluated against A. brasiliensis and Candida albicans (C. albicans), two

Table 4 Examples of cosmetic and pharmaceutical formulations of biosurfactant extract from CSL. Formulation type Pharmaceutical and cosmetic formulations

Biosurfactant role Solubilization and preservation of active ingredients in pharmaceutical and cosmetic products

Biosurfactant concentration 1 g/L

Studied variables

• Antimicrobial and

Results

• Biosurfactant extract

preservative capacity of biosurfactant extracts • Irritant effect on biological membranes

•

• Hair application

Replace lost lipids in hair

0.2 g/L

• • • •

Temperature pH Treatment time Adsorption capacity of hair

• • •

obtained from corn milling industry has important preservative capacity against S. aureus, E. coli, and P. aeruginosa, killing these germs in 7 to 14 days and an inhibitory effect on A. brasiliensis, reducing the growth up to 95% after 14 days. Extract from L. pentosus improved the growth of pathogenic microorganisms, except in the case of S. aureus and A. brasiliensis, after 28 and 14 days, respectively. Neither biosurfactant produced any irritation on biological membranes Higher biosurfactant adsorption at low temperatures pH effect only at middle or high temperatures Maximum capacity of adsorption (3679 μg/g) at the higher concentration of biosurfactant

Reference [20]

[45]

Continued

Table 4 Examples of cosmetic and pharmaceutical formulations of biosurfactant extract from CSL—cont’d Formulation type Hair application

Biosurfactant role Replace lost lipids in hair

Biosurfactant concentration 0.06–0.53 g/L

Studied variables

• Temperature • Treatment time • Biosurfactant concentration

• Adsorption capacity of hair

Hair application

Stabilization in antidandruff formulations based on Zn pyrithione powder

0%–5% (w/w)

• Emulsion characterization and emulsion stability • Zn pyrithione solubility • Factorial design based on tea tree oil/water ratio and both surfactant and biosurfactant concentration

Results

• Maximum capacity of

adsorption (10,549 μg/g) achieved at 0.295 g/L of biosurfactant (above the critical micellar concentration of the biosurfactant) • The biosurfactant maintains hair structure in a good state and obtains more defined cuticles • Biosurfactant extract produces formulations with small particle size, higher stability, and good solubility of Zn pyrithione (The formulation that provided the most favorable results contains Tween 80 (5%) and BS extract (2.5%), with an O/W ratio of 0.01. This provides the smallest particle size (40.5 μm), good stability after 30 days (91%), and the highest solubility of Zn pyrithione (59%))

Reference [46]

[47]

Dermal application

Co-stabilization of dermal formulations with antioxidative and penetration enhancing properties at the same time

0.05%–0.1%

• • • • • •

• • • •

Wettability Foaming capacity HLB value Emulsifying properties Antioxidant capacity Size characterization of nanocarriers and determination of zeta potential Biophysical properties Effect of BS on skin properties Cleansing capacity Penetration enhancing properties

• Improvement of •

•

• •

• Sunscreen formulation

Increase the sunscreen protection factor (SPF) of micas

0.5 g/L

Pharmaceutical and cosmetic formulations

Pickering emulsion stabilizer containing vitamin E

25%–50%

• Sunscreen protection

•

factor (SPF)

• Emulsifying capacity • Emulsion

•

characterization

• Synergistic effect between silicate mineral (mica) and the biosurfactant extract using a triangular design

•

•

wettability (but not of oil) on different surfaces Foaming, cleansing capacity, and efficacy to form stable o/w emulsions lower than classical surfactants Emulsification efficacy is pH-dependent and increases with pH values around 5 Suitability of the biosurfactant to be used as co-emulsifier Biosurfactant possesses antioxidative and penetration enhancing properties Biosurfactant possesses good skin tolerability Increase in the sun protection factor (SPF) value due to the addition of the biosurfactant extract Biosurfactant extract presence improves the emulsion volume up to 70% after 22 days for an emulsion composed of vitamin E and biosurfactant Mica increases the emulsion stability until values of 80% after 30 days for those emulsions with 10% of mica Both novel ingredients produce a synergistic effect on the Pickering emulsions

[49]

[50]

[51]

Continued

Table 4 Examples of cosmetic and pharmaceutical formulations of biosurfactant extract from CSL—cont’d Formulation type

Biosurfactant role

Biosurfactant concentration

Cosmetic and health care formulations

Stabilizing agent of formulations

0–2 g/L

Dermal application

Alteration in the rate and extent of permeation of some drugs

0.005 and 0.5 g/L

Dermal application

Acne vulgaris treatment

0%–5%

Studied variables

• Vitamin C degradation (Box-Behnken factorial design based on concentration of vitamin C, concentration of biosurfactant, and storage time) • Biosurfactant concentration • Permeation of ten pharmaceutical compounds though polydimethylsiloxane membrane (PDMS)

• Factorial design based on concentration of nonnano zinc oxide (ZnO), concentration of biosurfactant from corn milling industry (BS-CSW) and concentration of salicylic acid

Results

• Biosurfactant presence

Reference [52]

can inhibit the degradation of vitamin C in water up to 58% after 14 days

• Increase in permeation with biosurfactant in compounds with a relative molecular mass below 200 • Decrease in permeation with biosurfactant present in compounds with a relative molecular mass above 200 • Synergetic effect on the inhibition of Cutibacterium acnes between ZnO and BS-CSW in absence of salicylic acid (The biosurfactant extract alone possesses by itself antimicrobial activity against C. acnes, therefore 1% of ZnO alone gave inhibitory hales of 10.2 mm close to the inhibitory hales (9.4 mm) achieved with 2.5% of BS-CSW. However, the inhibition hales can achieve a double effect with the combination of 1% ZnO and 5% of biosurfactant extract (22.2 mm))

[54]

[53]

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143

different fungi that may affect a wide variety of food products and crops. The results obtained showed a fungicidal effect (100% inhibition) against A. brasiliensis at concentrations of 0.33 g/L at an incubation temperature of 4°C. In contrast, C. albicans was more resistant, achieving growth inhibitions of only 76.3% at a temperature of incubation of 40°C and by using a biosurfactant concentration of 0.99 g/L [62]. Nowadays, it is important to highlight the potential of biosurfactants in sustainable agriculture for seed protection, plant pathogen control, and fertility [63].

2.5 Food applications In the food industry, biosurfactants may be used as wetting agents to improve the water retention capacity of food components, as bioemulsifiers, due to their facility to form stable emulsions and help ingredient mixing [64,65], and as green additives and preservatives, due to their bactericidal activity, to extend the shelf-life of foods by avoiding their microbial contamination [61,62]. In this sense, the use of a biosurfactant extract from CSL to assess the morphological changes in grape surfaces (area, roughness, perimeter, etc.) when they were coated with the biosurfactant solution (1 g/L) should be highlighted relative to noncoated grapes as shown by Martı´nez-Arcos et al. [66]. The results showed that the grapes coated with the biosurfactant solution decreased their surface roughness and considerably reduced shape changes over time in comparison with noncoated ones [66]. Additionally and for the first time, the same biosurfactant extract from CSL was tested in the maceration process of two red wine-grape varieties (Vitis vinifera L. cv. Cabernet Sauvignon and Aglianico) to evaluate its influence on the color properties of red wine [67]. The results showed that the presence of the biosurfactant extract from CSL enhanced the color properties of red wine, probably due to co-pigmentation interactions. In addition, Lo´pez-Prieto et al. [18] also proved that the use of the biosurfactant extract from CSL enhanced the functional properties of probiotic food products. Therefore, there was a positive effect on the growth of Lactobacillus casei (L. casei), using an incubation temperature in the range of 30–40°C and without modification of its homofermentative pathway when the biosurfactant extract was added to a drinkable yogurt.

3. Use of other nonconventional sources to produce biosurfactants Apart of CSL, as a nonconventional source for the production of biosurfactants, there are other sources [68–70], considered an excellent cost-efficient substrates, to produce them as follows.

3.1 Dairy industrial wastes Alkan et al. [71] have evaluated a whey residue as fermentation medium to produce biosurfactants by lactic acid bacteria (Streptococcus thermophilus, Lactobacillus acidophilus, and Lactobacillus rhamnosus), and then compared with a commercial culture medium (MRS broth). Considering the beneficial properties of biosurfactants produced by lactic acid bacteria on food industry due to their compatible emulsifying and antiadhesive activities, in this work were determined several physicochemical (surface tension, emulsification index, biomass, and oil spreading test) and antiadhesive properties of biosurfactants produced from the two compared culture medium (whey waste and MRS broth). Although the

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values of surface tension obtained for biosurfactants from MRS broth (43.44–45.35 mN/m) were lower than those obtained with biosurfactants from whey medium (48.85–53.51 mN/m) no significant differences were obtained between them. The same behavior was observed for the results of biomass (ranged from 9.20–11.80 g/L and 6.38–8.20 g/L from whey medium and MRS broth, respectively) and for oil spreading (with zone diameter ranged from 1.87 to 5.92 cm), where data differences were statistically insignificant for biosurfactants from both whey culture and MRS broth. The emulsification index was obtained for lactic acid bacteria after 1, 24, and 168 h using different culture conditions and the emulsification stability was reached throughout 168 h of contact time, with values ranged from 19.5% to 32.5%. In general, the values of emulsification index reported by both whey and MRS broth were statistically insignificant. In contrast, when the lactic acid specie is considered, no differences were found between the emulsification index provided by Streptococcus thermophilus and Lactobacillus acidophilus (values ranged from 26.75% to 32.50% for whey medium after 168 h), however the values obtained for Lactobacillus rhamnosus (20.75% for whey medium and after 168 h) were significantly lower than to the other. The antiadhesive properties of biosurfactants were tested against different pathogens (E. coli, S. aureus, and P. aeruginosa) at biosurfactant concentrations of 2.5, 5 and 10 mg/mL, observing that the last dose presents the most antiadhesive effect. So, when using a biosurfactant concentration of 10 mg/L, it was preserved the adhesion of S. aureus, P. aeruginosa, and E. coli between 37.25%–52.5%, 10.25%–23.25%, and 5.32%–11.50%, respectively, being E. coli the microorganism more resistant to the biosurfactants than the other tested pathogens. In addition, biosurfactants from S. thermophilus shown the most inhibition percentages against all bacteria tested, being the biosurfactants produced by L. rhamnosus the least effective. Daverey et al. [72] have produced sophorolipids (SLs), a glycolipid biosurfactant produced by Starmerella bombicola (previously called Candida bombicola) using dairy industry wastewater. The maximum production of SLs (62 g/L) was obtained when the wastewater was supplemented with glucose (10% w/v), yeast extract (0.2% w/v), and soybean oil (10% v/v); the yeast growth in this medium was able to reduce the chemical oxygen demand (COD) with an elevated removal efficiency of 86%. The properties of the produced SLs in terms of CMC and minimum surface tension were 30 mg/L and 33.6 mN/m, respectively. On the other hand, the effect of temperature, pH, and ionic strength on surface tension reduction of the SLs was also evaluated, and a great stability of the biosurfactant in a wide range of pH (2 10), with a maximum activity at pH 6 was found. In relation to temperature influence, the biosurfactant under study showed a great stability to boiling during a time of 2 h and it was also stable in presence of sodium chloride concentrations between 0% and 20% (w/v), thus confirming the utility of SLs produced from dairy industry wastewater, for environmental applications.

3.2 Fruit and vegetable wastes Rocha et al. [73] have evaluated the use of cashew apple juice (CAJ), an agroindustrial residue rich in carbohydrate, fibers, vitamins, and mineral salts, as fermentation medium for the production of biosurfactants by Acinetobacter calcoaceticus using submerged fermentation in a rotatory shaker with an incubation temperature of 30°C, during a fermentation time of 5 days. It was investigated the biosurfactant production by A. calcoaceticus in a natural cashew apple juice medium in terms of microbial growth, surface tension of the broth, substrate uptake (total reducing sugars) and emulsifying activity and these results were compared with those reported by a defined mineral medium. The data reported

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indicate that the microorganism was able to growth and produce biosurfactant in both evaluated fermentation medium (mineral and natural CAJ), reducing the surface tension in both cases. It was also able to emulsify kerosene by reaching an emulsion index of 58.8% in natural CAJ, after 34 h of incubation at 30°C and pH 7.0, and an emulsifying activity of 85.7% in a synthetic mineral medium after 90 h incubation at the same temperature and pH values. These results indicate that natural CAJ culture medium could replace other conventional or synthetic substrates for biosurfactant production. Silva et al. [74] evaluated the potential of fruit wastes as substrates for the biosynthesis of biosurfactants. In this study, six waste fruit samples including oranges, mangoes, and mixed fruits were self-fermented and submitted to taxonomic analysis by the MEGAN software based on DIAMOND alignments to establish the identity of the microorganisms that contain the genes involved in the biosurfactant production. The MEGAN analysis indicates that the six fruit samples studied were dominated by the Gamma-proteobacteria, with most reads assigned to the genera Klebsiella ( 63%), Enterobacter ( 19%), Stenotrophomonas ( 7%), Escherichia ( 7%), and Acinetobacter ( 2%). In addition, a functional analysis was carried out by using the BiosurfDB classification system, integrated into the MEGAN analysis, to assess the relative abundance of genes associated with the biosynthesis of biosurfactants. In this sense, the functional profiles of the microbiota found in all fruit samples tested were very similar and they have a great potential for the biosynthesis of lipopeptides. In this respect, a commonly found type of biosurfactant in the fruit samples studied was the lipopeptide iturin A, belonging to the iturin family of biosurfactants, in whose biosynthesis were mainly involved genes of the Brevibacillus, Klebsiella, and Enterobacter genera. These results proved that the studied fruit wastes can be considered as a promising source of microbiota that can produce biosurfactants, mainly lipopeptides, potentially useful in various industrial sectors such as agriculture, food, chemical, and pharmaceutical. Chooklin et al. [75] focused in the evaluation of the banana peel as a substrate for biosurfactant production by screening the best microorganisms to maximize its production. Among 55 bacterial isolates only 30 presented biosurfactant activity, being Halobacteriaceae archaeon AS65 strain the most active in terms of reducing the surface tension of the culture medium from 68 to 30 mN/m. The highest biosurfactant production (5.30 g/L) was achieved after 54 h of incubation at 30°C in a mineral salt medium containing 35% (w/v) of banana peel and 1 g/L of commercial monosodium glutamate. The biosurfactant obtained showed a surface tension of 25.5 mN/m and a CMC of 10 mg/L. Thermal and pH stability of the biosurfactant was evaluated in terms of surface tension and emulsification activity, observing a great stability at temperatures over up 100°C and at pH values in the range of 5–10 as well as a high level of salt tolerance, when concentrations of sodium chloride up to 12% almost no affect its emulsification activity. The lipopeptide nature of the biosurfactant produced by Halobacteriaceae archaeon AS65 was confirmed by Fourier Transform Infrared Spectroscopy (FTIR), Nuclear Magnetic Resonance (NMR) and Electrospray Ionization-Mass Spectrometry (ESI-MS). Among the potential uses of the crude biosurfactant it should be mentioned its high antimicrobial activity against several pathogens such us B. cereus, C. albicans, P. aeruginosa, and S. aureus. It also had ability to enhance the solubility of PAHs in aqueous phase and it was able to remove high percentages of metals (between 56.87%–75.51% and 74.51%–92.87% for Cd and Pb, respectively) when using biosurfactant concentrations about 10 times higher than its CMC. Furthermore, the maximum biodegradation of crude oil (70.5%) was achieved in the presence of the biosurfactant under study, which suggests a potent accelerator for hydrocarbon biodegradation by increasing the oily substances bioavailability.

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3.3 Starch-rich wastes Gurjar et al. [76] used a rice mill polishing residue (RMPR) as substrate to produce surfactin using Bacillus subtilis MTCC 2423 by submerged fermentation in a rotatory shaker with an incubation temperature of 30°C, during a maximum fermentation time of 72 h. The biosurfactant was concentrated by multistage foam fractionation previously to its recovery by acid precipitation with HCl 6 N and, finally, the precipitate of biosurfactant obtained was dried in an oven at 5°C until weighted. The recovery of surfactin obtained was 4.17 g/Kg of residue employed, representing the 69% of the total biosurfactant produced and recovered from the foam formed. The residual supernatant contains about 30% of the surfactin yield and shows values of 23 and 69 mg/L for 5-day biological oxygen demand (DBO) and COD, respectively, permitting the direct use of these residual solutions for soil remediation purposes. On the other hand, the surfactin produced was characterized by ESI-MS and the spectra confirmed a dominant molecular weight of 1076 Da. In addition, the surfactin produced from RMPR was applied for removal of copper ions by foam separation, achieving a maximum removal percentage of 72.5% for copper solutions of 25 mg/L at pH 10. For other pH values (between 8 and 9.5) the removal capacity was decreased by 10%–20%. For higher copper concentrations of 50 or 100 mg/L the removal of copper was also reduced by 5%–8%, which can be attributed to a binding saturation of the biosurfactant tested. Andrade et al. [77] focused on the production and purification of the biosurfactant mannosylerythritol lipids-B (MEL-B) instead of the other three homologues (A, C, and D) by Pseudozyma tsukubaensis using cassava wastewater (an agroindustrial waste) as substrate. It is important to highlight that the main objective of this work was to develop a low cost and effective purification strategies based on ultrafiltration processes as well as to introduce cheap substrates, like cassava wastewater, as culture medium to reduce the effective cost of the biosurfactant MEL-B production. The use of cassava wastewater provided a production of MEL-B of 1.26 g/L of culture medium and the ultrafiltration process at small scale (20 mL) with membranes (100 kDa PES) allowed an 80% retention of MEL micelles, whereas more than 95% of the proteins were found in the permeate. Very similar results were found when this ultrafiltration system was scaled up to 500 mL by using a cross flow filtrations unit. On the other hand, different analysis of the purified biosurfactant by Gas Chromatography couple to Mass Spectrometry (GC–MS), FTIR, Matrix-Assisted Laser Desorption/Ionization with Time of Flight analyzer couple to Mass Spectrometry (MALDI-TOF-MS) and NMR were combined to confirm the chemical structure of the purified biosurfactant MEL-B. In this sense, the NMR analysis confirmed the majority production of the MEL-B homologue (around 90%) and the production of a second minority stereoisomer (around 9%). In addition, the GC–MS and the MALDI-TOF-MS analysis also revealed the identification of MEL-B with the presence of the main fatty acids within its structure [C8:0 and C12:0 (657 m/z) and C8:0 and C14:1 (683 m/z)] and indicated the production of minority stereoisomers, about 8%–10%. So, the results of this work suggest a promising alternative for the conventional production of MEL that is produced using a synthetic culture medium, it is extracted with organic solvents and purified using column chromatography.

3.4 Lignocellulosic wastes Konishi et al. [78] demonstrated the efficient production of SLs by S. bombicola using a corncob hydrolysate medium derived from lignocellulosic feedstocks. The production under different conditions was studied in a jar fermenter to optimize the saccharification of corncobs and the fermentation

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conditions. The results showed that the optimal concentration of sulfuric acid was 1% and the SLs production and cell growth was inhibited by autoclaving, something that can be solved by adding ammonium nitrate to the autoclaved medium. Finally, an SL production of 49.2 g/L with a volumetric productivity of 12.3 g/L/day was achieved. Panjiar et al. [79] achieved successfully the biosurfactant production from rich rice straw hydrolysates by a soil bacterium, Serratia nematodiphila. The production was optimized in different ways: ratio of C:N, adding palm oil as inducer, increasing the substrate concentrations, and adding some growth-inhibitory compounds into the media. Furthermore, the glycolipids obtained were extracted, characterized, and analyzed to explore some properties. Finally, the results showed that the production of glycolipids was higher using xylose-rich media (instead of using glucose-rich media), the inhibitors did not affect the growth nor the production, and the biosurfactant obtained possessed good emulsifying and antimicrobial activities.

3.5 Oily and glycerol-based wastes Chen et al. [80] have demonstrated the efficient production of the biosurfactant rhamnolipid by P. aeruginosa using kitchen waste oil as carbon source. The results with this medium were compared with those using other four carbon sources and the pH and nitrogen concentration of cultivation were optimized. Moreover, some properties of the biosurfactants (structure, stability, and oil removal effect) were analyzed. Finally, results showed the best fermentation performance with kitchen waste oil as carbon source, a pH of 8 and a nitrogen concentration of 2 g/L. On the other hand, the produced rhamnolipid presented excellent physicochemical properties in terms of interfacial activity and environmental stability and a good ability to be used in oil recovery. Sharma et al. [81] carried out rhamnolipid production with P. aeruginosa P7815 using waste cooking oil as the sole carbon source in a batch and fed-batch bioreactors. The product obtained was characterized, the emulsion stability was investigated at different environmental conditions (pH, temperature, and salinity) and the antibacterial activity was also studied. Results showed the highest production (16 g/L) using fed-batch bioreactor, being the rhamnolipid obtained stable in a wide range of conditions and capable of reducing the surface tension to 29 mN/m. Moreover, this biosurfactant exhibited antibacterial activity against some gram-positive and gram-negative bacteria. Luna et al. [82] evaluated a medium of distilled water with 9% ground-nut oil refinery residue plus 9% corn steep liquor for the production of biosurfactant by Candida sphaerica UCP 0995. The biosurfactant production was carried out in shake flasks and in a bioreactor and some physicochemical properties were determined. The results showed higher biosurfactant yield when using bioreactor (21 g/L) instead of flasks, a capacity to reduce water surface tension to 27 mN/m and a good capacity to destabilize oil-water emulsions. In addition, the product was proved to be nontoxic to indigenous marine microbiota, which allows the application in remediation of marine environments contaminated with hydrophobic pollutants. Radzuan et al. [83] used the P. aeruginosa type strain PAO1 for biosurfactant production in a 2 L bioreactor using a 2% palm oil refinery by-product as carbon source in minimal medium. Cell growth, rhamnolipid production, identification, and characterization of crude rhamnolipid extract were studied. The results showed a maximum production of 2.1 g/L with a reduction in surface tension to 28 mN/m and a CMC of 53 mg/L.

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4. Concluding remarks Biosurfactants are interesting and promising molecules used in different sectors such as environmental, nanotechnology, food, agrochemicals, cosmetics, pharmaceuticals, and personal care. Among biosurfactants, this book chapter is focused on the natural production of a biosurfactant extract from CSL and their application in different industries. For example, this extract contains a lipopeptide which is extracted from a residual stream of corn wet-milling industry. This lipopeptide is remarkable because it is produced as a natural product in a liquid stream during the wet steeping process of corn. Therefore, CSL is an interesting alternative to obtain cost-competitive and value-added biosurfactants, since its biotechnological production has zero cost. At the same time, it is increasing the market opportunities for these secondary raw materials (e.g., from nonconventional sources as CSL) under the circular economy framework by using the biosurfactant obtained in the same corn industry or in other industries. The only cost is related to the downstream processing, and the biosurfactant can be obtained by simple liquid-liquid extraction with organic solvents. On the other hand, the biosurfactant extract obtained from CSL possesses surfactant and antioxidant activities; thus, it can be considered a multifunctional extract. Additionally, the biosurfactant extract obtained from CSL presents good properties like being environmentally friendly, biodegradable, biocompatible, and less toxic than chemical surfactants, which opens potential uses in different industrial sectors as described in this chapter. Finally, the trends and challenges, in the case of biosurfactant extract obtained from CSL, could be (i) to perform further analytical tests to incorporate the biosurfactant extract safely and under industry-dependent regulations, (ii) study this biosurfactant extract in more cosmetic, food and agrochemical applications, (iii) evaluate other recovery processes to obtain a more biocompatible biosurfactant extract by applying membrane technology or liquid-solid procedures, and/ or (iv) bring it to the market.

Acknowledgment This research was supported by the Spanish Ministry of Science, Innovation and Universities under the project RTI2018-093610-B-100, by the Spanish Ministry of Science and Innovation under the project PID2019103873RJ-I00 and by the Xunta de Galicia under the project GPC-ED431B 2020/17.

References [1] Markets and markets aviation connectors market worth 5.52 Billion USD by 2022 Available online: https:// www.marketsandmarkets.com/PressReleases/aviation-connectors.asp. [2] Markets and markets surfactants market worth $52.4 Billion by 2025 - Exclusive report by marketsandmarkets™. [3] Desai JD, Banat IM. Microbial production of surfactants and their commercial potential. Microbiol Mol Biol Rev 1997;61:47–64. [4] Lo´pez-Prieto A, Martı´nez-Padro´n H, Rodrı´guez-Lo´pez L, Moldes AB, Cruz JM. Isolation and characterization of a microorganism that produces biosurfactants in corn steep water. CyTA J Food 2019;17:509– 16. https://doi.org/10.1080/19476337.2019.1607909.

References

149

[5] Santos DKF, Rufino RD, Luna JM, Santos VA, Sarubbo LA. Biosurfactants: multifunctional biomolecules of the 21st century. Int J Mol Sci 2016;17:401. https://doi.org/10.3390/ijms17030401. [6] Cruz JM. Microbial glycoprotein and lipopeptide biosurfactants production, properties and applications. In: Moldes AB, Vecino X, Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Banat IM, Thavasi R, editors. Microbial biosurfactants and their environmental and industrial applications. Taylor & Francis Group; 2018. p. 106–28. [7] Vecino X, Devesa-Rey R, Cruz JM, Moldes AB. Study of the synergistic effects of salinity, PH, and temperature on the surface-active Properties of Biosurfactants produced by lactobacillus Pentosus. J Agric Food Chem 2012;60:1258–65. [8] Moldes A, Vecino X, Rodrıı´guez-Lo´pez L, Rinco´n-Fonta´n M, Cruz JM. Biosurfactants: the use of biomolecules in cosmetics and detergents. In: Rodrigues AG, editor. New and future developments in microbial biotechnology and bioengineering. Elsevier; 2020. p. 163–85, ISBN:9780444643018. [9] Velraeds MMC, Van Der Mei HC, Reid G, Busscher HJ. Inhibition of initial adhesion of uropathogenic enterococcus faecalis by biosurfactants from lactobacillus isolates. Appl Environ Microbiol 1996;62: 1958–63. https://doi.org/10.1128/aem.62.6.1958-1963.1996. [10] Kim PI, Ryu J, Kim YH, Chi YT. Production of biosurfactant Lipopeptides Iturin A, Fengycin, and Surfactin A from bacillus subtilis CMB32 for control of colletotrichum gloeosporioides. J Microbiol Biotechnol 2010;20:138–45. https://doi.org/10.4014/jmb.0905.05007. [11] Perez KJ, Dos Santos Viana J, Lopes FC, Pereira JQ, dos Santos DM, Oliveira JS, Velho RV, Crispim SM, Nicoli JR, Brandelli A, et al. Bacillus Spp. isolated from Puba as a source of Biosurfactants and antimicrobial Lipopeptides. Front Microbiol 2017;8:1–14. https://doi.org/10.3389/fmicb.2017.00061. [12] Manga EB, Celik PA, Cabuk A, Banat IM. Biosurfactants: opportunities for the development of a sustainable future. Curr Opin Colloid Interface Sci 2021;101514. https://doi.org/10.1016/j.cocis.2021.101514. [13] Patria RD, Wong JWC, Johnravindar D, Uisan K, Kumar R, Kaur G. Food waste digestate-based biorefinery approach for Rhamnolipids production: a techno-economic analysis. Sustain Chem 2021;2:237–53. https:// doi.org/10.3390/suschem2020014. [14] Zhao Z, Wong JWC. Biosurfactants from Acinetobacter Calcoaceticus BU03 enhance the solubility and biodegradation of Phenanthrene. Environ Technol 2009;30:291–9. https://doi.org/10.1080/ 09593330802630801. [15] Vecino X, Barbosa-Pereira L, Devesa-Rey R, Cruz JM, Moldes AB. Optimization of liquid–liquid extraction of biosurfactants from corn steep liquor. Bioprocess Biosyst Eng 2015;38:1629–37. https://doi.org/10.1007/ s00449-015-1404-9. [16] Vecino X, Barbosa-Pereira L, Devesa-Rey R, Cruz JM, Moldes AB. Study of the surfactant properties of aqueous stream from the corn milling industry. J Agric Food Chem 2014;62:5451–7. https://doi.org/ 10.1021/jf501386h. [17] Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Vecino X, Cruz JM, Moldes AB. Extraction, separation and characterization of Lipopeptides and phospholipids from corn steep water. Sep Purif Technol 2020;248. https://doi. org/10.1016/j.seppur.2020.117076, 117076. [18] Lo´pez-Prieto A, Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Moldes AB, Cruz JM. Effect of biosurfactant extract obtained from the corn-milling industry on probiotic bacteria in drinkable yogurt. J Sci Food Agric 2019;99:824–30. https://doi.org/10.1002/jsfa.9251. [19] Rodrı´guez-Lo´pez L, Vecino X, Barbosa-Pereira L, Moldes AB, Cruz JM. A multifunctional extract from corn steep liquor: antioxidant and surfactant activities. Food Funct 2016;7:3724–32. https://doi.org/10.1039/ c6fo00979d. [20] Lo´pez-Prieto A, Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Cruz JM, Moldes AB. Characterization of extracellular and cell bound Biosurfactants produced by Aneurinibacillus Aneurinilyticus isolated from commercial corn steep liquor. Microbiol Res 2021;242. https://doi.org/10.1016/j.micres.2020.126614, 126614. [21] Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Vecino X, Cruz JM, Moldes AB. Preservative and irritant capacity of biosurfactants from different sources: a comparative study. J Pharm Sci 2019;108:2296–304. https://doi.org/ 10.1016/j.xphs.2019.02.010.

150

Chapter 7 Their application in different industries

[22] Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Vecino X, Moldes AB, Cruz JM. Biodegradability study of the biosurfactant contained in a crude extract from corn steep water. J Surfactant Deterg 2020;23:79–90. https://doi. org/10.1002/jsde.12338. ´ lvarez M, P
erez-Davila S, Serra J, Gonza´lez P, Cruz JM, [23] Rodrı´guez-Lo´pez L, Lo´pez-Prieto A, Lopez-A Moldes AB. Characterization and cytotoxic effect of Biosurfactants obtained from different sources. ACS Omega 2020;5:31381–90. https://doi.org/10.1021/acsomega.0c04933. [24] Vecino X, Rodrı´guez-Lo´pez L, Cruz JM, Moldes AB. Sewage sludge polycyclic aromatic hydrocarbon (PAH) decontamination technique based on the utilization of a Lipopeptide biosurfactant extracted from corn steep liquor. J Agric Food Chem 2015;63:7143–50. https://doi.org/10.1021/acs.jafc.5b02346. [25] Bustos-Va´zquez G, Vidal-Fontela A, Vecino-Bello X, Cruz-Freire JM, Moldes-Menduin˜a AB. Uso de biosurfactantes extraidos de los licores de lavado de maı´z para la eliminacio´n de aceite quemado de motor en suelo arenoso; 2018. p. 581–91. [26] Perez-Ameneiro M, Vecino X, Cruz JM, Moldes AB. Wastewater treatment enhancement by applying a Lipopeptide biosurfactant to a lignocellulosic biocomposite. Carbohydr Polym 2015;131:186–96. https:// doi.org/10.1016/j.carbpol.2015.05.075. [27] Nitschke M, Marangon CA. Microbial surfactants in nanotechnology: recent trends and applications. Crit Rev Biotechnol 2021;0:1–17. https://doi.org/10.1080/07388551.2021.1933890. [28] Vallet-Regı´ M, Balas F, Arcos D. Mesoporous materials for drug delivery. Angew Chem Int Ed 2007;46:7548–58. https://doi.org/10.1002/anie.200604488. [29] Li N, Zhao P, Astruc D. Anisotropic gold nanoparticles: synthesis, properties, applications, and toxicity. Angew Chem Int Ed 2014;53:1756–89. https://doi.org/10.1002/anie.201300441. [30] Gomez-Gran˜a S, Le Beulze A, Treguer-Delapierre M, Mornet S, Duguet E, Grana E, Cloutet E, Hadziioannou G, Leng J, Salmon JB, et al. Hierarchical self-assembly of a bulk metamaterial enables isotropic magnetic permeability at optical frequencies. Mater Horiz 2016;3:596–601. https://doi.org/10.1039/c6mh00270f. [31] Saha K, Agasti SS, Kim C, Li X, Rotello VM. Gold nanoparticles in chemical and biological sensing. Chem Rev 2012;112:2739–79. https://doi.org/10.1021/cr2001178. [32] Vecino X, Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Cruz JM, Moldes AB. Nanomaterials synthesized by biosurfactants. Compr Anal Chem 2021;94:267–301. https://doi.org/10.1016/bs.coac.2020.12.008. [33] Płaza GA, Chojniak J, Banat IM. Biosurfactant mediated biosynthesis of selected metallic nanoparticles. Int J Mol Sci 2014;15:13720–37. https://doi.org/10.3390/ijms150813720. [34] Go´mez-Gran˜a S, Perez-Ameneiro M, Vecino X, Pastoriza-Santos I, Perez-Juste J, Cruz JM, Moldes AB. Biogenic synthesis of metal nanoparticles using a biosurfactant extracted from corn and their antimicrobial properties. Nanomaterials 2017;7. https://doi.org/10.3390/nano7060139. [35] Narayanan J, Ramji R, Sahu H, Gautam P. Synthesis, stabilisation and characterisation of rhamnolipidcapped ZnS nanoparticles in aqueous medium. IET Nanobiotechnol 2010;4:29–34. https://doi.org/ 10.1049/iet-nbt.2009.0010. [36] Basnet M, Ghoshal S, Tufenkji N. Rhamnolipid biosurfactant and soy protein act as effective stabilizers in the aggregation and transport of palladium-doped Zerovalent iron nanoparticles in saturated porous media. Environ Sci Technol 2013;47:13355–64. https://doi.org/10.1021/es402619v. [37] Satpute SK, Kulkarni GR, Banpurkar AG, Banat IM, Mone NS, Patil RH, Cameotra SS. Biosurfactant/s from lactobacilli species: properties, challenges and potential biomedical Applications. J Basic Microbiol 2016;56:1140–58. https://doi.org/10.1002/jobm.201600143. [38] Vecino X, Cruz JM, Moldes AB, Rodrigues LR. Biosurfactants in cosmetic formulations: trends and challenges. Crit Rev Biotechnol 2017;37:911–23. https://doi.org/10.1080/07388551.2016.1269053. [39] Adu SA, Naughton PJ, Marchant R, Banat IM. Microbial biosurfactants in cosmetic and personal skincare pharmaceutical formulations. Pharmaceutics 2020;12:1–21. https://doi.org/10.3390/ pharmaceutics12111099.

References

151

[40] Ceresa C, Fracchia L, Fedeli E, Porta C, Banat IM. Recent advances in biomedical, therapeutic and pharmaceutical applications of microbial surfactants. Pharmaceutics 2021;13. https://doi.org/10.3390/ pharmaceutics13040466. [41] Ismail R, Baaity Z, Cso´ka I. Regulatory status quo and prospects for biosurfactants in pharmaceutical applications. Drug Discov Today 2021;26:1929–35. https://doi.org/10.1016/j.drudis.2021.03.029. [42] Moldes AB, Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Lo´pez-Prieto A, Vecino X, Cruz JM. Synthetic and bioderived surfactants versus microbial biosurfactants in the cosmetic industry: an overview. Int J Mol Sci 2021;22:1–23. https://doi.org/10.3390/ijms22052371. [43] Ferreira A, Vecino X, Ferreira D, Cruz JM, Moldes AB, Rodrigues LR. Novel cosmetic formulations containing a biosurfactant from lactobacillus paracasei. Colloids Surf B Biointerfaces 2017;155:522–9. https://doi. org/10.1016/j.colsurfb.2017.04.026. [44] Farias CBB, Almeida FCG, Silva IA, Souza TC, Meira HM, Soares da Silva RDCF, Luna JM, Santos VA, Converti A, Banat IM, et al. Production of green surfactants: market prospects. Electron J Biotechnol 2021;51:28–39. https://doi.org/10.1016/j.ejbt.2021.02.002. [45] Morita T, Fukuoka T, Imura T, Kitamoto D. Mannosylerythritol lipids: production and applications. J Oleo Sci 2015;64:133–41. https://doi.org/10.5650/jos.ess14185. [46] Rinco´n-Fonta´n M, Rodrı´guez-Lo´pez L, Vecino X, Cruz JM, Moldes AB. Adsorption of natural surface active compounds obtained from corn on human hair. RSC Adv 2016;6:63064–70. https://doi.org/10.1039/ c6ra13823c. [47] Rinco´n-Fonta´n M, Rodrı´guez-Lo´pez L, Vecino X, Cruz JM, Moldes AB. Influence of micelle formation on the adsorption capacity of a biosurfactant extracted from corn on dyed hair. RSC Adv 2017;7:16444– 52. https://doi.org/10.1039/c7ra01351e. [48] Rinco´n-Fonta´n M, Rodrı´guez-Lo´pez L, Vecino X, Cruz JM, Moldes AB. Novel multifunctional biosurfactant obtained from corn as a stabilizing agent for antidandruff formulations based on Zn pyrithione powder. ACS Omega 2020. https://doi.org/10.1021/acsomega.9b03679. [49] Yuan CL, Xu ZZ, Fan MX, Liu HY, Xie YH, Zhu T. Study on characteristics and harm of surfactants. J Chem Pharm Res 2014;6:2233–7. [50] Knoth D, Rinco´n-Fonta´n M, Stahr PL, Pelikh O, Eckert RW, Dietrich H, Cruz JM, Moldes AB, Keck CM. Evaluation of a biosurfactant extract obtained from corn for dermal application. Int J Pharm 2019;564:225– 36. https://doi.org/10.1016/j.ijpharm.2019.04.048. [51] Rinco´n-Fonta´n M, Rodrı´guez-Lo´pez L, Vecino X, Cruz JM, Moldes AB. Design and characterization of greener sunscreen formulations based on mica powder and a biosurfactant extract. Powder Technol 2018;327. https://doi.org/10.1016/j.powtec.2017.12.093. [52] Rinco´n-Fonta´n M, Rodrı´guez-Lo´pez L, Vecino X, Cruz JM, Moldes AB. Study of the synergic effect between mica and Biosurfactant to stabilize Pickering emulsions containing vitamin E using a triangular design. J Colloid Interface Sci 2019;537:34–42. https://doi.org/10.1016/j.jcis.2018.10.106. [53] Rinco´n-Fonta´n M, Rodrı´guez-Lo´pez L, Vecino X, Cruz JM, Moldes AB. Potential application of a multifunctional biosurfactant extract obtained from corn as stabilizing agent of vitamin C in cosmetic formulations. Sustain Chem Pharm 2020;16. https://doi.org/10.1016/j.scp.2020.100248. [54] Rodrı´guez-Lo´pez L, Rinco´n-Fonta´n M, Vecino X, Cruz JM, Moldes AB. Study of biosurfactant extract from corn steep water as a potential ingredient in Antiacne formulations. J Dermatolog Treat 2020;1–8. https://doi. org/10.1080/09546634.2020.1757016. [55] Rodrı´guez-Lo´pez L, Shokry DS, Cruz JM, Moldes AB, Waters LJ. The effect of the presence of biosurfactant on the permeation of pharmaceutical compounds through silicone membrane. Colloids Surf B Biointerfaces 2019;176:456–61. https://doi.org/10.1016/j.colsurfb.2018.12.072. [56] Sachdev DP, Cameotra SS. Biosurfactants in agriculture. Appl Microbiol Biotechnol 2013;97: 1005–16. https://doi.org/10.1007/s00253-012-4641-8.

152

Chapter 7 Their application in different industries

[57] Kim SK, Kim YC, Lee S, Kim JC, Yun MY, Kim IS. Insecticidal activity of rhamnolipid isolated from pseudomonas Sp. EP-3 against green peach aphid (Myzus Persicae). J Agric Food Chem 2011;59:934–8. https:// doi.org/10.1021/jf104027x. [58] Sharma D, Kalita S, Duarah K. Isolation and characterization of bio surfactant producing micro organisms and to determine i t’ s in vitro antagonistic activity against Phytopathogenic fungi. Int J Curr Trends Sci Technol 2017;7:20355–67. [59] Blackwell PS. Management of water Repellency in Australia, and risks associated with preferential flow, Pesticide Concentration and Leaching. J Hydrol 2000;231–232:384–95. https://doi.org/10.1016/S00221694(00)00210-9. [60] Petrovi
c M, Barcelo´ D. Analysis and fate of surfactants in sludge and sludge-amended soils. TrAC - Trends Anal Chem 2004;23:762–71. https://doi.org/10.1016/j.trac.2004.07.015. [61] Lo´pez-Prieto A, Vecino X, Rodrı´guez-Lo´pez L, Moldes AB, Cruz JM. A multifunctional biosurfactant extract obtained from corn steep water as bactericide for agrifood industry. Foods 2019;8:410. https://doi.org/ 10.3390/foods8090410. [62] Lo´pez-Prieto A, Vecino X, Rodrı´guez-Lo´pez L, Moldes AB, Cruz JM. Fungistatic and fungicidal capacity of a biosurfactant extract obtained from corn steep water. Foods 2020;9:662. https://doi.org/10.3390/ foods10061318. [63] Naughton PJ, Marchant R, Naughton V, Banat IM. Microbial biosurfactants: current trends and applications in agricultural and biomedical industries. J Appl Microbiol 2019;127:12–28. https://doi.org/10.1111/ jam.14243. [64] Campos JM, Stamford TLM, Sarubbo LA. Production of a bioemulsifier with potential application in the food industry. Appl Biochem Biotechnol 2014;172:3234–52. https://doi.org/10.1007/s12010-014-0761-1. [65] Sałek K, Euston SR. Sustainable Microbial biosurfactants and bioemulsifiers for commercial exploitation. Process Biochem 2019;85:143–55. https://doi.org/10.1016/j.procbio.2019.06.027. [66] Martı´nez-Arcos A, Lo´pez-Prieto A, Rodrı´guez-Lo´pez L, P
erez-Cid B, Vecino X, Moldes AB, Manuel CJ. Evaluation of morphological changes in grapes coated with a biosurfactant extract obtained from corn steep liquor. Appl Sci 2021;11. https://doi.org/10.3390/app11135904. [67] Scalzini G, Lo´pez-Prieto A, Paissoni MA, Englezos V, Giacosa S, Rolle L, Gerbi V, Segade SR, Cid BP, Moldes AB, et al. Can a corn-derived biosurfactant improve colour traits of wine? First insight on its application during Winegrape skin maceration versus oenological tannins. Foods 2020;9:1–23. https://doi.org/ 10.3390/foods9121747. ´ , Martı´nez Urbina MA ´ , Lo´pez y Lo´pez VE. Advances on research in the use of agro[68] Domı´nguez Rivera A Industrial waste in biosurfactant production. World J Microbiol Biotechnol 2019;35:1–18. https://doi.org/ 10.1007/s11274-019-2729-3. [69] Chebbi A, Franzetti A, Duarte Castro F, Gomez Tovar FH, Tazzari M, Sbaffoni S, Vaccari M. Potentials of winery and olive oil residues for the production of Rhamnolipids and other biosurfactants: a step towards achieving a circular economy model. Waste Biomass Valoriz 2021;12:4733–43. https://doi.org/10.1007/ s12649-020-01315-8. [70] Mohanty SS, Koul Y, Varjani S, Pandey A, Ngo HH, Chang JS, Wong JWC, Bui XT. A critical review on various feedstocks as sustainable substrates for biosurfactants production: a way towards cleaner production. Microb Cell Fact 2021;20:1–13. https://doi.org/10.1186/s12934-021-01613-3. € [71] Alkan Z, Erginkaya Z, Konuray G, Unal Turhan E. Production of biosurfactant by lactic acid bacteria using whey as growth medium. Turk J Vet Anim Sci 2019;43:676–83. https://doi.org/10.3906/vet-1903-48. [72] Daverey A, Pakshirajan K, Sumalatha S. Sophorolipids production by Candida Bombicola using dairy industry wastewater. Clean Techn Environ Policy 2011;13:481–8. https://doi.org/10.1007/s10098-010-0330-4.

References

153

[73] Rocha MVP, Oliveira AHS, Souza MCM, Gonc¸alves LRB. Natural cashew apple juice as fermentation medium for biosurfactant production by acinetobacter calcoaceticus. World J Microbiol Biotechnol 2006;22:1295–9. https://doi.org/10.1007/s11274-006-9175-8. [74] da Silva GF, Gautam A, Silveira Duarte IC, Delforno TP, de Oliveira VM, Huson DH. Interactive analysis of biosurfactants in fruit-waste fermentation samples using BioSurfDB and MEGAN. bioRxiv 2021. 2021.11.11.468240. [75] Chooklin CS, Maneerat S, Saimmai A. Utilization of Banana Peel as a novel substrate for biosurfactant production by halobacteriaceae archaeon AS65. Appl Biochem Biotechnol 2014;173:624–45. https://doi.org/ 10.1007/s12010-014-0870-x. [76] Gurjar J, Sengupta B. Production of Surfactin from Rice mill polishing residue by submerged fermentation using bacillus subtilis MTCC 2423. Bioresour Technol 2015;189:243–9. https://doi.org/10.1016/j. biortech.2015.04.013. [77] de Andrade CJ, de Andrade LM, Rocco SA, Sforc¸a ML, Pastore GM, Jauregi P. A novel approach for the PRODUCTION and purification of mannosylerythritol lipids (MEL) by Pseudozyma Tsukubaensis using cassava wastewater as substrate. Sep Purif Technol 2017;180:157–67. https://doi.org/10.1016/j. seppur.2017.02.045. [78] Konishi M, Yoshida Y, Horiuchi J. Ichi efficient production of sophorolipids by starmerella bombicola using a corncob hydrolysate medium. J Biosci Bioeng 2015;119:317–22. https://doi.org/10.1016/j. jbiosc.2014.08.007. [79] Panjiar N, Mattam AJ, Jose S, Gandham S, Velankar HR. Valorization of xylose-rich hydrolysate from rice straw, an Agroresidue, through biosurfactant production by the soil bacterium Serratia Nematodiphila. Sci Total Environ 2020;729. https://doi.org/10.1016/j.scitotenv.2020.138933. [80] Chen C, Sun N, Li D, Long S, Tang X, Xiao G, Wang L. Optimization and characterization of biosurfactant production from kitchen waste oil using pseudomonas aeruginosa. Environ Sci Pollut Res 2018;25: 14934–43. https://doi.org/10.1007/s11356-018-1691-1. [81] Sharma S, Verma R, Dhull S, Maiti SK, Pandey LM. Biodegradation of waste cooking oil and simultaneous production of rhamnolipid biosurfactant by pseudomonas aeruginosa P7815 in batch and fed-batch bioreactor. Bioprocess Biosyst Eng 2021. https://doi.org/10.1007/s00449-021-02661-0. [82] Luna JM, Rufino RD, Jara AMAT, Brasileiro PPF, Sarubbo LA. Environmental Applications of the biosurfactant produced by Candida Sphaerica cultivated in low-cost substrates. Colloids Surf A Physicochem Eng Asp 2015;480:413–8. https://doi.org/10.1016/j.colsurfa.2014.12.014. [83] Radzuan MN, Winterburn J, Banat I. Bioreactor rhamnolipid production using palm oil agricultural refinery by-products. Processes 2021;9:1–15.

CHAPTER

Metabolic and process engineering on the edge—Rhamnolipids are a true challenge: A review

8

Melanie Filbiga, Sonja Kubickib, Isabel Batora, Rudolf Hausmannc, Lars Mathias Blanka, Marius Henkelc, Stephan Thiesb, and Till Tisoa a

iAMB - Institute of Applied Microbiology, ABBt - Aachen Biology and Biotechnology, RWTH Aachen University, € Dusseldorf, € € Julich, Aachen, Germany, bInstitute of Molecular Enzyme Technology, Heinrich-Heine-Universitat Germany, cDepartment of Bioprocess Engineering, University of Hohenheim, Institute of Food Science and Biotechnology, Stuttgart, Germany

1. Introduction Biosurfactants play an important role for the future chemical industry in the shift to ‘green’ compounds and the renunciation of fossil-based resources. Rhamnolipids (RL) are glycolipid biosurfactants produced by different microorganisms. They consist of one (mono-RL) or two (di-RL) rhamnose molecules in the hydrophilic part and a dimer of esterified fatty acids in the hydrophobic moiety (Fig. 1). The best-known native producer of RL, P. aeruginosa, is an opportunistic human pathogen, which complicates its use in industry. Furthermore, RL production with P. aeruginosa is subject to complicated fermentation strategies due to the complex regulation of RL production [1]. Alternative nonpathogenic native RL producers were reported, e.g., Burkholderia glumae [2], Thermus sp. [3] or Marinobacter [4], but none of them can compete with P. aeruginosa regarding RL titers. The recombinant production of RL in nonpathogenic hosts is an alternative to avoid the drawbacks that come along with the use of P. aeruginosa. Escherichia coli [5], Saccharomyces cerevisiae [6], Cellvibrio japonicus [7], and Pseudomonas putida [8] were enabled to produce RL when the rhlAB (C) operon from P. aeruginosa was expressed heterologously. One of the best-studied heterologous hosts for the biosynthesis of RL is the bacterium P. putida KT2440 [9–19]. Mono-RL biosynthesis in this strain is enabled by the expression of the genes rhlA, encoding for an acyltransferase that produces a dimer of esterified fatty acids (HAA), and rhlB, encoding for rhamnosyltransferase I. Additional expression of rhlC, encoding for rhamnosyltransferase II, allows for the production of di-RL (Fig. 1). Since the pathways yielding the precursors activated β-hydroxy-fatty acid and dTDP-L-rhamnose are natively present in P. putida, expression of rhlA and rhlB(C) is sufficient to achieve RL biosynthesis. Further advantages of this strain include the tolerance to RL titers of up to 90 g L-1 rendering it suitable as a host for large-scale biotechnological RL production [17]. So far, no recombinant P. putida RL processes or strains have been published that achieve specific productivities or titers comparable to those of the native producer P. aeruginosa. The reason for this is Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00008-9 Copyright # 2023 Elsevier Inc. All rights reserved.

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FIG. 1 RL molecule. Arrows indicate bonds that are formed during RL biosynthesis and the corresponding enzymes.

largely a matter of speculation. One reason might be low expression of the synthesis genes, which might be counteracted by the means discussed in Section 2. The limited availability of precursor molecules is another possible reason for lower RL titers achieved with the recombinant producer. Possibilities to overcome this limitation are discussed in Section 3. In this chapter, we summarize the studies carried out regarding (1) cloning of optimized expression cassettes for recombinant RL production using P. putida, (2) construction of enhanced P. putida chassis strains to further enhance RL production, and (3) fermentation approaches on lab scale using recombinant P. putida strains.

2. Design of an optimal expression cassette As introduced previously, P. putida KT2440 does not naturally produce RL or any other biosurfactant by itself; hence, all those approaches rely on successfully expressing rhlAB operon and rhlC, the respective biosynthetic genes, in recombinant strains [20–22]. Therefore, the expression module is the heart of each P. putida RL production strain and much effort has been put into the development of suitable strategies. In a nutshell, five aspects (IdV) gathered particular interest within this process (Fig. 2): Obviously, the selection of genes (I) encoding RL biosynthesis (i.e., variants of rhlA, rhlB, rhlC) is the first step [23,24]. Those genes are combined (II) with promoter sequences to establish/control the gene expression [25]. Particular emphasis might be (III) spent on the design of the untranslated regions (UTR) of the resulting transcript influencing the translation initiation of the biosynthetic genes [26,27]. The fast-growing biobricks toolbox derived within the conceptual frame of synthetic biology offers an increasing number of opportunities to tune both transcription and translation to this end. The fourth (IV) aspect is the inclusion of accessory genes, i.e., genes whose transfer is not essential to establish RL biosynthesis in P. putida, but help to improve the yields. This can be achieved by enhancing the pool of precursor molecules provided by the native metabolism for rhlA and rhlB/rhlC [28,29]. Finally, the biosynthetic module for RL synthesis constructed this way has to be implemented (V) in P. putida, e.g., using plasmid or chromosomal integrons [20]. In the following section, we will highlight the strategies explored so far regarding each of these aspects. Notably, systematic expression module construction often overlaps with engineering of the host metabolism (see Section 3) and, hence, both cannot be considered independent of each other in an integrated strain development strategy.

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159

FIG. 2 Critical aspects of an expression module to construct P. putida strains for RL production.Choice of biosynthetic genes (I, green) with regard to the desired product, and expression parameters e.g., using strong promoters (II, orange) like the constitutive synthetic 14-ffg stacked promoter or the salicylate inducible nagAa-promoter, UTR engineering (III, blue) e.g., by standardizing translation initiation through the application of bicistronic designs (BCD), optional inclusion of accessory genes (IV, violet) for improved production performance and finally the mode of implementation of the expression module (V, gray) (i.e., plasmid or chromosomal integration).

2.1 Biosynthetic genes The core of an expression module for RL production with P. putida is, of course, constituted by the genes encoding the biosynthetic enzymes, RhlA and RhlB for the synthesis of mono-RL from activated rhamnose and hydroxy fatty acids, and, if desired, RhlC for the conversion of mono-RL to di-RL. Most studies so far rely on genes amplified from P. aeruginosa [20,29], probably because of the two species’ phylogenetic relationship and finding a nonpathogenic alternative to specifically P. aeruginosa for less complex regulated RL biosynthesis being the primary motivation to develop the P. putida

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RL production platform [29]. RL are always produced as a mixture of molecules. Since those molecules are structurally related, they are referred to as congeners. The congeners in the RL mixture can differ in the number of rhamnose moieties but also in the amount of hydroxy fatty acids (one or two), the chain length, as well as the degree of unsaturation in those fatty acids. P. aeruginosa produces a characteristic spectrum of RL congeners consisting mainly of di-RL, with minor fractions of mono-RL and HAA. The carbon chain spectrum includes fatty acids with lengths between C8 and C12, but the main congener for mono- and di-RL, as well as for HAA, is the molecule containing two C10 chains. Expression of the P. aeruginosa rhl genes in other organisms yields the same congener distribution as reported for the donor [24]. More recent studies showed that the origin of the rhl genes influences the congener spectrum produced by the respective P. putida strains. Species of Burkholderia predominantly produce RL with a C14–C14 lipid moiety [2,30], while synthesis of RL with a C16–C16 lipid moiety has been reported for Thermus sp. [3]. Since mainly the acyltransferase RhlA determines the chain length of the hydrophobic moiety in the resulting HAA/RL [19], the choice of the rhlA-origin for the recombinant production has a great impact. When rhlAB(C) from Burkholderia glumae PG1 were expressed in P. putida, the host produced RL-mixtures typical for Burkholderia and more hydrophobic than with the P. aeruginosa genes [19]. Germer et al. tested eight rhlA homologs from different organisms in E. coli and produced 53 different HAA congeners with chain lengths reaching from C4 to C18 and including mono saturations, as well as odd chain lengths [24]. However, the titers achieved with P. putida expressing B. glumae genes were much lower than comparable strains expressing P. aeruginosa genes [19]. The reasons behind this discrepancy have not yet been investigated. A lower specific activity of the enzymes or limited availability of specific 3-hydroxy fatty acids were discussed to this end. It might also be possible that long-chain RL production exhibits more stress to the host cells, or that the high GC-content of the Burkholderia sp. derived genes prevent an efficient expression comparable to the Pseudomonas derived genes. Indeed, codon adaption of rhl genes has not explicitly been investigated so far in P. putida RL production strain development but synthetic and probably optimized operons were already applied successfully. For example, Gehring et al. report the use of an artificially synthesized operon encoding P. aeruginosa-like rhlABC to successfully establish production of di-RL from butane, but whether codon adaption was considered here remains unclear and was at least not explicitly investigated [31]. Studies with engineered S. cerevisiae and E. coli indicated that codon adaption might indeed help to improve the production of RL [6,32]. The selection between a single expression of rhlA or co-expression of rhlAB or even rhlABC can further control the composition of the product: the single expression of rhlA yields HAA only, the addition of rhlB yields mono-RL and HAA and the further addition of rhlC yields a mixture of di-RL, mono-RL and HAA. Mixing rhl genes from different origins was shown to yield variations in the RL composition in E. coli [32]. Modifications of the rhl genes, e.g., by exchanging segments of rhl genes from different species, as has been shown for Burkholderia, further increase the possibility to produce designer RL [33]. It has further been shown that the properties of the resulting RL produced in E. coli can be altered by directed mutagenesis of the rhlB gene [34]. Since the physico-chemical properties including the critical micelle concentration (CMC) or the emulsification behavior heavily depend on the structure of RL, the possibility to create tailored molecules with specific properties is of high interest.

2.2 Promoter The second essential brick to think of while constructing an expression module is a promoter element to control the expression of the rhl genes of choice (Table 1).

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Table 1 Overview on rhl expression modules so far applied for RL production in P. putida Promoter/ origin

50 -UTR/RBS of rhlA

Expression mode

Ptac + lacI/ synthetic

50 UTRlacX RBSrhlA

Inducible (if LacI is provided)

50 UTRrhlA

Constitutive

Vector Plasmid

Plasmid

SynPro variants / synthetic

SPL variants/ synthetic, based on PrRNA RhaRSrhaPBAD/ E. coli NagR- PnagAa/ Comamonas testosteroni

mRL/0.6– 2.2 g L
1 HAA/0,05 g L
1 mRL/0,05 g L
1 mRL and dRL/ 0,015 and 0,22 g L
1 Long chain mRL and dRL/0,075 and 0,059 g L
1 HAA/1.5 g L
1 mRL/0.2 to 14.9 g L
1

References [8,14,17,41] [18], [42,43]

[19]

[15] [15], [11,12], [14], [42,43], [9], [44,45], [13,16,46] [15] [40]

5 UTRrhlA (exchanged four U RNAThairpin2/S. enterica) Biscistronic design BCD2

Temperaturedependent

Plasmid

dRL/3.3 g L
1 mRL/2.15 to 2.42 g L
1

Constitutive

Tn7 transposon

mRL/0.72 to 2.23 g L
1

[10,37], [16,38,47]

50 UTRrhlA

Autoinducer, based on RhlI synthesized AHL

Plasmid

mRL/0.02 to 7.3 g L
1 mRL/1.68 g L
1

[8,36]

mRL/>0.01 g L
1

[48]

0

Pffg /stacked synthetic promoters RhlR/I - Prhl via native/P. aeruginosa

Product/range of achieved titers

50 UTRrhlA

Constitutive

Tn5 transposon Plasmid

50 UTRrhlA

Inducible

Plasmid

dRL/1.2 g L
1

[31]

Biscistronic design BCD2

Inducible

Tn7 transposon Plasmid

mRL/1.3 g L
1

[16]

mRL/0.73 g L
1

[39]

[35]

In P. aeruginosa, the expression of rhlAB is controlled, among others, by a homoserine lactone binding transcription factor, i.e., RhlR. The respective quorum-sensing signal molecules are synthesized by RhlI [49]. The genes encoding both proteins are on the P. aeruginosa chromosome located downstream of rhlAB. By transferring the whole rhlABRI cluster into P. putida to maintain the native regulation system, RL titers up to the g L
1 scale were achieved [35,36].

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During the development of P. putida toward a synthetic biology chassis, a versatile set of wellcharacterized, adjustable expression systems has become available. These include several libraries of synthetic constitutive promoters with defined activities and inducible promoters with low leakiness (e.g., XylS/Pm, RhaRS/PrhaBAD, nagR/PnagAa, AraC/ParaB) [25,50,51]. Aiming to establish and enhance autoinducer-independent expression of the RL biosynthesis in P. putida, artificial promoter systems were applied in most studies. RL biosynthesis in P. putida was demonstrated using the inducible Ptac. In different approaches synthetic promoters were used, resulting in the highest titers and yields reported so far [12,29]. Tiso et al. investigated a library of synthetic promoters for the plasmid-based expression of rhlAB(C) from P. aeruginosa, with the highest titer of 2.8 g L
1 being produced using promoter pSynPro8 [14], which was therefore chosen for several other studies [9,11,12,15,18]. The most recent approach utilized three strong synthetic promoters stacked in a row to achieve the strongest expression of rhl genes [10,37,38,52]. Besides, some inducible promoter systems were explored, i.e., RhaRS/PrhaBAD and nagR/PnagAa [16,31,39]. A recent comparative study indicated that inducible systems with low leakiness might have remarkable advantages in strain stability compared to strong constitutive promoters and their transcript levels are comparable [16]. Hence, such systems might be an option for large-scale processes that necessitate several rounds of seed fermentations. Given the wealth of controllable systems available, the fact that only three inducible expression systems have been explored so far leaves space for further studies to characterize other systems concerning production performance and strains stability. Differential fine-tuning of expression strengths of rhlA, rhlB, and rhlC by assigning a specific promoter to each gene can further be used to adjust the proportions of HAA/ mono-RL/di-RL and hence might facilitate the control of product composition. Tiso et al. illustrated this approach and showed that with transcriptional uncoupling of rhlC from rhlAB a high proportion of di-RL in the final RL mixture can be achieved [15].

2.3 Untranslated regions and translation initiation Considering UTR as a critical element of expression modules besides the two essential parts and design of UTR as another tuning knob for gene expression came up with the biobrick concept of synthetic biology. Functional and engineerable mRNA elements for Pseudomonas biotechnology are ribosome binding sites (RBSs), riboregulators (riboswitches) that cover the RBS upon the respective signal, and ribozymes [25,27]. Further, translation initiation might also be unintendedly affected by interaction between the 50 end of the gene of interest (GOI) and the UTR leading to secondary structure formation, which must be kept in mind when combining designed elements into an expression module [53]. In this context, the sequence space between RBS and start codon influences translation initiation [54,55]. Finally, RNAse recognition sites in the UTR (or the target genes themselves) might deplete the transcript and decrease the overall expression of the GOI [56,57]. In a nutshell, UTR design may be motivated by two goals: (i) introduce regulatory elements on mRNA level and (ii) avoid undesired regulation of the expression module or interference with the intended control mechanisms. Only a few studies so far targeted UTR design in the context of RL production with P. putida, but notably, most studies transferred the native UTR of P. aeruginosa rhlA along with rhlAB to the recombinant host (Table 1). Although not deliberately engineered, these constructs appear to contain a regulatory element in the form of an only recently described temperature-sensitive secondary structure element (RNA-thermometer) [40]. The concept of thermoregulation of rhlA expression was subsequently further exploited by the introduction of a recombinant four U RNA thermometer, showing that UTR engineering can be a practical approach to develop novel strategies to control P. putida RL producers.

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163

Besides, some recent studies replaced the original rhlA 50 UTR with a translation initiation concept called bicistronic design (BCD). This approach exploits a strong RBS for translation of the GOI (here: rhlA) along with the helicase activity of ribosomes to circumvent any interaction of the RBS with the UTR or the coding sequence of the GOI [16]. To this end, a short peptide (plus its RBS) is placed upstream of the GOI. The latter’s RBS is included in the coding sequence of the peptide. Upon translation of the peptide, the ribosome unfolds possibly occurring secondary structure that might hamper ribosome binding to the RBS in front of the GOI [58,59]. This way, a reliable sequence-independent strong translation initiation for the GOI is secured. The presented strategies indicate that UTR design might be a useful add-on to the currently applied toolbox for constructing expression modules for RL production with P. putida. Notably, both described approaches for UTR engineering only addressed rhlA, whereas the UTR /RBS of rhlB was not addressed so far to the best of our knowledge. The addition of an artificial RBS in combination with a synthetic promoter upstream of rhlC facilitated a high proportion of diRL [15]. The progressing synthetic biology provides further elements to explore UTR design, for example, aptamer-like riboregulators that bind small molecules [60], like the recently for P. putida established fluoroswitch [61], to address goal (i) and ribozymes to remove undesired UTR to address goal (ii) [27,62].

2.4 Accessory genes It might prove helpful to include genes beyond rhlABC within the expression module to increase yields or facilitate process design, even though such strategies have hardly ever been explored in P. putida. For example, the expression of additional genes might enhance the availability of precursor molecules (see Section 3.2). Critical genes for new-to-Pseudomonas catabolic pathways might also be integrated into the rhlAB expression module to facilitate strain construction. Moreover, reporter genes to follow the expression of the biosynthetic genes online might be valuable for process development.

2.5 Integration concepts: Genome-based vs plasmid-based expression The final step toward a P. putida RL production strain is the transfer of the thoughtfully designed rhl expression module into the host organisms. Traditionally, small operons comparable to rhlAB(C) have been predominantly introduced on plasmids, despite the potential metabolic burden of plasmid maintenance [31,63,64]. Plasmids enable the rapid creation of production strains and the flexible use of one construct in multiple strains [65]. Hence it is not surprising that most of the studies on RL production relied on the expression module encoding plasmids [20] (Table 1). On the other hand, intensive research during the last decades on tools for chromosomal integration via homologous recombination, recombinase-assisted recombination, or transposons, especially in the context of Pseudomonas strain development, resulted in a broad set of strategies to construct antibiotic-independent stable expression strains, even for large biosynthetic genes clusters [20,66,67]. A few, mostly very recent, studies already exploited transposons to stably integrate the expression module into the host’s chromosome. In particular, the Tn7 transposon was proven very useful to construct efficient RL production strains [16,37]. Notably, it was shown that, although the gene dose in the strains bearing plasmids was considerably higher than in the Tn7-born strains, the production performance was better and more reliable in the latter [16]. Especially in terms of a large-scale process where

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robustness (high stress tolerance), high product titers (resulting from balanced transcript levels and homogeneous expression), and low cost (omission of expensive additives such as antibiotics) are required [68,69], the use of plasmids to generate a stable production strain may be detrimental. It may be assumed that with the further progress of assembly cloning and genomic engineering methods, and identification of novel, suitable integration loci, chromosomal integration of rhl expression modules may become the favored strategy for P. putida RL production strain construction in the near future.

3. Development of an enhanced chassis cell Several strategies have been pursued to enhance the RL production in P. putida and other native or heterologous RL-producers by metabolic engineering (Fig. 3). In general, they can be classified into (I) enhancement of the precursor availability, (II) reduction of unwanted by-product formation and metabolic burden, (III) the accessibility of ‘uncommon’ substrates, and (IV) strain engineering for improved bioprocess characteristics. The (over-) expression of the involved key genes, i.e., the rhl genes, also plays an important role in metabolic engineering for heterologous RL production. The product spectrum can be adjusted to obtain a tailored product composition (see Section 2.2).

3.1 Enhancing precursor-availability The two main precursors for RL biosynthesis derive from central metabolic pathways: activated β-hydroxy fatty acids derived from fatty acid de novo synthesis is the precursor for the fatty acid moiety, while activated dTDP-L-rhamnose constitutes the sugar moiety. The synthesis of activated rhamnose requires glucose-6-phosphate, whereas fatty acid de novo synthesis depends on the central carbon metabolism intermediate acetyl-CoA. To improve the biosynthesis of RL the availability of precursor molecules plays an important role. First, glucose-6-phosphate is converted to glucose-1-phosphate by phosphoglucomutase (Pgm). dTDP-L-rhamnose is then formed from glucose-1-phosphate in four subsequent reactions catalyzed by RmlA, RmlB, RmlC, and RmlD and considered to be a bottleneck for RL production in P. putida [21]. When the rml operon, encoding for the enzymes involved in the formation of dTDP-L-rhamnose from glucose-1-phosphate, was overexpressed in E. coli, the availability of dTDP-L-rhamnose was increased. This also led to enhanced RL titers, revealing a limitation in rhamnose-availability for RL-biosynthesis in E. coli [5]. Intermediates of the biosynthesis pathway yielding activated rhamnose are also used for the synthesis of exopolysaccharides (EPS). The enzymes encoded by pslAB use dTDP-D-glucose for the synthesis of EPS and thus detract precursors from the RL biosynthesis [70]. The deletion of pslAB in P. aeruginosa increased the availability of dTDP-Lrhamnose and improved the RL yield by 21%. In addition, depletion of activated hydroxy fatty acids might also be considered a limiting factor. To enhance the flux of intermediates toward fatty acid de novo synthesis, which is involved in the synthesis of HAA, increasing the pool of acetyl-CoA is one option. Since acetyl-CoA is synthesized from pyruvate by pyruvate-dehydrogenase, this enzyme is one target for overexpression. Overexpression of the subunit acoA of the enzyme showed a positive impact on HAA production [71]. Alternatively, the acetyl-CoA pool can be increased by reducing the flux through the TCA cycle. By inactivating aceA, more acetyl-CoA is available for fatty acid de novo synthesis. A reduced flux through the TCA cycle has been shown to increase polyhydroxyalkanoate (PHA) production in P. putida [72]. Since PHA and

3 Development of an enhanced chassis cell

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FIG. 3 Metabolic engineering strategies for heterologous synthesis of RL in Pseudomonas putida. Enhancement of the precursor availability (I, green), reduction of unwanted by-product formation and metabolic burden (II, orange), the accessibility of ‘uncommon’ substrates (III, blue), and strain engineering for improved bioprocess characteristics (IV, purple) with applied examples.

HAA share the same precursor for their biosynthesis, this approach might also improve RL production. A further possibility to enhance the flux toward HAA synthesis is an increased flux through fatty acid de novo synthesis itself. The overexpression of fabG, involved in type II fatty acid biosynthesis, together with the expression of rhlABC also led to RL titers improved by 27% in E. coli [32].

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FabG catalyzes the generation of β-hydroxyacyl-ACP, which is used for HAA formation as the substrate of rhlA. An overexpression of FabG thus enhances the supply of precursors for RhlA and consequently for RL biosynthesis. To enhance the availability of activated hydroxy fatty acids, the coexpression of the FabG-similar β-ketoacyl reductase RhlG might further increase the RL production in P. putida KT2440. In P. aeruginosa, RhlG appears to almost supply alkanoic hydroxy acids exclusively to produce HAA and also RL [73]. Furthermore, fatty acid synthesis was increased sixfold in E. coli by overexpression of the acetyl-CoA carboxylase, which catalyzes the formation of malonyl-CoA from acetyl-CoA [74] and could also be applicable to heterologous RL biosynthesis.

3.2 Reduction of by-product formation and observed metabolic burden The introduction of genes for the synthesis of a heterologous product into P. putida KT2440 increases the metabolic burden on the cells. To reduce the overall metabolic burden, nonessential resource- and energy-consuming cellular activities can be deleted, allowing a higher supply of resources and energy for the target reaction. The additional elimination of unwanted byproduct formation further increases the pool of available intermediates and energy. Especially in Pseudomonads, the biosynthesis of RL competes for precursor molecules with the biosynthesis of PHA, which are formed from central metabolism intermediates deriving from fatty acid de novo synthesis or β-oxidation. While (R)-3hydroxyacyl-CoA, derived from β-oxidation, has been shown to be a shared precursor for RL and PHA biosynthesis in P. aeruginosa [75], the origin of the RL- and PHA-precursor in P. putida is not yet fully elucidated. Nevertheless, it is believed that the acyltransferase, encoded by rhlA, and PHA polymerases, encoded by phaC1 and phaC2, compete for the same precursor. Thus, the formation of PHA withdraws precursor molecules from the RL biosynthesis. To avoid this, the genes involved in the formation of PHA can be deleted in order to block PHA synthesis. By knocking out phaC1DC2DFI, Adbel-Mawgoud et al. reached a 13% increase in RL production with P. aeruginosa PA14 [75]. By simultaneous overexpression of rhlAB-R, a mutant deficient in phaG, phaC1, and phaC2 produced 59% more RL than the P. aeruginosa PA14 wild type [76]. RL biosynthesis with P. putida was increased sevenfold by interrupting phaC1 [17]. The single deletion of the (R)-3-hydroxyacyl-acyl carrier protein-coenzyme A transferase phaG resulted in a 27% increase in the RL titer produced by P. putida, while a complete knock-out of the PHA operon led to an increase in the RL concentration of 115% in the same strain [16]. A decrease of the activity of PHA synthase or (R)-3-hydroxyacyl-acyl carrier protein-coenzyme A transferase is also applied for the industrial production of RL in P. putida [77]. The apparent advantage of blocking PHA synthesis on RL production indicates a strong competition for precursors between the two pathways. A further approach to direct the carbon toward RL biosynthesis is the deletion of glucose dehydrogenase, encoded by gcd. In a gcd-deficient mutant of P. putida, the PHA-content was increased by 60%–100%, depending on the cultivation conditions, compared to the wild type [78,79]; RL production could be improved by 10% in the gcd-deletion strain [71]. In P. putida Δgcd, the periplasmatic conversion of glucose into gluconate was blocked and the redirected carbon flux could be used for product synthesis via central carbon metabolism. By reducing the carbon-flux toward unwanted by-products, more carbon can be channeled into the biosynthesis of the desired product. A different strategy to improve the biosynthesis of heterologous products is the improvement of energy availability. The assembly and export of the flagellar machinery for example demands high energy amounts; the lack of flagellar-motion further enhances the availability of ATP [80]. Deleting

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the flagellar machinery has been shown to enhance RL titers by 130% [16]. Combining the individual attempts could improve product biosynthesis by cumulative effects.

3.3 Accessibility of uncommon substrates Many recent studies targeted the production of RL from a variety of substrates besides glucose. While most of them, such as glycerol, do not require additional strain engineering, other substrates require genetic engineering to be made accessible. Due to its high abundance and its renewability, plant biomass is an interesting substrate for biotechnological processes. Besides glucose, xylose and arabinose are the most abundant sugars in lignocellulose. To efficiently use xylose for RL biosynthesis, P. putida was equipped with three different metabolic pathways (Weimberg pathway, Isomerase pathway, and Dahms pathway) followed by adaptive laboratory evolution. The highest RL titer of 0.72 g L
1 was achieved from xylose by the evolved strain equipped with the Weimberg pathway [10]. Arabinose was made accessible for P. putida by expression of the araBAD operon from E. coli [81]. Efforts have also been made to utilize waste streams for the production of RL in P. putida. For the use of small organic acids (formate, acetate, and propionate, as well as mixtures thereof), no engineering is necessary [9]. Additionally, ethanol represents an attractive feedstock since it can be produced biotechnologically from CO2 or it can be produced from biomass. To establish an efficient production process on ethanol, a combined approach of strain and process engineering was taken: while adaptive laboratory evolution resulted in a P. putida KT2440 mutant able to efficiently convert ethanol into RL, the substrate also worked as a defoamer in a bioreactor fermentation [10]. In a different approach Tiso et al. addressed the reduction of plastic waste: P. umsongensis GO16 was engineered to convert enzymatically hydrolyzed PET monomers into HAA, the RL precursor, as well as PHA [82]. Ackermann et al. implemented the metabolization of adipic acid, which is abundant in many plastics, by heterologous expression of the dcaAKJP genes from Acinetobacter baylyi and subsequent adaptive laboratory evolution [83]. Furthermore, P. putida KT2440 was engineered to consume ethylene glycol and convert it into PHA [84,85]. Another alternative substrate is sucrose, which is highly abundant in molasses. It was made accessible to P. putida by recombinant expression of a sucrose invertase [86]. Furthermore, lactose could be used as a substrate for RL production with P. aeruginosa by expression of the lacZY genes from E. coli since P. aeruginosa, just like P. putida, lacks the genes required for lactose catabolization [87,88]. Additionally, PHA production with P. putida from levoglucosan, appearing in biomass deconstruction, was enabled by the expression of the levoglucosan kinase from Lipomyces starkeyi. After additional expression of a β-glucosidase from Agrobacterium sp., the metabolization of cellobiosan was enabled [89]. Another approach is the efficient use of acetate for bioproduction with P. putida. The acetate assimilation pathway has been overexpressed to efficiently produce PHA from acetate with P. putida [90]. In order to use butane as a substrate for RL production in P. putida, Thum et al. introduced an alkB-type oxidoreductase from P. putida GPo1, which converts butane to butanol [77]. Overall, a broad substrate spectrum is made accessible by genetic engineering for bioconversion with P. putida enabling a sustainable bioproduction.

3.4 Strain engineering for improved process control Strain engineering was also performed in order to improve the behavior of the production strain in a bioreactor setup. The production of RL and their precursors, HAA, leads to excessive foaming of the fermentation broth in aerated bioreactors due to the amphiphilic structure of biosurfactant molecules.

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This can be used to separate the product via foam fractionation [91,92]. Disadvantageous for this approach is the adhesion of the cells in the foam, caused by hydrophobic cell surface structures, which leads to a reduction of the available biocatalyst in the bioreactor. Engineering P. putida KT2440 for reduced cell surface hydrophobicity, e.g., by deleting the flagellum, as well as the cell surface proteins LapA and LapF, led to reduced foam adhesion and thus allowed for an improved separation of the product from biomass [38]. As mentioned before, ethanol, which can be efficiently converted into RL by engineered P. putida KT2440, simultaneously serves as a defoamer by destabilizing the foam and thus allowing a stable fed-batch bioreactor cultivation [10]. Foaming can also be avoided by eliminating the need for aeration, which is required to allow for a sufficient oxygen supply for the aerobic growth of P. putida. Askitosai et al. thus enabled P. putida to grow and produce RL under oxygen limitation in a bioelectrochemical system (BES) by equipping the strain with the genes required to produce phenazines, acting as redox mediators [39].

3.5 Alternative genetic targets for improved RL synthesis Despite factors for a more efficient biosynthesis, other beneficial additional genes might be product transport-related genes. However, in contrast to fungal glycolipids, transport mechanisms for RL have not been described yet, neither in the native producers nor in P. putida. It is however well-known that co-expression of exporters can increase glycolipid yields and product tolerance [93]. Tiso et al. found evidence for the involvement of a major facilitator superfamily (MFS) transporter, which is colocalized with rhlC in P. aeruginosa, in RL biosynthesis or RL export: a P. aeruginosa mutant lacking PA1131 showed decreased extracellular di-RL titers [71]. To enhance the export of RL, the MFS transporter from P. aeruginosa, or an efflux transporter from B. thailandensis, was coexpressed in P. putida [77]. Further, it was shown that the autotransporter esterase EstA plays an important role in RL biosynthesis in P. aeruginosa: an overexpression of estA led to higher RL titers in the supernatant [94]. Expression of this gene or its homologue in recombinant hosts [95] might also boost RL production. As mentioned before, Noll et al. investigated RL production in P. putida under the control of the native P. aeruginosa ROSE-like RNA thermometer and showed that it can be used to control RL production by the cultivation temperature and thus offers the possibility to establish a temperature-dependent process [13]. However, the increase in the specific RL-production rate was mainly caused by a higher overall metabolic rate at the higher cultivation temperature. The approach was refined in a follow-up study using a fourU RNA thermometer from Salmonella to allow separating phases of preferred growth from product formation during bioreactor cultivation [40]. The RL biosynthesis in P. putida has been improved by several means of genetic engineering. Factors to be considered are, e.g., the expression level of involved genes. Besides, the deletion of by-product forming reactions competing for the same metabolic intermediates or energy units successfully enhanced RL titers in P. putida. Further modifications that have been proved successful in other bacteria are potential additional targets. By improving the RL-production in nonpathogenic hosts via genetic engineering, these biosurfactants can be produced efficiently with other bacteria than the pathogenic P. aeruginosa. Some of those approaches are also applied for recombinant large-scale industrial production of RL with P. putida by Evonik Industries AG [77]. More improvement can be achieved by combining process and genetic engineering to overcome existing challenges.

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4. Fermentation of P. putida for production of RL In general, fermentation strategies used in the production of biosurfactants are diverse, and no uniform concepts and strategies can be derived from existing approaches [37,45,47,91,96,97]. As such, batch, fed-batch, and continuous cultivation strategies have been reported for biosurfactant production in the past. Overall, the efficient production of biosurfactants is associated with several obstacles, the largest of which is excessive foaming. This challenge of biosurfactant-production processes is a major distinction to other bioprocesses and is one of the main topics discussed in the following section. To overcome challenges coming along with processes for the production of biosurfactants, several concepts have been investigated in the past, which are based on alternatives to conventional sparger aeration [47]. These different approaches are often less efficient, thus resulting in oxygen limitation. This may affect both growth and product formation negatively. Another concept to address high foaming is represented by in situ product removal methods such as foam fractionation [11,44,91,98]. As foam formation and coalescence of bubbles originating from the fermentation broth are complex processes with many different mechanisms and influencing factors involved [99], concepts for foam fractionation are mainly based on heuristic approaches. It is for this reason that reliable prediction of foam formation during the fermentation process is not yet possible. Due to the heterogeneity and different physical properties of microbial biosurfactants, especially regarding foaming, surface tension reduction, and emulsifying ability, processes to produce biosurfactants in bioreactors have to be adjusted to suit the individual target product (Fig. 4). Various bioreactor concepts for biosurfactant production have been described in the past, ranging from conventional stirred tank bioreactor designs [12] to less common designs such as rotating disk bioreactors [100].

4.1 Cultivation strategies Batch cultivation is a common and simple setup, which has been reported in many biosurfactant production processes in the past [101,102]. The production of RL using P. aeruginosa PAO1, a wellknown and widely used model organism, is feasible by a simple batch fermentation process [102]. In batch cultivations, no regulation of the amount of substrate in the bioreactor is performed. It is for this reason that using substrate limitation does not play a role in common batch cultivation setups. Furthermore, the applied batch cultivation strategies in RL production are subject to design restrictions due to the complex regulation of RL production being associated with the quorum sensing system in its native host P. aeruginosa [21]. In addition to batch fermentation setups, sequential batch processes have also successfully been used in RL production in the past [103,104]. One of the key advantages of using a fed-batch strategy, the ability to control substrate concentrations and growth rate by feeding, was successfully employed to achieve increased RL formation, both in wild-type P. aeruginosa [105] as well as in heterologous approaches [12]. In fed-batch culture, the production process is commonly divided into two phases [106]. In the first phase, biomass is produced while product formation is relatively low. In a consecutive phase, limitation of a substrate component or an external stimulus, trigger or enhance product formation. A two-stage fed-batch process for high-efficient production of RL was reported in the past to yield a final product concentration of 14.9 g L
1 [12]. Another cultivation strategy that was used for RL production is continuous mode of operation. This concept was investigated, however only comparably low dilution rates below 0.1 h
1 and consequently low overall productivities compared to batch or fed-batch processes could be achieved [107–109].

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FIG. 4 Process design considerations for the production of rhamnolipids in a bioreactor. Aspects to be considered are the cultivation strategy (I, green), the reactor design (II, orange), and the downstream processing (III, blue).

4.2 Design considerations and hardware concepts The microbial production of biosurfactants during cultivation in a bioreactor involves several critical design considerations. It was reported that a hydrophobic carbon source is beneficial for inducing biosurfactant production in P. aeruginosa due to its effect on the bacterial quorum sensing system [110] as well as by unspecific mechanisms and substrate availability resulting from using a two-phase system [111–113]. However, these two-phase systems must be properly dispersed to ensure efficient availability of substrates, as well as efficient biosurfactant production. If a hydrophobic carbon source is applied as part of a two-phase culture medium, it typically inhibits foaming initially [114,115]. However, the hydrophobic carbon source is consumed during the production process, which may lead to a reduced antifoaming effect. In addition, cleavage products such as fatty acids have an amphiphilic character that can in turn result in a state of increased foaming [116]. As such, limiting and countering foaming during biosurfactant production is one of the main challenges toward implementing production-scale efficient processes.

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4.2.1 Dispersion and mixing Due to their amphiphilic character, RL and biosurfactants, in general, are able to disperse hydrophobic substances in the culture broth [117] which are a substrate for the native producer P. aeruginosa. As such, inefficient dispersion occurs especially at the beginning of the process, where product concentrations are low. In many biosurfactant production processes, stirred tank reactors are equipped with radial impellers. The most common design is referred to as Rushton turbine, a flat disk with vertically oriented blades. Rushton turbines lack the ability to mix in axial direction, and are consequently responsible for inefficient dispersion of the hydrophobic carbon source due to phase separation on top of the fermentation medium [96]. In addition, Rushton turbines are suboptimally involving the ability to efficiently mix the fermentation broth at the end of the production process, where a frothed-up and consequently less dense constitution of the culture medium is present in the bioreactor. To counter these effects, different concepts have been proposed in the past which involve special stirrer layouts, a combination of axial and radial mixing due to multiple stirrers or the application of specific foam stirring concepts, typically using large paddles to achieve foam mixing [1].

4.2.2 Preventing and disrupting foam formation Problems with foam formation occur primarily in the production of biosurfactants with high foaming potential in aerated and agitated (stirred) processes that use hydrophilic carbon sources. Several strategies can be used to interrupt foam formation during fermentation. A different approach to control foaming is to prevent foam from occurring in the first place. For this purpose, antifoaming agents are usually used from the beginning of the cultivation process. In addition to chemical defoaming agents, optimized fermentation strategies can also be used to prevent or control foam formation (see Section 4.2.3). For the very common concept of using chemical defoamers, detailed reviews are given by Pelton [118] and Pugh [119]. Nevertheless, the mode of action on foam formation and control is complex and not yet fully understood [119]. Although the use of defoamers appears to be a simple and generally convenient method of foam control, these agents also have significant disadvantages. The first effect that can be observed upon the use of defoamers is its effect on the oxygen transfer rate [120]. Furthermore, defoamers are in many cases an additional cost factor, which cannot be neglected, especially considering the production of low-cost surfactants. In addition, further processing of biosurfactants is extremely difficult due to the similar physiological and chemical behavior of defoamers and biosurfactants. These problems result in higher overall costs for a biosurfactant production process. Mechanical means of foam destruction have been developed as an alternative to chemical defoamers. The main principles of mechanical foam destruction are shear stress and pressure within the foam breaker. In addition, the collision of the condensed foam with the primary foam and the walls of the bioreactor plays an important part in the concept of foam disruption [121]. A mechanical foam breaker can be attached to the agitator shaft [122], or placed either in a stirred tank reactor or in a bubble column bioreactor [123,124]. However, it should be considered that mechanical foam breakers may also have a relevant effect on production costs due to maintenance and power consumption [125,126]. Another possibility is the collecting of the foam expelled from the fermenter in a separate vessel, in which the foam collapses over time and can be refed into the fermenter. This has been shown in RL production using recombinant P. putida [16]. Nevertheless, due to excessive foaming, no additional carbon source could be fed. The fermentation thus had to be terminated after the batch phase. Using a genetically modified strain, which could use ethanol as sole carbon source (Section 3.4), the authors

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were able to increase the efficiency of the system. By spraying ethanol on the collapsing foam, the alcohol acts as a chemical defoamer; the foam collapsing could be significantly increased. With this setup even fed-batch fermentation could be facilitated [37].

4.2.3 Design of foam-free production processes One strategy for avoiding foam formation that was reported in the past is the rotating disk bioreactor [100]. By using this concept, foam formation is dramatically limited by avoiding gas bubbles in the liquid. Oxygen supply is enabled by surface contact of the fermentation broth, as well as on the rotating disks. It should be noted however, that limitations in oxygen transfer were observed resulting in low overall microbial growth rates. Another strategy for aeration without bubbling is the use of bubble-free membrane bioreactors, which were originally used in animal cell culture and wastewater treatment [127]. This concept was used in the past for biosurfactant production [128]. Oxygen transfer is realized by diffusion through a hollow fiber membrane into the surrounding liquid without bubble formation [129]. A different example of a bubble-free bioreactor is described by Pinzon et al., where cells were immobilized outside the fibers and the medium circulated inside the fibers [130]. Using this concept, higher microbial growth as well as biosurfactant production was reported compared to initial work with membrane bioreactors. Bongartz et al. reported on the design of a bubble-free aeration system by combining simulations and experiments of a submerged membrane module for efficient bubble-free aeration [47]. A digital twin of the combined bioreactor and membrane aeration module was used to optimize mixing in computational fluid dynamics studies.

4.2.4 Considerations for future bioeconomical-technical substrates In a future bio-based economy, substrates, which are not in competition with food are preferred, the most important source being lignocellulosic biomass or lignocellulose-derived side-streams [131]. This transition toward nonfood alternative substrates is especially relevant for biosurfactants since chemically synthesized surfactants are a mass product of the chemical industry and thus a prime reference product for establishing sustainable alternative processes. While there are different potentially conceivable concepts for utilization of lignocellulosic biomass using prior depolymerization and monomerization of the contained sugars, there are also approaches which rely on using mechanically broken-down whole biomass. It is for this reason that solid state fermentation may play and increasingly important role for RL production in the future, e.g., in concepts that are referred to as simultaneous saccharification and fermentation (SSF) or consolidated bioprocessing (CBP).

4.3 Bioreactor-coupled integrated downstream processing One of the most common approaches used for the purification of biosurfactants is precipitation followed by extraction with organic solvent. Demling et al. developed a fermentation concept with integrated in situ product removal (ISPR) by extraction for RL production using recombinant P. putida [97]. Even though it is possible to perform this as a dedicated continuous process with recycling of the applied solvent, the generation of problematic waste stream cannot completely be avoided. As an economical and environmentally friendly approach, the physical properties, notably the abilities to form micelles and accumulate in the foam fraction, can be used as a basis to develop a purification concept, which can ideally be integrated into existing approaches.

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4.3.1 Foam fractionation concepts Foam fractionation is based on the physicochemical properties of biosurfactants. Due to their surface activity, biosurfactants adsorb on the surface of air bubbles. Adsorption depends on various parameters, such as pH and ionic strength of the solution [132]. During foam fractionation, the resulting foam is collected in an external foam collector where it is collapsed typically either due to time, shear stress, use of acid, or combinations thereof. In addition to using the gas outlet of a bioreactor, specialized devices for foam fractionation such as fractionation columns have been reported [133,134]. In foam fractionation, many different parameters have complex and opposing effects on enrichment and recovery. Therefore, settings need to be adjusted to the specific application. For this purpose, empiric process design is common due to the complexity of interactions regarding relevant parameters. To prevent the enrichment of cells in foam, Heyd et al. reported the immobilization of P. aeruginosa on magnetic particles, resulting in strong cell retention [135]. Another possibility to reduce the adhesion of cells on the foam is the modification of the cell surface as described for P. putida [38]. Based on previous findings [98], Anic et al. designed a foam fractionation procedure for purification of RL from a fermentation broth [44]. Blesken et al. reported a process for continuous RL separation by using an external fractionation column to decouple biosurfactant production from foam fractionation [91]. In a subsequent product recovery step, continuous foam adsorption was integrated into the process.

4.3.2 Purification based on micelle-forming properties The unique property of surfactant molecules is that they tend to form supramolecular structures above the CMC, as was described in Chapter 1. These supramolecular structures such as micelles or vesicles display nominal molecular diameters up to two to three orders of magnitude larger than those of individual unassociated molecules. It was demonstrated in the past that surfactant molecules in this form can be retained applying ultrafiltration membranes using appropriate and specific molecular weight cut-offs. Mulligan et al. successfully used this principle for the recovery and purification of RL from complex fermentation broths [136]. Another method described by Lin and Jiang [137] focuses on the recovery and purification of biosurfactants from fermentation broths and consists of two coupled ultrafiltration steps. In the first step, biosurfactants are effectively retained by membranes above their CMC, while subsequently, the micelles of the retentate are broken by the addition of a chemical, usually a 50% (v/v) alcohol solution (e.g., methanol), to allow recovery in the permeate. Due to the characteristic physico-chemical properties of biosurfactant molecules, a bioprocess for RL production is accompanied by special challenges, for the native producer P. aeruginosa, as well as for the heterologous producer P. putida. Several of those challenges can be counteracted or even overcome by appropriate reactor and/or process design. A combination of successful approaches presented here might result in an economically competitive large-scale production of biosurfactants.

5. Concluding remarks Establishing RL synthesis in recombinant P. putida enabled the biotechnological community to apply existing and newly developed tools to enhance the productivity of the microbial cell factory. These methods for efficient gene expression as well as metabolic streamlining can be combined for enhancing yield and rate while process design including sophisticated cultivations strategies facilitates high titers. The efforts have in part been carried out in parallel in the academic and the industrial world and

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culminated finally in the construction of a pilot RL production plant in Slovenska´ Lˇupca in Slovakia by Evonik Industries AG, which was announced in 2016 [138]. While not much is known about strain and process engineering work carried out in industry, the rich collection of academic literature indicates several contact points between synthetic biology, metabolic engineering, and process development that call for multidisciplinary actions to tackle current limitations in P. putida-based rhamnolipid production. This collection may serve as an appeal to pursue such integrated approaches, of which a few already exist. The combination of an efficient overall process with the manifold developments to establish usage of alternative carbon sources by P. putida constitutes a promising step within the transition toward a bioeconomy. The here outlined workflow may serve as a general blueprint for recombinant biosurfactant production.

Acknowledgments S.K., I.B., S.T., and T.T. received funding from the Ministry of Culture and Science of the German State of North Rhine-Westphalia within in the framework of the NRW Strategieprojekt BioSC (No. 313/323-400-00213) in the projects Bio2, DesignR and SurfIn. The work of M.F., L.M.B., and T.T. is partially funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) under Germany’s Excellence Strategy— Exzellenzcluster 2186 “The Fuel Science Center” ID: 390919832. The scientific activities of S.K., L.M.B., S.T., and T.T. were financially supported by the Federal Ministry of Education and Research in the project GlycoX (Grant number 031B0866A). S.K. and S.T. received additional funding by the Federal Ministry of Education and Research in the project NO-STRESS (Grant number 031B0852B).

References [1] Abdel-Mawgoud AM, Hausmann R, Lepine F, M€ uller MM, Deziel E. Rhamnolipids: detection, analysis, biosynthesis, genetic regulation, and bioengineering of production. In: Biosurfactants. Springer; 2011. p. 13–55. [2] Costa SG, Deziel E, Lepine F. Characterization of rhamnolipid production by Burkholderia glumae. Lett Appl Microbiol 2011;53(6):620–7. [3] Rezanka T, Siristova L, Sigler K. Rhamnolipid-producing thermophilic bacteria of species Thermus and Meiothermus. Extremophiles 2011;15(6):697–709. [4] Tripathi L, Twigg MS, Zompra A, Salek K, Irorere VU, Gutierrez T, et al. Biosynthesis of rhamnolipid by a Marinobacter species expands the paradigm of biosurfactant synthesis to a new genus of the marine microflora. Microb Cell Fact 2019;18(1):164. [5] Cabrera-Valladares N, Richardson AP, Olvera C, Trevin˜o LG, Deziel E, Lepine F, et al. Monorhamnolipids and 3-(3-hydroxyalkanoyloxy)alkanoic acids (HAAs) production using Escherichia coli as a heterologous host. Appl Microbiol Biotechnol 2006;73(1):187–94. [6] Bahia FM, de Almeida GC, de Andrade LP, Campos CG, Queiroz LR, da Silva RLV, et al. Rhamnolipids production from sucrose by engineered Saccharomyces cerevisiae. Sci Rep 2018;8(1):2905. [7] Horlamus F, Wittgens A, Noll P, Michler J, M€uller I, Weggenmann F, et al. One-step bioconversion of hemicellulose polymers to rhamnolipids with Cellvibrio japonicus: a proof-of-concept for a potential host strain in future bioeconomy. GCB Bioenergy 2019;11(1):260–8. [8] Ochsner UA, Reiser J, Fiechter A, Witholt B. Production of Pseudomonas aeruginosa Rhamnolipid biosurfactants in heterologous hosts. Appl Environ Microbiol 1995;61(9):3503–6.

References

175

[9] Arnold S, Henkel M, Wanger J, Wittgens A, Rosenau F, Hausmann R. Heterologous rhamnolipid biosynthesis by P. putida KT2440 on bio-oil derived small organic acids and fractions. AMB Express 2019;9 (1):80. [10] Bator I, Wittgens A, Rosenau F, Tiso T, Blank LM. Comparison of three xylose pathways in Pseudomonas putida KT2440 for the synthesis of valuable products. Front Bioeng Biotechnol 2020;7(480). [11] Beuker J, Steier A, Wittgens A, Rosenau F, Henkel M, Hausmann R. Integrated foam fractionation for heterologous rhamnolipid production with recombinant Pseudomonas putida in a bioreactor. AMB Express 2016;6(1):1–10. [12] Beuker J, Barth T, Steier A, Wittgens A, Rosenau F, Henkel M, et al. High titer heterologous rhamnolipid production. AMB Express 2016;6(1):1–7. [13] Noll P, Treinen C, M€uller S, Senkalla S, Lilge L, Hausmann R, et al. Evaluating temperature-induced regulation of a ROSE-like RNA-thermometer for heterologous rhamnolipid production in Pseudomonas putida KT2440. AMB Express 2019;9(1):154. [14] Tiso T, Sabelhaus P, Behrens B, Wittgens A, Rosenau F, Hayen H, et al. Creating metabolic demand as an engineering strategy in Pseudomonas putida—Rhamnolipid synthesis as an example. Metab Eng Commun 2016;3:234–44. [15] Tiso T, Zauter R, Tulke H, Leuchtle B, Li W-J, Behrens B, et al. Designer rhamnolipids by reduction of congener diversity: production and characterization. Microb Cell Fact 2017;16(1):1–14. [16] Tiso T, Ihling N, Kubicki S, Biselli A, Schonhoff A, Bator I, et al. Integration of genetic and process engineering for optimized Rhamnolipid production using Pseudomonas putida. Front Bioeng Biotechnol 2020;8:976. [17] Wittgens A, Tiso T, Arndt TT, Wenk P, Hemmerich J, M€ uller C, et al. Growth independent rhamnolipid production from glucose using the nonpathogenic Pseudomonas putida KT2440. Microb Cell Fact 2011;10(1):80. [18] Wittgens A, Kovacic F, M€uller MM, Gerlitzki M, Santiago-Sch€ ubel B, Hofmann D, et al. Novel insights into biosynthesis and uptake of rhamnolipids and their precursors. Appl Microbiol Biotechnol 2017;101 (7):2865–78. [19] Wittgens A, Santiago-Schuebel B, Henkel M, Tiso T, Blank LM, Hausmann R, et al. Heterologous production of long-chain rhamnolipids from Burkholderia glumae in Pseudomonas putida—a step forward to tailor-made rhamnolipids. Appl Microbiol Biotechnol 2018;102(3):1229–39. [20] Loeschcke A, Thies S. Engineering of natural product biosynthesis in Pseudomonas putida. Curr Opin Biotechnol 2020;65:213–24. [21] Sobero´n-Cha´vez G, Gonza´lez-Valdez A, Soto-Aceves MP, Cocotl-Yan˜ez M. Rhamnolipids produced by Pseudomonas: from molecular genetics to the market. J Microbial Biotechnol 2021;14(1):136–46. [22] Tiso T, Thies S, M€uller M, Tsvetanova L, Carraresi L, Br€ oring S, et al. Rhamnolipids—production, performance, and application. In: Lee SY, editor. Consequences of microbial interactions with hydrocarbons, oils and lipids: production of fuels and chemicals. Handbook of hydrocarbon and lipid microbiology series. Berlin: Springer International Publishing AG; 2017. [23] Abdel-Mawgoud AM, Lepine F, Deziel E. Rhamnolipids: diversity of structures, microbial origins and roles. Appl Microbiol Biotechnol 2010;86(5):1323–36. [24] Germer A, Tiso T, M€uller C, Behrens B, Vosse C, Scholz K, et al. Exploiting the natural diversity of RhlA acyltransferases for the synthesis of the rhamnolipid precursor 3-(3-hydroxyalkanoyloxy) alkanoic acid. Appl Environ Microbiol 2020;86(6):e02317–9. [25] Martin-Pascual M, Batianis C, Bruinsma L, Asin-Garcia E, Garcia-Morales L, Weusthuis RA, et al. A navigation guide of synthetic biology tools for Pseudomonas putida. Biotechnol Adv 2021;107732. [26] Apura P, Saramago M, Peregrina A, Viegas SC, Carvalho SM, Saraiva LM, et al. Tailor-made sRNAs: a plasmid tool to control the expression of target mRNAs in Pseudomonas putida. Plasmid 2020;109, 102503.

176

Chapter 8 Metabolic and process engineering on the edge

[27] Neves D, Vos S, Blank LM, Ebert BE. Pseudomonas mRNA 2.0: boosting gene expression through enhanced mRNA stability and translational efficiency. Front Bioeng Biotechnol 2020;7:458. [28] Kumar R, Das AJ. Advancement of genetic engineering in Rhamnolipid (s) production. In: Rhamnolipid Biosurfactant. Springer; 2018. p. 43–50. [29] Wittgens A, Rosenau F. Heterologous rhamnolipid biosynthesis: advantages, challenges, and the opportunity to produce tailor-made rhamnolipids. Front Bioeng Biotechnol 2020;8(1263). [30] Dubeau D, Deziel E, Woods DE, Lepine F. Burkholderia thailandensis harbors two identical rhl gene clusters responsible for the biosynthesis of rhamnolipids. BMC Microbiol 2009;9:263. [31] Gehring C, Wessel M, Schaffer S, Thum O. The power of biocatalysis: a one-pot total synthesis of rhamnolipids from butane as the sole carbon and energy source. ChemistryOpen 2016;5(6):513. [32] Du J, Zhang A, Ja H, Wang J. Biosynthesis of di-rhamnolipids and variations of congeners composition in genetically-engineered Escherichia coli. Biotechnol Lett 2017;39(7):1041–8. [33] Dulcey CE, Lo´pez de los Santos Y, Letourneau M, Deziel E, Doucet N. Semi-rational evolution of the 3-(3hydroxyalkanoyloxy)alkanoate (HAA) synthase RhlA to improve rhamnolipid production in Pseudomonas aeruginosa and Burkholderia glumae. FEBS J 2019;286(20):4036–59. [34] Han L, Liu P, Peng Y, Lin J, Wang Q, Ma Y. Engineering the biosynthesis of novel rhamnolipids in Escherichia coli for enhanced oil recovery. J Appl Microbiol 2014;117(1):139–50. [35] Cao L, Wang Q, Zhang J, Li C, Yan X, Lou X, et al. Construction of a stable genetically engineered rhamnolipid-producing microorganism for remediation of pyrene-contaminated soil. World J Microbiol Biotechnol 2012;28(9):2783–90. [36] Cha M, Lee N, Kim M, Kim M, Lee S. Heterologous production of Pseudomonas aeruginosa EMS1 biosurfactant in Pseudomonas putida. Bioresour Technol 2008;99(7):2192–9. [37] Bator I, Karmainski T, Tiso T, Blank LM. Killing two birds with one stone – strain engineering facilitates the development of a unique Rhamnolipid production process. Front Bioeng Biotechnol 2020;8(899). [38] Blesken CC, Bator I, Eberlein C, Heipieper HJ, Tiso T, Blank LM. Genetic cell-surface modification for optimized foam fractionation. Front Bioeng Biotechnol 2020;8(1116). [39] Askitosari TD, Berger C, Tiso T, Harnisch F, Blank LM, Rosenbaum MA. Coupling an electroactive Pseudomonas putida KT2440 with bioelectrochemical Rhamnolipid production. Microorganisms 2020;8(12):1959. [40] Noll P, Treinen C, M€uller S, Lilge L, Hausmann R, Henkel M. Exploiting RNA thermometer-driven molecular bioprocess control as a concept for heterologous rhamnolipid production. Sci Rep 2021;11(1):14802. [41] Setoodeh P, Jahanmiri A, Eslamloueyan R, Niazi A, Ayatollahi SS, Aram F, et al. Statistical screening of medium components for recombinant production of Pseudomonas aeruginosa ATCC 9027 rhamnolipids by nonpathogenic cell factory Pseudomonas putida KT2440. Mol Biotechnol 2014;56(2):175–91. [42] Behrens B, Baune M, Jungkeit J, Tiso T, Blank LM, Hayen H. High performance liquid chromatographycharged aerosol detection applying an inverse gradient for quantification of rhamnolipid biosurfactants. J Chromatogr A 2016;1455:125–32. [43] Behrens B, Engelen J, Tiso T, Blank LM, Hayen H. Characterization of rhamnolipids by liquid chromatography/mass spectrometry after solid-phase extraction. Anal Bioanal Chem 2016;408(10):2505–14. [44] Anic I, Nath A, Franco P, Wichmann R. Foam adsorption as an ex situ capture step for surfactants produced by fermentation. J Biotechnol 2017;258:181–9. [45] Anic I, Apolonia I, Franco P, Wichmann R. Production of rhamnolipids by integrated foam adsorption in a bioreactor system. AMB Express 2018;8(1):122. [46] Utomo RNC, Li W-J, Tiso T, Eberlein C, Doeker M, Heipieper HJ, et al. Defined microbial mixed culture for utilization of polyurethane monomers. ACS Sustain Chem Eng 2020;8(47):17466–74. [47] Bongartz P, Bator I, Baitalow K, Keller R, Tiso T, Blank LM, et al. A scalable bubble-free membrane aerator for biosurfactant production. Biotechnol Bioeng 2021;118(9):3545–58. [48] Wigneswaran V, Nielsen KF, Sternberg C, Jensen PR, Folkesson A, Jelsbak L. Biofilm as a production platform for heterologous production of rhamnolipids by the nonpathogenic strain Pseudomonas putida KT2440. Microb Cell Fact 2016;15(1):1–13.

References

177

[49] Reis RS, Pereira AG, Neves BC, Freire DMG. Gene regulation of rhamnolipid production in Pseudomonas aeruginosa - a review. Bioresour Technol 2011;102(11):6377–84. [50] Sathesh-Prabu C, Tiwari R, Kim D, Lee SK. Inducible and tunable gene expression systems for Pseudomonas putida KT2440. Sci Rep 2021;11(1):1–8. [51] Schuster LA, Reisch CR. A plasmid toolbox for controlled gene expression across the Proteobacteria. Nucleic Acids Res 2021. [52] K€obbing S, Blank LM, Wierckx N. Characterization of context-dependent effects on synthetic promoters. Front Bioeng Biotechnol 2020;8:551. [53] Tietze L, Lale R. Importance of the 50 regulatory region to bacterial synthetic biology applications. J Microbial Biotechnol 2021;14(6):2291–315. [54] Komarova ES, Chervontseva ZS, Osterman IA, Evfratov SA, Rubtsova MP, Zatsepin TS, et al. Influence of the spacer region between the Shine–Dalgarno box and the start codon for fine-tuning of the translation efficiency in Escherichia coli. J Microbial Biotechnol 2020;13(4):1254–61. [55] Volkenborn K, Kuschmierz L, Benz N, Lenz P, Knapp A, Jaeger K-E. The length of ribosomal binding site spacer sequence controls the production yield for intracellular and secreted proteins by Bacillus subtilis. Microb Cell Fact 2020;19(1):1–12. [56] Apura P, Gonc¸alves LG, Viegas SC, Arraiano CM. The world of ribonucleases from pseudomonads: a short trip through the main features and singularities. J Microbial Biotechnol 2021. [57] Hui MP, Foley PL, Belasco JG. Messenger RNA degradation in bacterial cells. Annu Rev Genet 2014;48:537–59. [58] Mutalik VK, Guimaraes JC, Cambray G, Lam C, Christoffersen MJ, Mai Q-A, et al. Precise and reliable gene expression via standard transcription and translation initiation elements. Nat Methods 2013;10 (4):354–60. [59] Zobel S, Benedetti I, Eisenbach L, de Lorenzo V, Wierckx N, Blank LM. Tn7-based device for calibrated heterologous gene expression in Pseudomonas putida. ACS Synth Biol 2015;4(12):1341–51. [60] Senoussi A, Lee Tin Wah J, Shimizu Y, Robert J, Jaramillo A, Findeiss S, et al. Quantitative characterization of translational riboregulators using an in vitro transcription–translation system. ACS Synth Biol 2018;7(5):1269–78. [61] Calero P, Volke DC, Lowe PT, Gotfredsen CH, O’Hagan D, Nikel PI. A fluoride-responsive genetic circuit enables in vivo biofluorination in engineered Pseudomonas putida. Nat Commun 2020;11 (1):1–11. [62] Otto M, Wynands B, Drepper T, Jaeger K-E, Thies S, Loeschcke A, et al. Targeting 16S rDNA for stable recombinant gene expression in Pseudomonas. ACS Synth Biol 2019;8(8):1901–12. [63] Gyorgy A, Jimenez JI, Yazbek J, Huang H-H, Chung H, Weiss R, et al. Isocost lines describe the cellular economy of genetic circuits. Biophys J 2015;109(3):639–46. [64] Kelly JR, Rubin AJ, Davis JH, Ajo-Franklin CM, Cumbers J, Czar MJ, et al. Measuring the activity of BioBrick promoters using an in vivo reference standard. J Biol Eng 2009;3(1):1–13. [65] Calero P, Jensen SI, Nielsen AT. Broad-host-range ProUSER vectors enable fast characterization of inducible promoters and optimization of p-coumaric acid production in Pseudomonas putida KT2440. ACS Synth Biol 2016;5(7):741–53. [66] Wang G, Zhao Z, Ke J, Engel Y, Shi Y-M, Robinson D, et al. CRAGE enables rapid activation of biosynthetic gene clusters in undomesticated bacteria. Nat Microbiol 2019;4(12):2498–510. [67] Zhang JJ, Tang X, Huan T, Ross AC, Moore BS. Pass-back chain extension expands multimodular assembly line biosynthesis. Nat Chem Biol 2020;16(1):42–9. [68] Henkel M, M€uller MM, K€ugler JH, Lovaglio RB, Contiero J, Syldatk C, et al. Rhamnolipids as biosurfactants from renewable resources: concepts for next-generation rhamnolipid production. Process Biochem 2012;47(8):1207–19. [69] Yang S-T. Bioprocessing for value-added products from renewable resources: new technologies and applications. Elsevier; 2011.

178

Chapter 8 Metabolic and process engineering on the edge

[70] Lei L, Zhao F, Han S, Zhang Y. Enhanced rhamnolipids production in Pseudomonas aeruginosa SG by selectively blocking metabolic bypasses of glycosyl and fatty acid precursors. Biotechnol Lett 2020;42 (6):997–1002. [71] Tiso T, Wichmann R, Blank LM. Accessing natural diversity of rhamnolipids by metabolic engineering of Pseudomonas putida. In: Fachgruppe Biologie; 2016. [72] Klinke S, Dauner M, Scott G, Kessler B, Witholt B. Inactivation of isocitrate lyase leads to increased production of medium-chain-length poly(3-hydroxyalkanoates) in Pseudomonas putida. Appl Environ Microbiol 2000;66(3):909–13. [73] Miller DJ, Zhang YM, Rock CO, White SW. Structure of RhlG, an essential beta-ketoacyl reductase in the rhamnolipid biosynthetic pathway of Pseudomonas aeruginosa. J Biol Chem 2006;281(26):18025–32. [74] Davis MS, Solbiati J, Cronan JE. Overproduction of acetyl-CoA carboxylase activity increases the rate of fatty acid biosynthesis in Escherichia coli. J Biol Chem 2000;275(37):28593–8. [75] Abdel-Mawgoud Ahmad M, Lepine F, Deziel E. A stereospecific pathway diverts β-oxidation intermediates to the biosynthesis of Rhamnolipid biosurfactants. Chem Biol 2014;21(1):156–64. [76] Gutierrez-Go´mez U, Soto-Aceves MP, Servı´n-Gonza´lez L, Sobero´n-Cha´vez G. Overproduction of rhamnolipids in Pseudomonas aeruginosa PA14 by redirection of the carbon flux from polyhydroxyalkanoate synthesis and overexpression of the rhlAB-R operon. Biotechnol Lett 2018;40(11
12):1561–6. [77] Thum O, Engel P, Gehring C, Schaffer S, Wessel M, inventors; Evonik Degussa GmbH, assignee. Methods of producing rhamnolipids. United Stated patent 10174353; 2019. [78] Borrero-de Acun˜a JM, Bielecka A, H€aussler S, Schobert M, Jahn M, Wittmann C, et al. Production of medium chain length polyhydroxyalkanoate in metabolic flux optimized Pseudomonas putida. Microb Cell Fact 2014;13(1):88. [79] Poblete-Castro I, Binger D, Rodrigues A, Becker J, Martins dos Santos VAP, Wittmann C. In-silico-driven metabolic engineering of Pseudomonas putida for enhanced production of poly-hydroxyalkanoates. Metab Eng 2013;15:113–23. [80] Martı´nez-Garcı´a E, Nikel PI, Chavarrı´a M, de Lorenzo V. The metabolic cost of flagellar motion in Pseudomonas putida KT2440. Environ Microbiol 2014;16(1):291–303. [81] Wang Y, Horlamus F, Henkel M, Kovacic F, Schl€afle S, Hausmann R, et al. Growth of engineered Pseudomonas putida KT2440 on glucose, xylose, and arabinose: hemicellulose hydrolysates and their major sugars as sustainable carbon sources. GCB Bioenergy 2019;11(1):249–59. [82] Tiso T, Narancic T, Wei R, Pollet E, Beagan N, Schr€ oder K, et al. Toward bio-upcycling of polyethylene terephthalate. Metab Eng 2021;66:167–78. [83] Ackermann YS, Li W-J, de Hipt LO, Niehoff P-J, Casey W, Polen T, et al. Engineering adipic acid metabolism in Pseudomonas putida. Metab Eng 2021;67:29–40. [84] Franden MA, Jayakody LN, Li W-J, Wagner NJ, Cleveland NS, Michener WE, et al. Engineering Pseudomonas putida KT2440 for efficient ethylene glycol utilization. Metab Eng 2018;48:197–207. [85] Li WJ, Jayakody LN, Franden MA, Wehrmann M, Daun T, Hauer B, et al. Laboratory evolution reveals the metabolic and regulatory basis of ethylene glycol metabolism by Pseudomonas putida KT2440. Environ Microbiol 2019;21(10):3669–82. [86] L€owe H, Schmauder L, Hobmeier K, Kremling A, Pfl€ uger-Grau K. Metabolic engineering to expand the substrate spectrum of Pseudomonas putida toward sucrose. MicrobiologyOpen 2017;6(4). [87] Koch AK, Reiser J, K€appeli O, Fiechter A. Genetic construction of lactose-utilizing strains of Pseudomonas aeruginosa and their application in biosurfactant production. Bio/Technology 1988;6(11):1335–9. [88] Nikel PI, de Lorenzo V. Pseudomonas putida as a functional chassis for industrial biocatalysis: from native biochemistry to trans-metabolism. Metab Eng 2018;50:142–55. [89] Linger JG, Hobdey SE, Franden MA, Fulk EM, Beckham GT. Conversion of levoglucosan and cellobiosan by Pseudomonas putida KT2440. Metab Eng Commun 2016;3:24–9.

References

179

[90] Yang S, Li S, Jia X. Production of medium chain length polyhydroxyalkanoate from acetate by engineered Pseudomonas putida KT2440. J Ind Microbiol Biotechnol 2019;46(6):793–800. [91] Blesken CC, Str€umpfler T, Tiso T, Blank LM. Uncoupling foam fractionation and foam adsorption for enhanced biosurfactant synthesis and recovery. Microorganisms 2020;8(12):2029. [92] Sarachat T, Pornsunthorntawee O, Chavadej S, Rujiravanit R. Purification and concentration of a rhamnolipid biosurfactant produced by Pseudomonas aeruginosa SP4 using foam fractionation. Bioresour Technol 2010;101(1):324–30. [93] Claus S, Sa´nchez LJ, Van Bogaert INA. The role of transport proteins in the production of microbial glycolipid biosurfactants. Appl Microbiol Biotechnol 2021;1-15. [94] Wilhelm S, Gdynia A, Tielen P, Rosenau F, Jaeger K-E. The autotransporter esterase EstA of Pseudomonas aeruginosa is required for rhamnolipid production, cell motility, and biofilm formation. J Bacteriol 2007;189(18):6695–703. [95] Lesˇcic Asˇler I, Ivic N, Kovacic F, Schell S, Knorr J, Krauss U, et al. Probing enzyme promiscuity of SGNH hydrolases. Chembiochem 2010;11(15):2158–67. [96] Beuker J, Syldatk C, Hausmann R. Bioreactors for the production of biosurfactants. In: Biosurfactants: production and utilization; processes, technologies, and economics; 2014. p. 117–28. [97] Demling P, von Campenhausen M, Gr€utering C, Tiso T, Jupke A, Blank LM. Selection of a recyclable in situ liquid–liquid extraction solvent for foam-free synthesis of rhamnolipids in a two-phase fermentation. Green Chem 2020;22(23):8495–510. [98] Siemann-Herzberg M, Wagner F. Prospects and limits for the production of biosurfactants using immobilized biocatalysts. In: Kosaric N, editor. Biosurfactants. Surfactant science series. 48. New York: Marcel Dekker Inc.; 1993. [99] Furchner B, Mersmann A. Foam breaking by high speed rotors. Chem Eng Technol 1990;13(1):86–96. [100] Chtioui O, Dimitrov K, Gancel F, Dhulster P, Nikov I. Rotating discs bioreactor, a new tool for lipopeptides production. Process Biochem 2012;47(12):2020–4. [101] Davis D, Lynch H, Varley J. The production of surfactin in batch culture by Bacillus subtilis ATCC 21332 is strongly influenced by the conditions of nitrogen metabolism. Enzyme Microb Technol 1999;25 (3–5):322–9. [102] M€uller MM, H€ormann B, Syldatk C, Hausmann R. Pseudomonas aeruginosa PAO1 as a model for rhamnolipid production in bioreactor systems. Appl Microbiol Biotechnol 2010;87(1):167–74. [103] Jiang J, Zhang D, Niu J, Jin M, Long X. Extremely high-performance production of rhamnolipids by advanced sequential fed-batch fermentation with high cell density. J Clean Prod 2021;326, 129382. [104] Pornsunthorntawee O, Maksung S, Huayyai O, Rujiravanit R, Chavadej S. Biosurfactant production by Pseudomonas aeruginosa SP4 using sequencing batch reactors: effects of oil loading rate and cycle time. Bioresour Technol 2009;100(2):812–8. [105] Zhu L, Yang X, Xue C, Chen Y, Qu L, Lu W. Enhanced rhamnolipids production by Pseudomonas aeruginosa based on a pH stage-controlled fed-batch fermentation process. Bioresour Technol 2012;117:208–13. [106] Davila A-M, Marchal R, Vandecasteele J-P. Kinetics and balance of a fermentation free from product inhibition: sophorose lipid production by Candida bombicola. Appl Microbiol Biotechnol 1992;38 (1):6–11. [107] Guerra-Santos L, K€appeli O, Fiechter A. Pseudomonas aeruginosa biosurfactant production in continuous culture with glucose as carbon source. Appl Environ Microbiol 1984;48(2):301–5. [108] Heyd M, Franzreb M, Hausmann R, Syldatk C, Berensmeier S. Integral continuous microbial rhamnolipid production. J Biotechnol 2007;2(131):S77. [109] Reiling H, Thanei-Wyss U, Guerra-Santos L, Hirt R, K€appeli O, Fiechter A. Pilot plant production of rhamnolipid biosurfactant by Pseudomonas aeruginosa. Appl Environ Microbiol 1986;51(5):985–9.

180

Chapter 8 Metabolic and process engineering on the edge

[110] Henkel M, Schmidberger A, K€uhnert C, Beuker J, Bernard T, Schwartz T, et al. Kinetic modeling of the time course of N-butyryl-homoserine lactone concentration during batch cultivations of Pseudomonas aeruginosa PAO1. Appl Microbiol Biotechnol 2013;97(17):7607–16. [111] Cameotra SS, Makkar RS. Biosurfactant-enhanced bioremediation of hydrophobic pollutants. Pure Appl Chem 2010;82(1):97–116. [112] Drouin C, Cooper D. Biosurfactants and aqueous two-phase fermentation. Biotechnol Bioeng 1992;40(1):86–90. [113] Noordman WH, Janssen DB. Rhamnolipid stimulates uptake of hydrophobic compounds by Pseudomonas aeruginosa. Appl Environ Microbiol 2002;68(9):4502–8. [114] Rau U, Nguyen L, Roeper H, Koch H, Lang S. Fed-batch bioreactor production of mannosylerythritol lipids secreted by Pseudozyma aphidis. Appl Microbiol Biotechnol 2005;68(5):607–13. [115] Vardar-Sukan F. Efficiency of natural oils as antifoaming agents in bioprocesses. J Chem Technol Biotechnol 1988;43(1):39–47. [116] Kanicky J, Poniatowski A, Mehta N, Shah D. Cooperativity among molecules at interfaces in relation to various technological processes: effect of chain length on the p K a of fatty acid salt solutions. Langmuir 2000;16(1):172–7. [117] Zhang Y, M MR. Enhanced octadecane dispersion and biodegradation by a Pseudomonas rhamnolipid surfactant (biosurfactant). Appl Environ Microbiol 1992;58(10):3276–82. [118] Pelton R. A review of antifoam mechanisms in fermentation. J Ind Microbiol Biotechnol 2002;29 (4):149–54. [119] Pugh R. Foaming, foam films, antifoaming and defoaming. Adv Colloid Interface Sci 1996;64:67–142. [120] Vardar-Sukan F. Foaming: consequences, prevention and destruction. Biotechnol Adv 1998;16 (5–6):913–48. [121] Zlokarnik M. R€uhrtechnik: Theorie und Praxis. Springer; 1999. [122] Deshpande NS, Barigou M. Performance characteristics of novel mechanical foam breakers in a stirred tank reactor. J Chem Technol Biotechnol 1999;74(10):979–87. [123] Andou S, Yoshida M, Yamagiwa K, Ohkawa A. Performance characteristics of mechanical foam-breakers with rotating parts fitted to bubble column. J Chem Technol Biotechnol 1997;68(1):94–100. [124] Takesono S, Onodera M, Yoshida M, Yamagiwa K, Ohkawa A. Performance characteristics of mechanical foam-breakers fitted to a stirred-tank reactor. J Chem Technol Biotechnol 2003;78(1):48–55. [125] Gong Z, Yang G, Che C, Liu J, Si M, He Q. Foaming of rhamnolipids fermentation: impact factors and fermentation strategies. Microb Cell Fact 2021;20(1):1–12. [126] Hoeks FW, Van Wees-Tangerman C, Luyben KCA, Gasser K, Schmid S, Mommers HM. Stirring as foam disruption (SAFD) technique in fermentation processes. Can J Chem Eng 1997;75(6):1018–29. [127] Schneider M, Reymond F, Marison I, Von Stockar U. Bubble-free oxygenation by means of hydrophobic porous membranes. Enzyme Microb Technol 1995;17(9):839–47. [128] Coutte F, Lecouturier D, Yahia SA, Lecle`re V, Bechet M, Jacques P, et al. Production of surfactin and fengycin by Bacillus subtilis in a bubbleless membrane bioreactor. Appl Microbiol Biotechnol 2010;87 (2):499–507. [129] Ahmed T, Semmens MJ. Use of transverse flow hollow fibers for bubbleless membrane aeration. Water Res 1996;30(2):440–6. [130] Pinzon NM, Cook AG, Ju LK. Continuous rhamnolipid production using denitrifying Pseudomonas aeruginosa cells in hollow-fiber bioreactor. Biotechnol Prog 2013;29(2):352–8. [131] Kiefer D, Merkel M, Lilge L, Henkel M, Hausmann R. From acetate to bio-based products: underexploited potential for industrial biotechnology. Trends Biotechnol 2021;39(4):397–411. € [132] Ozdemir G, Peker S, Helvaci S. Effect of pH on the surface and interfacial behavior of rhamnolipids R1 and R2. Colloids Surf A Physicochem Eng Asp 2004;234(1–3):135–43.

References

181

[133] Davis D, Lynch H, Varley J. The application of foaming for the recovery of surfactin from B. subtilis ATCC 21332 cultures. Enzyme Microb Technol 2001;28(4–5):346–54. [134] Winterburn J, Russell A, Martin P. Integrated recirculating foam fractionation for the continuous recovery of biosurfactant from fermenters. Biochem Eng J 2011;54(2):132–9. [135] Heyd M, Weigold P, Franzreb M, Berensmeier S. Influence of different magnetites on properties of magnetic Pseudomonas aeruginosa immobilizates used for biosurfactant production. Biotechnol Prog 2009;25 (6):1620–9. [136] Mulligan CN, Yong RN, Gibbs BF. On the use of biosurfactants for the removal of heavy metals from oil-contaminated soil. Environ Prog 1999;18(1):50–4. [137] Lin S-C, Jiang H-J. Recovery and purification of the lipopeptide biosurfactant of Bacillus subtilis by ultrafiltration. Biotechnol Tech 1997;11(6):413–6. [138] Evonik commercializes biosurfactants [press release]. accessed 22.11.2021 2016.

CHAPTER

Improved production of novel (bola) glycolipid biosurfactants with the yeast Starmerella bombicola through an integrative approach combining genetic engineering and multiomics analyses

9

Martijn Casteleina, Nicolas de Fooza,b, Goedele Luytena, Lisa Van Renterghema, Sven Dierickxa,b, Stijn Bovijna, Sophie Roelantsa, Lynn Vanhaeckeb, and Wim Soetaerta a

Centre for Industrial Biotechnology and Biocatalysis (InBio.be), Department of Biotechnology, Faculty of Bioscience Engineering Ghent University, Coupure Links, Ghent, Belgium, bLaboratory of Integrative Metabolomics (LIMET), Department of Translational Physiology, Infectiology and Public Health, Faculty of Veterinary Medicine Ghent University, Salisburylaan, Merelbeke, Belgium

1. Introduction Starmerella bombicola (initially referred to as Torulopsis bombicola and Candida bombicola) was first isolated from the honey of a bumblebee in 1970 [1]. It is classified as a nonpathogenic and osmotolerant yeast. The Starmerella clade is typically associated with (bumble)bees (Bombus spp.) and various flowers that are often visited by (bumble)bees [2]. A recent overview of the latter is given by de Graeve et al. [3]. S. bombicola was only the third discovered sophorolipid (SL) producing species, after Starmerella apicola and Pseudohyphozyma bogoriensis in 1961 and 1968, respectively [1,4,5]. This particular yeast strain has become the most intensively studied nowadays, due to its natural capacity to obtain high SL production titers and volumetric productivities, making it most relevant for industrial SL production [6–8], resulting in valorization of SL production by companies such as Evonik, Holiferm, Locus IP, Amphi-Star, etc. SLs are secondary metabolites, so their production is boosted after a period of exponential growth of the yeast cells, when they attain their stationary phase and nitrogen and/or phosphorous sources are depleted [9,10]. For S. bombicola, all genes encoding the enzymes involved in the intracellular steps of SL biosynthesis are present in a large subtelomeric gene cluster [11] as depicted in Fig. 1, which are clearly upregulated in the stationary phase [12]. Until today, the exact molecular Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00009-0 Copyright # 2023 Elsevier Inc. All rights reserved.

183

A

6

3

5

4

2

SBLE

UGTB1

MDR

AT

UGTA1

1 CYP52M1

Telomeric end

2.5 Mb

26.9 kb 2000

0

4000

6000

8000

10 000

bp

B Oleyl alcohol

HO O2

I II III

Fay alcohol oxidase

H2O2

O

Rapeseed oil

nucleus

NAD++ H2O

Lipase

NADH + 2H+ O

Fay acids

De novo fay acid synthesis

Oleic acid

HO NADPH + H+ + O2

1

NADP+ + H2O O

Z or Z-1 hydroxy fay acid

HO OH

UDP-glucose

2

UDP

OH

O O

HO HO

H3C O

O

OH

O

Glucolipid

O

HO HO

HO O

UDP-glucose

3

UDP

HO OH HO O

O O

OH

HO

Mono-acetylated acidic SL OH

O

HO HO

HO OH HO O

O

O

H3C

O H3C Acetyl-CoA

HO

CoA-SH

4

Non-acetylated acidic SL

ATP

OR1 O

O

O HO

O O

OH

Di-acetylated acidic SL ADP + Pi

5

O

O HO O HO O

HO HO

O HO

OR2 HO HO O

O

O

O

OH

OR1 O

O

6

OH

H2O

O

HO HO

HO OR2 HO O

O O

OH

HO

Lactonic SLs (Non-, mono- and di-acetylated)

Acidic SLs (Non-, mono- and di-acetylated)

FIG. 1 (A) Illustration of chromosome II of S. bombicola containing the full SL biosynthetic gene cluster (
11 kilobases) and the gene responsible for lactonization (SBLE) at the other side of the same chromosome. (B) The full SL biosynthetic pathway consists of (1) hydroxylation of a fatty acid by a CYP52M1 monooxygenase, (2) glucosylation of the hydroxy fatty acid by the first glucosyltransferase UGTA1, and (3) second glucosylation step of the formed glucolipid by a second glucosyltransferase UGTB1 giving rise to an acidic SL, which can be (4) acetylated by the action of an acetyltransferase AT. The different SLs are transported into the extracellular space by a multidrug transporter protein MDR (5). Lactonization (6) occurs extracellularly as the responsible enzyme SBLE is secreted. Adapted from Roelants SLKW, Ciesielska K, de Maeseneire SL, Moens H, Everaert B, Verweire S, et al. Towards the industrialization of new biosurfactants: Biotechnological opportunities for the lactone esterase gene from Starmerella bombicola. Biotechnol Bioeng 2016;113(3):550–9. https://doi.org/10.1002/BIT.25815.

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basis of the regulation of SL biosynthesis has not yet been elucidated. An analogy with other secondary metabolite production is anticipated [13] and is one of the research focuses of our lab. During the past decade, we elucidated the SL biosynthesis pathway for S. bombicola (Fig. 1) [11,14–18]. SLs produced by wild-type S. bombicola typically contain a hydrophobic hydroxy fatty acid part that is ω-1/ω (subterminal/terminal) hydroxystearate (C18:0) or hydroxyl-oleate (C18:1), and hydroxypalmitate (C16:0) and hydroxypalmitoleate (C16:1) moieties are detected [5]. The specificity for the fatty acid moiety is determined by the specificity of the CYP52M1 cytochrome P450 monooxygenase enzyme, responsible for the first step of the production pathway, which selectively hydroxylates the fatty acid chain on the subterminal or terminal position (ω-1/ω) [18]. Subsequently, glycosylation of the hydroxy fatty acid is performed by the glucosyltransferase UGTA1 to form a glucolipid (GL, and further glycosylation to an acidic SL by adding another glucose molecule by the action of a second glucosyltransferase UGTB1 [16,17]. Next, acetyl groups can be added at the 60 and 600 positions of the sophorose moiety by an acetyltransferase AT [15]. The acidic SLs are then transported out of the cell by the ATP-consuming MDR transporter [11]. Recently, a second SL transporter MDR2 was discovered [19] by analyzing the complete transportome (the collection of all transporter proteins of an organism) of S. bombicola. The MDR2 gene is located outside of the SL biosynthetic gene cluster and thus not subject to its regulation. In the extracellular space, the lactone esterase SBLE enzyme can form a covalent bond between the 400 hydroxyl group of the second glucose molecule and the free carboxylic acid group of the acidic SL, giving rise to a macrocyclic lactone structure [20,21]. The SLs produced by S. bombicola mainly consist of di-acetylated lactonic SLs and minor amounts of acidic SLs, although the ratio can be influenced by changing the culturing conditions, the substrate used, culture medium, etc. A lot of different hydrophobic substrates are reported for SL production using S. bombicola: alkanes, fatty acids, fatty esters, plant-based oils, and so on [22–25], and an overview is provided in a recent book chapter [26]. Mostly, C16–C18 chain substrates were assessed, as this is the preferred chain length of the CYP52M1 enzyme (Fig. 1) [27]. Additionally, shorter or special unconventional substrates were already investigated in the past, giving rise to SLs based on petroselinic acid (C18:1 (ω-12)), coconut oil (C12:0), meadowfoam seed oil (mainly C20:1 (ω-7)), eicosapentaenoic acid (EPA, 20:5 (ω-3)), and docosahexaenoic acid (DHA, 22:6 (ω-3)) [28–30]. A strategy to bypass the CYP52M1 enzyme is to feed S. bombicola with already hydroxylated substrates, such as medium-chain alcohols or diols (1- or 2-dodecanol, 1,12-dodecanediol, 1-tetradecanol, or myristyl alcohol) [25,31–33]. Even though the SL titers from feeding with these hydroxylated substrates are quite low, some of these authors detected novel glycolipids and some have very promising antimicrobial potential [34,35]. More information about S. bombicola fermentation parameters and development has been thoroughly discussed by Roelants et al. [36].

2. Diversifying and boosting glycolipid production with S. bombicola Genetic engineering has definitely made its entry to alter glycolipid synthesis with S. bombicola over the past decade. Our lab had and has a pioneering role in the domestication of S. bombicola. Genetic engineering methods have been developed since 2007 at our lab, and we have significantly contributed to this field since. This work has resulted in the full elucidation of the SL biosynthetic pathway at our lab and enabled the bending of the well-oiled SL machinery toward the production of tailored glycolipids. All this would not have been possible without the development of a molecular toolbox for

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Chapter 9 Improved production of novel (bola) glycolipid biosurfactants

S. bombicola. This toolbox and the general principles for genetic engineering of S. bombicola are thoroughly described by [37], to which the interested reader is referred. Moreover, CRISPR (clustered regulatory interspaced short palindromic repeats) technology for S. bombicola genetic engineering has recently been developed at our lab (Luyten et al., unpublished results). The development of two other CRISPR-based genetic engineering systems has been published recently, one employing the Cas9 gene [38], while the other makes use of the activity of Cas12a [39]. Taken together, S. bombicola is not limited to classic SL production anymore but should be considered as a platform organism for the production of novel glycolipids by employing (part of) the SL biosynthetic enzymes (Fig. 1). An overview of the modified strains and their main glycolipid end products that will be discussed in the remainder of this section is represented in Fig. 2 and Table 1.

FIG. 2 Novel S. bombicola strains developed at InBio.be and the main glycolipid product produced by these strains as also summarized in Table 1. By-product formation is described in the respective papers describing these novel strains. Rx shows the location of possible acetylation (CH3dCOd). SL, sophorolipid; SS: sophoroside; 20-HETE, 20-hydroxyeicosatetraenoic acid based; GL, glucolipid; GS, glucoside.

Table 1 Overview of engineered S. bombicola strains at Inbio.be and their produced glycolipids.a Strain

Substrates

Produced glycolipid

FT (scale)

Titer (g/L)

Prod (g/L h)

YC (g/g)

Reference

Δmfe-2

Lauryl alcohol, glucose 20-HETE, glucose Rapeseed oil, glucose Rapeseed oil, glucose Rapeseed oil, glucose Rapeseed oil, glucose Oleic acid, glucose Rapeseed oil, glucose Ethyl palmitate (C16:0), glucose Petroselinic acid, glucose Oleic acid, glucose Rapeseed oil, glucose HOSO, glucose

C12:0-based acidic SL

B (SF)

29

0.12

0.21

[40]

20-HETE-based SL

B (SF)

19

0.11

0.16

[41]

Hydroxy fatty acids

B (SF)

n.a.

n.a.

n.a.

[17]

Acetylated GLs

B (SF)

4

0.11

n.a.

[16]

Cellobiose lipids

B (SF)

0.5-1

n.a.

n.a.

[13]

Nonacetylated SLs

B (SF)

5

0.01

0.03

[15]

No SL production

B (SF)

n.a.

n.a.

n.a.

[18]

Polyhydroxyalkanoate (PHA) C16:0 SLs

B (SF)

0.6

n.a.

n.a.

[13]

FB (30 L)

12

0.02

n.a.

[42]

Petroselinic-based acidic SLs

FB (3 L)

40

0.18

0.19

[28]

Acetylated lactonic SLs Acetylated acidic SLs

FB (150 L) FB (150 L) FB (150 L)

199

0.9

0.70

[43]

138

0.83

n.a.

[43]

63

0.22

0.19

[44]

CFb (10 L)

120

0.63

n.a.

[26]

B (SF)

20

0.07

n.a.

[44]

B (SF)

18

0.09

n.a.

(unpublished results)

Δugta1 Δugtb1 Δugtb1:: UGT1 Δat Δcyp52m1 Δcyp52m1:: phaC1 Δcyp52m1:: UmCYP1

OE SBLE

Δsble Δat Δsble

Oleic acid, glucose Δat Δsble Δfao1

Δat Δsble Δfao1 Δugtb1

Palmityl (C16:0) or oleyl alcohol (C18:1), glucose Oleyl alcohol (C18:1), glucose

Bola SLs and nonacetylated acidic SLs Bola SLs and nonacetylated acidic SLs Bola SSs and alkyl SSs

Bola GSs and alkyl GSs

a FT ¼ fermentation technique, WT ¼ wild type, B ¼ batch, FB ¼ fed-batch, SF ¼ shake flask, BR ¼ bioreactor, CF ¼ continuous fermentation, prod ¼ volumetric productivity (g/L h), YC ¼ carbon yield (g glycolipid/g substrate, %). n.a. ¼ not available, HOSO ¼ high-oleic sunflower oil. b Continuous fermentation with cell retention was successfully performed, but the steady state for constant productivity could only be maintained for 10 days.

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Chapter 9 Improved production of novel (bola) glycolipid biosurfactants

One of the first modified S. bombicola strains has been developed at our lab in collaboration with the ecological detergent company Ecover, aiming for the production of medium-chained SLs [45]. When a wild-type strain is fed with shorter-chain alcohols, most of the substrate is metabolized through the β-oxidation pathway, instead of being incorporated in SL production. Therefore the multifunctional enzyme 2 (MFE-2) gene, a key gene in the β-oxidation pathway, was blocked, increasing the production of medium-chain SLs. SLs between C8 and C14 were targeted instead of the wild-type C18:1-based SLs, which are largely water insoluble. When using lauryl alcohol (or 1-dodecanol) as a hydrophobic substrate, the production of medium-chain SL production was three times higher when using the Δmfe-2 strain compared to the wild type [40]. Similar to the production of medium-chain SLs, these authors also described the production of 20-hydroxyeicosatetraenoic acid (HETE)-based SLs with this modified strain by feeding with arachidonic acid (ARA, C20:4 (ω-6)) [41]. In 2011, Saerens et al. [17] published the discovery of the first glucosyltransferase UGTA1 of the SL production pathway of S. bombicola (Fig. 1). The results suggested the presence of two glucosyltransferases, each responsible for a specific glycosylation step in SL biosynthesis. Indeed, quickly after, the second glucosyltransferase UGTB1 was discovered (Fig. 1), and a corresponding UGTB1 knockout in S. bombicola was created, resulting in the Δugtb1 strain [16], which produced a mixture of acetylated GLs. The acetylation of GLs illustrates that the AT enzyme (Fig. 1), normally attaching acetyl groups on the 60 and 600 positions on de novo synthesized SLs, shows activity toward GLs [16] as well Unfortunately, the volumetric productivity of the original Δugtb1 strain was very low (0.01 g/L h). Nevertheless, recently a novel Δugtb1 strain was developed with a different genetic background, as will also be discussed [46]. After 270 h of cultivation using oleic acid and glucose in fedbatch fermentation, an impressive 135 g/L of GLs was produced, corresponding to a volumetric productivity of 0.5 g/L h. So, it was proven that the reason for limited GL production was mainly due to the use of a faulty background strain (with possible random mutations) and not due to the regulation of the glucosyltransferase activity or inability of the SL transporter MDR to efficiently transport GLs, as stated earlier. These observations underline the importance of the use of a fit background strain for engineering purposes. The latter is much easier nowadays then it was 12 years ago when no tools were available at all. Replacing the second glucosyltransferase UGTB1 gene with another glucosyltransferase UGT1 from Ustilago maydis [47] allowed us to produce cellobiose lipids with the resulting S. bombicola Δugtb1::UmUGT1 strain [13]. Masses of acetylated cellobiose lipids were detected, indicating that the acetyltransferase active on SLs (Fig. 1) was also able to acetylate the novel compounds, and no lactonization of the cellobiose lipids was observed. Nevertheless, very low amounts were produced, even with different modification strategies, indicating that the UGT1 is highly specific toward hydroxylated palmitic acid, and not to GLs with a C18:1 fatty acid chain [13]. However, the recent observations as discussed previously for the Δugtb1 strain that was used at that time as a background strain indicate that also for this strain changing the background strain into a rationally engineered auxotrophic mutant as described later might resolve the issue, which is the subject to an ongoing research at our lab. Another modified strain developed at InBio.be is the AT knockout of S. bombicola [15] (Fig. 2). This gene encodes the acetyltransferase within the SL biosynthetic gene cluster, which was proven to specifically attach acetyl groups on the 60 and 600 positions of the produced SLs (Fig. 1). Even though production levels with the Δat strain were low (5 g/L), the unique nonacetylated lactonic SLs are otherwise only present in very low amounts in the wild-type mixture [15]. It was later found that this strain also produces nonacetylated bola SLs [48] in addition to the previously reported nonacetylated acidic and lactonic SLs reported to be produced by this strain [15].

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Next, deletion of the CYP52M1 gene gives rise to a strain (Δcyp52m1) having lost its ability to produce SLs. Thus this strain can be used as a “fundamental base strain” to produce other interesting compounds with S. bombicola such as other types of glycolipids or other types of biochemicals, such as the bioplastic polyhydroxyalkanoates (PHAs), which consist of β-hydroxy fatty acids. By replacing the CYP52M1 gene with the phaC1 synthase of Pseudomonas resinovorans, small amounts of PHAs were obtained by the Δcyp52m1::phaC1 strain [13]. Unfortunately, no C16- or C18-based PHAs were identified, but only derivatives of up to 14 carbon atoms. Another strategy to alter the hydrophobic part of the produced SLs was published by Geys et al. [49], which consisted of replacing the cytochrome P450 CYP52M1 of S. bombicola with a clear preference of incorporating mainly C18:1 fatty acids into the glycolipid production pathway [27,50] with a heterologous cytochrome P450 enzyme with another substrate specificity. More precisely, the CYP1 of Ustilago maydis was selected, as this enzyme has been shown to be responsible for the terminal hydroxylation of C16 fatty acids. However, since CYP1 needs a cytochrome P450 reductase (CPR) for the delivery of electrons, a chimeric CYP1 had to be constructed whereby a reductase and linker of Bacillus megaterium (BMR) was coupled to CYP1. More precisely, by feeding the altered Δcyp52m1:: CYP1BMR S. bombicola strain with ethyl palmitate and glucose, the major found SLs were indeed found to be C16 acidic SLs instead of C18 SLs, as was confirmed by NMR analysis. The CYP52M1 replacement strategy has recently also been applied to alter the position of hydroxylation, and thus the “anchoring point” of the sophorose head group. Therefore Chatterjee et al. [51] expressed the Claviceps purpurea oleate hydroxylase (CpOHY) which hydroxylates oleic acid (C18:1) and palmitic acid (C16:0) at the C-12 position [52], which is in strong contrast to the specificity of the S. bombicola CYP52M1 [27]. The obtained 12-hydroxyoleic acid is commonly called ricinoleic acid, and when oleic acid was supplied as the hydrophobic substrate, the resulting strain proved to be capable of producing “branched” SLs containing ricinoleic acid in which the sophorose group is indeed attached to the C18:1 hydrophobic tail at the 12th carbon atom. In 2014, the enzyme responsible for lactonization was discovered in the secretome by Ciesielska et al. [21]. The gene encoding this enzyme is called SBLE or S. bombicola’s specific lactone esterase and is located at the other side of chromosome II of S. bombicola, compared to the SL biosynthetic gene cluster (Fig. 1). This enzyme was found to be responsible for lactonization of SLs, as a deletion strain Δsble strain does not produce any lactonic SLs anymore [21,43]. In contrast to knocking out the SBLE gene, one can also overexpress it (OE SBLE), to solely obtain lactonic SLs. This was enabled by integrating a second copy of the SBLE gene under the control of the strong constitutive phosphoglycerate kinase promoter (pGKI) at the URA3 locus [43]. In the end, a product mixture of 99% lactonic SLs was obtained (the remaining 1% corresponding to acidic SLs). By overexpressing the lactone esterase, the regulatory effect of citrate in the production medium (amongst others) was overcome, making the strain more robust toward industrial production of lactonic SLs [43]. Delbeke et al. [28] reported on the increased production of novel SLs based on petroselinic acid (C18:1, ω-12), an isomer of oleic acid (C18:1, ω-9) employing this new S. bombicola OE SBLE strain. Further, in 2016, at InBio.be, a strain was developed that efficiently produces so-called “bola(form) SLs,” displaying two sophorose heads and a lipophilic tail in between, resembling a “bola” throwing weapon, therefrom deriving its name [48,53]. When S. bombicola was disabled in its AT and SBLE genes, the resulting Δat Δsble strain surprisingly mainly produces bola SLs, consisting out of an acidic SL with an additional sophorose group linked to the free carboxylic end (confirmed by MS and NMR analysis) [48]. These bolaform SLs account for at least 74% of the SL mixture produced by this strain

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Chapter 9 Improved production of novel (bola) glycolipid biosurfactants

(the other SL type produced are nonacetylated acidic SLs due to incomplete glycosylation). Our lab has since then made a lot of progress in terms of increasing uniformity and productivity of bola SL biosynthesis (which will be discussed further on). However, still lower productivities are achieved when compared to the wild-type strain [43,44]. The latter can potentially be caused by downregulation of the remaining proteins encoded in the SL cluster. This possibility was proven by a recently developed multireaction monitoring (MRM) assay whereby expression levels of the SL cluster proteins were determined [46]. Recently, researchers at our lab described a strategy to obtain so-called “bola sophorosides (SS)” instead of bola SLs, for which the Δat Δsble Δfao1 strain was used [44]. Due to the presence of two glycosidic linkages of the sophorose units to the hydrophobic linker (instead of the bolaform SLs displaying both an ester and a glycosidic linkage), this molecule is more stable, alleviating the potential limitation of using bola SLs in formulations at increased pH. Similar end titer and productivity as for bolaform SLs were obtained for bolaform SSs. Our lab has since then continued to increase the productivity of these bolaform glycolipids by among others collaborating with the Bio Base Europe Pilot Plant for process development, optimization, and scale up. The production of bolaform SLs has been scaled up to 15 m3 scale and productivity levels of above 1 g/L h for bola SLs and bola SSs are currently obtained. Also, several -omic strategies have been performed to determine remaining bottlenecks in the strains described previously and resolve them using the developed molecular toolbox. An overview of the published achievements and strategies is described later. In a very recent achievement (unpublished results), the UGTB1 gene was deleted in the Δat Δsble Δfao1 strain, giving rise to a quadruple knock out strain: Δat Δsble Δfao1 Δugtb1 producing bola glucosides, as shown in Fig. 2.

3. Application of integrated -omics strategies for improved glycolipid biosynthesis with S. bombicola SL production as described in this chapter is a schoolbook example of how synthetic biology and pathway engineering can widen the possibilities regarding bioproduction of interesting or new-to-nature primary or secondary metabolites useful in, for example, pharmaceutical, food, agriculture, metallurgy, personal care, and cosmetic industries [54,55]. Semirational engineering approaches can be defined and refined through careful in-depth analysis and integration of so-called “omics” data of biochemical production hosts. Moreover, such an approach can be of crucial importance to characterize and subsequently optimize metabolic networks linked to biomanufacturing [56].

3.1 A multiomics approach in industrial biotechnology: A work in progress In an industrial biotechnology context, knowledge to characterize geno- and phenotypes is mostly gained from single-omics analyses such as genomics, transcriptomics, and to the lesser extent proteomics and metabolomics. The challenges encountered in the multiomic approach are manifold: limited golden standards in evaluating and classifying integration methodologies, development of novel computational approaches, unprecedented data dimensionality causing dataset overfitting and the intrinsic difficulties associated with biological data such as heterogeneity, missing values, and precision variations across layers [57,58]. In contrast to the industrial biotechnological sector, multiomics approaches have

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191

penetrated the medical field for some time now, as they offer the opportunity to thoroughly investigate the flow of information that underlies disease [59]. To holistically study diseases, biomarkers, and the complex interrelationships between biomolecules, specialized integration and interpretation methods are used, e.g., conceptual integration, model-based integration, networks, and pathway data integration. These successful approaches could also be implemented in industrial biotechnology [58,60,61]. However, unlike in the medical context, data acquisition is often more confined in biotechnological settings, and missing data often limits the system-level understanding of the microbial metabolism, especially in the case of nonconventional hosts such as S. bombicola. It is, for example, possible to gain insight into interaction mechanisms between different pathways, but not always possible to characterize clear bottlenecks to define metabolic engineering goals. Time- and labor-intensive research is required to elucidate causal relations, which is often not possible. The experimental design also often remains oversimplified due to practical limitations, undermining the usefulness of the observations. To date, only few publications implement a holistic view to identify bottlenecking locations in microbial metabolic networks. For example, in the case of lipid production in Yarrowia lipolitica, the integration of metabolic flux analysis, genomics studies, genome-scale model reconstruction and transcriptomic, metabolomic, lipidomic, and proteomic data was proven useful to achieve lipid overaccumulation and to optimize the production parameters [62]. When such information is not yet present, it is key to diversify experimental conditions as much as possible to gather information, which is useful in industrial-scale production settings. For example, the physiological responses of industrial yeasts Kluyveromyces marxianus and Saccharomyces cerevisiae were determined in multiple oxygen-limitation regimes through genome and transcriptome analyses, complemented with various sterol supplementation and uptake measurements to elucidate its importance in the oxygen requirements of K. marxianus [63]. When considering microbial biosurfactant-producing organisms, genome and transcriptome analyses are currently by far the most used analytical tools to unravel and optimize biosurfactant production pathways, e.g., rhamnolipids in Pseudomonas aeruginosa [64], mannosylerythritol lipids by Pseudozyma and Ustilago [65–67], and even much more complex structures such as surfactin produced by Bacillus species [68–71]. With the arrival of high-throughput analytical techniques, the amount of (big-) data is vastly expanding and single omics technologies are branching out into more in-depth omic layers such as epigenomics, epitranscriptomics, phosphoproteomics, etc. However, single “omics” analyses are still largely limited to black-box correlations, while the causality of processes is often unknown or misinterpreted. Therefore an integrated multiomics approach in which different -omic outputs, preferably taken simultaneously, is preferred to better understand the complex geno-phenoenvirotypic relationships of the organisms of interest and their production profiles [56]. To really reap the benefit from large multiomics data sets, effective and efficient data integration should be the number one priority.

3.2 Case studies in omics integration in glycolipid production In what follows, the developments and proceedings of (multi-)omics analyses for the development of the S. bombicola glycolipid biosynthesis platform developed at our lab will demonstrate the evolution toward a more holistic path for the production of novel glycolipids and optimization of production metrics (Fig. 3). The discovery of the SL biosynthetic gene cluster was an important first step in unravelling SL biosynthesis and broadening the industrial applicability of the yeast S. bombicola [11]. Since then, a combination of both targeted and nontargeted genomic, transcriptomic, proteomic, metabolomics, and

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Chapter 9 Improved production of novel (bola) glycolipid biosurfactants

FIG. 3 A schematic overview of the integrated multiomics approach to identify metabolites and pathways linked to glycolipid biosynthesis in S. bombicola.

lipidomic studies have been performed, aiming to further unravel this highly efficient biosynthesis. This will be described in this section. In a first step to unravel the genes involved in SL biosynthesis, our lab investigated the function of multiple P450 CYP52 enzymes present in the genome of S. bombicola. The function of different CYP

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enzymes was elucidated through transcriptomic analyses. The first -omic analysis published for S. bombicola and performed by our research group was a targeted transcriptomic analysis of three CYP genes present in the S. bombicola genome [18]. This expression analysis evaluated on different lipophilic substrates enabled us to determine the involvement of one CYP enzyme in SL biosynthesis: the CYP52M1 gene was found to be highly upregulated in the stationary phase on both hexadecane and rapeseed oil. Indeed, it was later confirmed that the corresponding enzyme is indeed responsible for the first—rate-limiting—step in SL biosynthesis [11]. The CYP52E3 and CYP52N1 were described to likely be only involved in alkane utilization and not in SL production [18,72]. A few years later, a nontargeted transcriptomic analysis (mRNA seq) was performed by our research group to investigate the parameters influencing SL biosynthesis. It is expected that SL biosynthesis is most efficient when both a hydrophilic and hydrophobic substrate is present, as they are both a constituent of this biosurfactant (see Fig. 1). In this study, we revealed that under glucose-limiting conditions, the five genes of the SL biosynthetic gene cluster are significantly downregulated [13], while no specific effect was observed for the presence/absence of the hydrophobic substrate. It was also found that all genes of the SL biosynthetic gene cluster are significantly upregulated in the stationary phase [13]. A nontargeted proteomic analysis was also performed by our group with the same goal: gather more insights in glycolipid biosynthesis with S. bombicola. The proteome of S. bombicola in the stationary and exponential phase was compared by means of a SILAC analysis [12]. In SILAC (or stable isotopelabeled amino acids) experiments, isotope-labeled amino acids are added to the medium and incorporated in the proteome. A strain deficient in its lysine biosynthesis (Δlys1) was supplemented with either 12 C6 or 13C6 L-lysine for discriminant labelling. Proteins were identified against an in-house annotated genome (4617 proteins). The proteomic data set yielded 615 quantifiable proteins of which the majority are involved in energy metabolism and translation. A subset of 147 proteins was significantly up- or downregulated between the growth and SL-producing stationary phase [12]. Later, these researchers performed an untargeted transcriptomic analysis (mRNA-seq) under these conditions and integrated this dataset with the proteomic data set. Protein expression, did for the most part, follows the general trends in mRNA transcripts. Peptides of CYP52M1 were solely identified in the stationary phase in agreement with the RT-qPCR data. A homologue of damage resistance protein 1 (DAP1) that was highly abundant in the stationary phase could act as a possible P450 enzyme enhancer [12]. All genes of the SL biosynthetic gene cluster were upregulated in the stationary phase at the gene and protein level. Changes in enzyme levels associated with the transition to the stationary phase and nutrient exhaustion include a downregulation in protein translation and an increase in protein breakdown, respectively. Metabolic changes in pathways interconnected with SL synthesis might lead to an increased pool of reducing power and required precursors. Enzymes involved in stress response and secretory processes were upregulated during SL production. Surprisingly, heat shock protein 12, a response to stress conditions, was down-regulated at both mRNA and protein level [12]. At that time, it was still unknown if an enzyme was involved in the lactonization reaction of SLs as no such gene was identified as part of the SL biosynthetic gene cluster [11]. Proteome and transcriptome analyses were thus performed, aiming to find a gene/protein which was clearly upregulated under conditions promoting the biosynthesis of lactonic SLs. Despite the extent of the proteome analysis (615 proteins), the protein responsible for lactonization of SLs was not identified [12]. Although a few candidates were identified based on the -omic data sets (in-house data), no varying phenotype was observed upon deletion of these genes. Hommel et al. [73] proposed an extracellular production of the lactonizing enzyme. It was therefore decided to perform an

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exoproteome analysis, focusing on the proteins in the extracellular environment at two different stages in both the exponential and stationary phase (four time points) [21]. At that time, the exoproteome of other biosurfactant-producing yeasts had not been published. The exoproteome was similar to other relevant yeasts and most identified enzymes were involved in cell wall organization or protein degradation (proteases). Among others, a protein with 33% homology to Candida antarctica lipase A (CALA) was identified. Upon deletion of the gene encoding this enzyme (SBLE), no lactonic SL was detected anymore [21]. This novel approach thus enabled us to confirm that lactonization of SLs is an activity catalyzed by a specific extracellular enzyme, which had until then not been identified, highlighting the strength of this “omics” strategy. The discovery of the SBLE gene also opened many perspectives for further diversification of glycolipid production in S. bombicola, i.e., production of solely lactonic SLs (by an SBLE overexpression strain) or solely nonlactonized SLs (by the △ sble strain) [14]. The lactonization activity in an aqueous environment is also a desired trait for green chemistry [74]. The discovery of a glycosylated invertase in the exoproteome also shed more light on the natural fructophilic nature of the Starmerella genera as reported [75]. This is in accordance with its natural sugar-rich niche such as floral nectar [1]. The preferential consumption of fructose over glucose in the presence of both carbon sources only occurs at high fructose concentrations [76]. Recently, Liu et al. [77] performed a nontargeted proteomic analysis of S. bombicola on different nitrogen sources aiming to produce a more uniform glycolipid product mixture. Wild-type S. bombicola produces a mixture of lactonic and acidic SLs in varying ratios depending on growth conditions and medium compositions [78]. Mainly, lactonic SLs were produced when grown on yeast extract as opposed to acidic SLs with (NH4)2SO4. When grown on ammonium sulphate, four secreted aspartic protease-like proteins (SAPL1, SAPL2, SAPL3, and SAPL4) were identified as part of the proteome. A quadruple knock-out mutant in the SAPL1-4 genes (Δsapl1-4) resulted in a 90% increase in SL titer to 61 g/L, higher yield on glucose, higher ratio of acidic SLs, and upregulation of all SL biosynthetic genes when grown on the inorganic nitrogen source [77]. This again shows the power of “omics” analyses and subsequent rational engineering strategies. Another example of a single “omics” analysis on S. bombicola, is represented by a metabolomics analysis performed by [79] to evaluate the reason for the phenotype of a S. bombicola △ mfe-2 knockout. Although SL production is possible on glucose as sole carbon source [10], supplementation with a hydrophobic substrate such as fatty acids or vegetable oil improves the productivity and titer of the process significantly. However, most of this hydrophobic substrate is channeled to the beta-oxidation pathway. A knockout in the beta-oxidation multifunctional enzyme type-2 (MFE-2), which catalyzes the second and third step of this pathway, did not result in a suspected increased yield, instead a fivefold decrease of the wild-type SL production was achieved [40]. A metabolic profiling and flux analysis of the Δmfe-2 versus the wild-type strain revealed a drastic downregulation of the TCA cycle in the former resulting in a limited supply of cytoplasmic acetyl-CoA in the Δmfe-2 knockout. Acetyl-CoA is a key metabolite in SL biosynthesis and its reduced supply likely also has a direct influence on the low acetylation degree of the SL mixture. Citric acid supplementation restored the SL production by 56% in the mutant strain [79]. The first time a multiomics approach was applied to S. bombicola when we wanted to reveal the reason for inefficient glycolipid biosynthesis in a novel strain designed at our lab. The original S. bombicola strain used for GL production (Δugtb1) displayed a very inefficient synthesis at a volumetric productivity of 0.01 g/L h [16], which is a factor 200 lower compared to the wild type operated under the same conditions. A multiomics approach comprising targeted transcriptomics (RT-qPCR), targeted

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proteomics (multireaction monitoring), genomics, and targeted metabolomics was employed to determine the inherent bottlenecks leading to inefficient GL synthesis [46]. We found that the GL biosynthetic genes were downregulated at both the transcriptional and translational level. Unexpectedly, we then found—from the metabolomics dataset—that the metabolism of the novel strain was strongly disrupted compared to the WT and also compared to strains deficient in other genes of the SL biosynthetic gene cluster: e.g., CYP52M1 or AT (acetyltransferase). We thus strongly suspected that this phenotype was not attributed to the single knock-out in UGTB1 (responsible for the second glucosylation step). The deregulation at all levels (transcriptomic, proteomic, and metabolomics) was subsequently found to be attributed to genomic changes in the base strain: a spontaneous URA3 mutant as rationally developed URA3 mutants were not available at that time [46]. The creation of rationally engineered URA3 deficient strains was later published by our research group [37]. A novel UGTB1 deletion strain was engineered based on such a rationally designed URA3 mutant (called S. bombicola PT36), and for the resulting strain, a 50-fold GL improvement was achieved (0.51 g/L h) compared to the original strain [16,46]. The multiomics approach, although not integrated, proved useful here as it helped to reveal the reason for suboptimal GL production. Our lab recently explored an in-depth metabolomics approach to investigate the productivity drop observed for a continuous bioprocess setup, through cell recycling, for bola SL [26]. In a fed-batch fermentation with the same strain, a mean volumetric productivity of bola SLs of 0.22 g/L h was achieved, with only minor acidic SL formation [44]. Cell-recycle cultivations or retentostat setups are well established for primary and secondary metabolites [80,81]. Dierickx et al. [82] further investigated such retentostat setup for continuous bolaform SL production with the S. bombicola strain △ at △ sble. In such a setup, broth is removed, cells are retained on a microfiltration unit and new nutrients are continuously added. A total of 1306 h of cultivation with the S. bombicola strain mentioned previously was achieved. Following a fed-batch phase of 108 h, a cell recycle was initiated. An initial high volumetric productivity of 0.91 g/L h decreased to 0.53 g/L h and decreased further until no more bola SLs were formed (545 h). A new batch phase was initiated to restore bola SL productivity; nevertheless, after a high productive phase, the bola SL productivity again gradually decreased to zero [82]. A feed deficient in N and P source was continuously fed as influent as SL synthesis has been described to occur under N and P limitation [8,10]. In an attempt to restore the productivity, a feed containing P sources (1 g/L KH2PO4, 0.16 g/L K2HPO4) and a feed containing both nitrogen and phosphate sources (1 g/L KH2PO4, 0.16 g/L K2HPO4, 0.4 g/L yeast extract (Gistex, DSM), 0.15 g/L NH4Cl) was introduced. Both supplementations did not restore bolaform SL production. To rule out that any genetic alterations were the cause of this productivity drop during prolonged fermentations, the genome was sequenced to look for genomic variants. No evidence for genomic variation was found, even after 1306 h of running the bioprocess, which is proof of the inherent fitness of the strain for long continuous bioprocesses [82]. The latter is an interesting fact which could help to reduce the downtime of bioprocessing installations. A new approach employing untargeted metabolomics was thus used to unravel the reasons for the described productivity drop. In total, 7722 and 2620 ions in positive and negative ionization mode, respectively, were significantly altered between the highly productive phase and the nonproducing phase. Twelve discriminating metabolites were successfully identified in this nontargeted approach [82]. The presence of 8-hydroxyguanosine during the high productive phase (prior to the productivity drop) points toward an increased oxidative RNA damage due to the presence of high levels of reactive oxygen species (ROS), which could have a downstream effect on protein synthesis as was

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hypothesized by the authors. The elevated ROS levels may be linked to the bola SL synthesis machinery through the activity of the CYP52M1 enzyme [82]. Subsequently, we decided to perform a more in-depth metabolomics and lipidomics study to further investigate the productivity drop in bola SL production [83]. The strategy was to perform two different bioprocess setups known to negatively impact bola SL biosynthesis and to look for overlapping discriminating metabolites. Two bioprocess setups were thus considered: a similar retentostat setup as already described previously and a fed batch setup where high stirring was compared with normal stirring. Under high stirring, it was found that this strain mainly produces acidic SLs and other unfinished bola SL intermediates and almost no bola SLs. Four discriminative cases with a high and low bola SL volumetric productivity could be deduced from these two set-ups [83]. Polar metabolomics on all discriminative cases yielded 22,873 components, while 21,958 components were found in the lipidomics analysis. These numbers are markedly higher than obtained in the first metabolomics study described previously. All OPLS-DA models comparing high and low bola SL productivity in the different datasets yielded a total of 200 discriminant polar metabolites and 276 discriminant lipids; 10 metabolites were identified. The metabolomic and lipidomic analyses point toward a decreased autophagy and recycling of RNA and protein degradation products. Essential nutrients gradually flushed out of the bioreactor are thus suggested to result in a decreased efficiency of bola SL synthesis by the authors [83]. Multiple omics strategies are currently further being employed at our lab, including untargeted metabolomics and proteomics in combination with genetic engineering to tackle suboptimal production efficiencies for specific glycolipid-producing strains (Fig. 1).

4. Omics development in microbial fermentations and future perspectives Over the past 100 years, there has been an immense optimization of petrochemical production processes, and these are all very well established with good efficiencies and economic viability, which is not the case for biotechnological production routes of so-called “drop ins” (i.e., the same molecules produced through another more sustainable route) or even completely novel compounds. Indeed, many biotechnological production processes are still rooted in development and their processes not yet economically viable to compete with the traditional industry [26]. Further digitization of process control allows for a more directed control of industrial fermentation processes, and there remains a large potential for continuous on-line monitoring through omics technologies. At InBio.be, we are developing an untargeted metabolomics approach permitting the real-time control of bioprocesses. The lack of extensive tools for real-time monitoring of bioprocesses impedes the robustness of the fermentation process to maintain high process efficiencies. Perturbations in the bioprocess might lead to decreased viability, unwanted side reactions or productivity decreases. Current state-of-the-art techniques are limited to the continuous monitoring of biomass viability via flow cytometry [84] or dielectric spectroscopy [85] and the targeted monitoring of key metabolites using midinfrared spectroscopy [86], nuclear magnetic resonance [87] or Raman spectroscopy [88]. Rapid evaporative ionization mass spectrometry (REIMS) eliminates the need for sample preparation and allows true real-time monitoring in a matter of seconds [89]. Contrary to other real-time monitoring techniques, such an untargeted approach covering the entire metabolome can provide profound insight into the intracellular state compared to targeted analysis of few key metabolites that only provide a narrow

References

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view. Such broad analysis allows the identification of metabolic disturbances and allows to steer the bioprocess early on to high production efficiencies. In future endeavors, it also remains key to integrate the complex data from different single omics layers to obtain a thorough systems-level understanding of the microbial metabolism of our nonconventional host, S. bombicola. Optimized integration and interpretation methods will enable to exploit the huge amount of data that is provided by the upcoming real-time tools to identify clear bottlenecks in biosynthesis or the production process of glycolipid biosurfactants.

Acknowledgments and funding The work described in this book chapter was supported by the European FP7 Project Biosurfing [Nr. 289219]; the European FP7 Project IB2Market [nr. 111043]; IWT SBO project BIOSURF [Nr. 80 050]; the European Horizon 2020 Bio-Based Industries (BBI) Consortium Project Carbosurf [Nr. 669003]; the Flemish Vlaio VIS project APPLISURF [HBC.2017.0704]; an FWO fellowship SB grant [Nr. 1SE3321N]; and an FWO fellowship FR grant [Nr. 11E0221N].

References [1] Spencer JFT, Gorin PAJ, Tulloch AP. Torulopsis bombicola sp. n. Ant Van Leeuwenh 1970;36(1):129– 33. https://doi.org/10.1007/BF02069014. [2] Rosa CA, Lachance M-A, Silva JOC, Teixeira ACP, Marini MM, Antonini Y, et al. Yeast communities associated with stingless bees. FEMS Yeast Res 2003;4(3):271–5. [3] de Graeve M, de Maeseneire SL, Roelants SLKW, Soetaert W. Starmerella bombicola, an industrially relevant, yet fundamentally underexplored yeast. FEMS Yeast Res 2018;18(7). https://doi.org/10.1093/FEMSYR/FOY072. [4] Gorin PAJ, Spencer JFT, Tulloch AP. Hydroxy fatty acid glycosides of sophorose from Torulopsis magnoliae. Can J Chem 1961;39(4):846–55. https://doi.org/10.1139/v61-104. [5] Tulloch AP, Spencer JFT, Gorin PAJ. The fermentation of long-chain compounds by Torulopsis magnoliae. I. Structures of the hydroxy fatty acids obtained by the fermentation of fatty acids and hydrocarbons. Can J Chem 1962;40(7):1326–38. https://doi.org/10.1139/v62-203. [6] Gao R, Falkeborg M, Xu X, Guo Z. Production of sophorolipids with enhanced volumetric productivity by means of high cell density fermentation. Appl Microbiol Biotechnol 2013;97(3):1103–11. https://doi.org/ 10.1007/s00253-012-4399-z. [7] Zhang Y, Jia D, Sun W, Yang X, Zhang C, Zhao F, et al. Semicontinuous sophorolipid fermentation using a novel bioreactor with dual ventilation pipes and dual sieve-plates coupled with a novel separation system. Microb Biotechnol 2018;11(3):455–64. https://doi.org/10.1111/1751-7915.13028. [8] Davila A-M, Marchal R, Vandecasteele J-P. Kinetics and balance of a fermentation free from product inhibition: sophorose lipid production by Candida bombicola. Appl Microbiol Biotechnol 1992;38(1):6– 11. https://doi.org/10.1007/BF00169410. [9] Alcon A, Santos VE, Casas JA, Garcı´a-Ochoa F. Use of flow cytometry for growth structured kinetic model development: application to Candida bombicola growth. Enzym Microb Technol 2004;34(5):399–406. [10] Albrecht A, Rau U, Wagner F. Initial steps of sophoroselipid biosynthesis by Candida bombicola ATCC 22214 grown on glucose. Appl Microbiol Biotechnol 1996;46(1):67–73. https://doi.org/10.1007/ S002530050784.

198

Chapter 9 Improved production of novel (bola) glycolipid biosurfactants

[11] van Bogaert INA, Holvoet K, Roelants SLKW, Li B, Lin Y-C, van de Peer Y, et al. The biosynthetic gene cluster for sophorolipids: a biotechnological interesting biosurfactant produced by Starmerella bombicola. Mol Microbiol 2013;88(3):501–9. https://doi.org/10.1111/MMI.12200. [12] Ciesielska K, Li B, Groeneboer S, van Bogaert I, Lin Y, Soetaert W, et al. SILAC-based proteome analysis of Starmerella bombicola sophorolipid production. J Proteome Res 2013;12(10):4376–92. https://doi.org/ 10.1021/pr400392a. [13] Roelants SLKW, Saerens KMJ, Derycke T, Li B, Lin Y-C, van de Peer Y, et al. Candida bombicola as a platform organism for the production of tailor-made biomolecules. Biotechnol Bioeng 2013;110(9):2494– 503. https://doi.org/10.1002/bit.24895. [14] Ciesielska K, Roelants SLKW, van Bogaert INA, de Waele S, Vandenberghe I, Groeneboer S, et al. Characterization of a novel enzyme-Starmerella bombicola lactone esterase (SBLE)-responsible for sophorolipid lactonization. Appl Microbiol Biotechnol 2016;100(22):9529–41. https://doi.org/10.1007/S00253-0167633-2. [15] Saerens KMJ, Saey L, Soetaert W. One-step production of unacetylated sophorolipids by an acetyltransferase negative Candida bombicola. Biotechnol Bioeng 2011;108(12):2923–31. https://doi.org/10.1002/bit.23248. [16] Saerens KMJ, Zhang J, Saey L, van Bogaert INA, Soetaert W. Cloning and functional characterization of the UDP-glucosyltransferase UgtB1 involved in sophorolipid production by Candida bombicola and creation of a glucolipid-producing yeast strain. Yeast 2011;28(4):279–92. https://doi.org/10.1002/yea.1838. [17] Saerens KMJ, Roelants SLKW, van Bogaert INA, Soetaert W. Identification of the UDP-glucosyltransferase gene UGTA1, responsible for the first glucosylation step in the sophorolipid biosynthetic pathway of Candida bombicola ATCC 22214. FEMS Yeast Res 2011;11(1):123–32. https://doi.org/10.1111/j.1567-1364.2010.00695.x. [18] van Bogaert INA, Demey M, Develter D, Soetaert W, Vandamme EJ. Importance of the cytochrome P450 monooxygenase CYP52 family for the sophorolipid-producing yeast Candida bombicola. FEMS Yeast Res 2009;9(1):87–94. https://doi.org/10.1111/J.1567-1364.2008.00454.X. [19] Claus S, Jezierska S, Elbourne LDH, van Bogaert I. Exploring the transportome of the biosurfactant producing yeast Starmerella bombicola. BMC Genomics 2022;23:1–17. https://doi.org/10.1186/S12864-02108177-X/FIGURES/9. [20] Tulloch AP, Hill A, Spencer JFT. A new type of macrocyclic lactone from Torulopsis apicola. Chem Commun (Lond) 1967;23:584–6. https://doi.org/10.1039/c19670000584. [21] Ciesielska K, van Bogaert IN, Chevineau S, Li B, Groeneboer S, Soetaert W, et al. Exoproteome analysis of Starmerella bombicola results in the discovery of an esterase required for lactonization of sophorolipids. J Proteome 2014;98:159–74. https://doi.org/10.1016/j.jprot.2013.12.026. [22] Lang S, Brakemeier A, Heckmann R, Sp€ockner S, Rau U. Production of native and modified sophorose lipids. Chim Oggi 2000;18(10):76–9. [23] Paulino BN, Pess^oa MG, Mano MCR, Molina G, Neri-Numa IA, Pastore GM. Current status in biotechnological production and applications of glycolipid biosurfactants. Appl Microbiol Biotechnol 2016;100 (24):10265–93. https://doi.org/10.1007/s00253-016-7980-z. [24] Saerens KMJ, van Bogaert INA, Soetaert W. Characterization of sophorolipid biosynthetic enzymes from Starmerella bombicola. FEMS Yeast Res 2015;15(7). https://doi.org/10.1093/femsyr/fov075. [25] van Bogaert I, Fleurackers S, van Kerrebroeck S, Develter D, Soetaert W. Production of new-to-nature sophorolipids by cultivating the yeast Candida bombicola on unconventional hydrophobic substrates. Biotechnol Bioeng 2011;108(4):734–41. https://doi.org/10.1002/bit.23004. [26] Roelants SLKW, Van Renterghem L, Maes K, Everaert B, Redant E, Vanlerberghe B, et al. Microbial biosurfactants: from lab to market. In: Banat IM, Thavasi R, editors. Microbial biosurfactants and their environmental and industrial applications. Boca Raton: CRC Press; 2018. p. 341–63. https://doi.org/10.1201/ b21950-13.

References

199

[27] Huang F-C, Peter A, Schwab W. Expression and characterization of CYP52 genes involved in the biosynthesis of sophorolipid and alkane metabolism from Starmerella bombicola. Appl Environ Microbiol 2014;80(2):766–76. https://doi.org/10.1128/AEM.02886-13. [28] Delbeke EIP, Everaert J, Uitterhaegen E, Verweire S, Verlee A, Talou T, et al. Petroselinic acid purification and its use for the fermentation of new sophorolipids. AMB Express 2016;6:28. https://doi.org/10.1186/ s13568-016-0199-7. [29] Li H, Ma X, Wang S, Song X. Production of sophorolipids with eicosapentaenoic acid and docosahexaenoic acid from Wickerhamiella domercqiae var. sophorolipid using fish oil as a hydrophobic carbon source. Biotechnol Lett 2013;35(6):901–8. https://doi.org/10.1007/s10529-013-1151-4. [30] van Bogaert INA, Roelants S, Develter D, Soetaert W. Sophorolipid production by Candida bombicola on oils with a special fatty acid composition and their consequences on cell viability. Biotechnol Lett 2010;32 (10):1509–14. https://doi.org/10.1007/s10529-010-0323-8. [31] Brakemeier A, Lang S, Wullbrandt D, Merschel L, Benninghoven A, Buschmann N, et al. Novel sophorose lipids from microbial conversion of 2-alkanols. Biotechnol Lett 1995;17(11):1183–8. https://doi.org/ 10.1007/BF00128383. [32] Brakemeier A, Wullbrandt D, Lang S. Candida bombicola: production of novel alkyl glycosides based on glucose/2-dodecanol. Appl Microbiol Biotechnol 1998;50:161–6. https://doi.org/10.1007/s002530051271. [33] Dengle Pulate V, Bhagwat S, Prabhune A. Microbial oxidation of medium chain fatty alcohol in the synthesis of sophorolipids by Candida bombicola and its physicochemical characterization. J Surfactant Deterg 2012;16:173–81. https://doi.org/10.1007/s11743-012-1378-4. [34] Dengle-Pulate V, Chandorkar P, Bhagwat S, Prabhune AA. Antimicrobial and SEM studies of sophorolipids synthesized using lauryl alcohol. J Surfactant Deterg 2014;17(3):543–52. https://doi.org/10.1007/s11743013-1495-8. [35] de Clercq V, Roelants SLKW, Castelein MG, de Maeseneire SL, Soetaert WK. Elucidation of the natural function of sophorolipids produced by Starmerella bombicola. J Fungi 2021;7(11). https://doi.org/ 10.3390/jof7110917. [36] Roelants S, Solaiman DKY, Ashby RD, Lodens S, Van Renterghem L, Soetaert W. Production and applications of sophorolipids. In: Biobased surfactants. Elsevier; 2019. p. 65–119. https://doi.org/10.1016/ B978-0-12-812705-6.00003-4. [37] Lodens S, de Graeve M, Roelants SLKW, de Maeseneire SL, Soetaert W. Transformation of an exotic yeast species into a platform organism: a case study for engineering glycolipid production in the yeast Starmerella bombicola. In: Braman JC, editor. Synthetic biology: methods and protocols, vol. 1772. New York, NY: Humana Press; 2018. p. 95–123. https://doi.org/10.1007/978-1-4939-7795-6_5. [38] Shi Y, Zhang L, Zhang M, Chu J, Xia Y, Yang H, et al. A CRISPR–Cas9 system-mediated genetic disruption and multi-fragment assembly in Starmerella bombicola. ACS Synth Biol 2022;11(4):1497–509. https://doi. org/10.1021/acssynbio.1c00582. [39] Zhang M, Shi Y, Zhang L, Zhu S, Yang H, Shen W, et al. A CRISPR–Cas12a system for multi-gene editing (CCMGE) and metabolic pathway assembly in Starmerella bombicola. Syst Microbiol Biomanuf 2022;1– 11. https://doi.org/10.1007/S43393-022-00093-9. [40] van Bogaert INA, Sabirova J, Develter D, Soetaert W, Vandamme EJ. Knocking out the MFE-2 gene of Candida bombicola leads to improved medium-chain sophorolipid production. FEMS Yeast Res 2009;9(4):610– 7. https://doi.org/10.1111/J.1567-1364.2009.00501.X. [41] van Bogaert I, Zhang G, Yang J, Liu J-Y, Ye Y, Soetaert W, et al. Preparation of 20-HETE using multifunctional enzyme type 2-negative Starmerella bombicola. J Lipid Res 2013;54(11):3215–9. https://doi.org/ 10.1194/jlr.D042226. [42] Geys R. Engineering the metabolism of Starmerella bombicola for the production of tailor-made glycolipids [dissertation]. Ghent: University of Ghent; 2016.

200

Chapter 9 Improved production of novel (bola) glycolipid biosurfactants

[43] Roelants SLKW, Ciesielska K, de Maeseneire SL, Moens H, Everaert B, Verweire S, et al. Towards the industrialization of new biosurfactants: Biotechnological opportunities for the lactone esterase gene from Starmerella bombicola. Biotechnol Bioeng 2016;113(3):550–9. https://doi.org/10.1002/BIT.25815. [44] Van Renterghem L, Roelants SLKW, Baccile N, Uyttersprot K, Taelman MC, Everaert B, et al. From lab to market: an integrated bioprocess design approach for new-to-nature biosurfactants produced by Starmerella bombicola. Biotechnol Bioeng 2018;115(5):1195–206. https://doi.org/10.1002/bit.26539. [45] Develter D, Fleurackers S, van Bogaert I. A method for the production of medium-chained sophorolipids. WO2009141407A2; 2009. [46] Lodens S, Roelants SLKW, Ciesielska K, Geys R, Derynck E, Maes K, et al. Unraveling and resolving inefficient glucolipid biosurfactants production through quantitative multiomics analyses of Starmerella bombicola strains. Biotechnol Bioeng 2020;117(2):453–65. https://doi.org/10.1002/BIT.27191. [47] Teichmann B, Linne U, Hewald S, Marahiel MA, B€ olker M. A biosynthetic gene cluster for a secreted cellobiose lipid with antifungal activity from Ustilago maydis. Mol Microbiol 2007;66(2):525–33. https://doi. org/10.1111/j.1365-2958.2007.05941.x. [48] van Bogaert INA, Buyst D, Martins JC, Roelants SLKW, Soetaert WK. Synthesis of bolaform biosurfactants by an engineered Starmerella bombicola yeast. Biotechnol Bioeng 2016;113(12):2644–51. https://doi.org/ 10.1002/BIT.26032. [49] Geys R, de Graeve M, Lodens S, van Malderen J, Lemmens C, de Smet M, et al. Increasing uniformity of biosurfactant production in Starmerella bombicola via the expression of chimeric cytochrome P450s. Colloids Interfaces 2018;2(4):42. https://doi.org/10.3390/colloids2040042. [50] Tulloch AP, Spencer JFT, Deinema MH. A new hydroxy fatty acid sophoroside from Candida bogoriensis. Can J Chem 1968;46(3):345–8. https://doi.org/10.1139/v68-057. [51] Chatterjee M, Patel JB, Stober ST, Zhang X. Heterologous synthesis and secretion of ricinoleic acid in Starmerella bombicola with sophorolipid as an intermediate. ACS Synth Biol 2022;11(3):1178–85. https://doi. org/10.1021/acssynbio.1c00457. [52] Meesapyodsuk D, Qiu X. An oleate hydroxylase from the fungus Claviceps purpurea: cloning, functional analysis, and expression in Arabidopsis. Plant Physiol 2008;147(3):1325–33. https://doi.org/10.1104/ PP.108.117168. [53] Price NPJ, Ray KJ, Vermillion KE, Dunlap CA, Kurtzman CP. Structural characterization of novel sophorolipid biosurfactants from a newly identified species of Candida yeast. Carbohydr Res 2012;348:33– 41. https://doi.org/10.1016/j.carres.2011.07.016. [54] Castelein M, Verbruggen F, Van Renterghem L, Spooren J, Yurramendi L, du Laing G, et al. Bioleaching of metals from secondary materials using glycolipid biosurfactants. Miner Eng 2021;163. https://doi.org/ 10.1016/j.mineng.2020.106665. [55] Dierickx S, Castelein M, Remmery J, de Clercq V, Lodens S, Baccile N, et al. From bumblebee to bioeconomy: recent developments and perspectives for sophorolipid biosynthesis. Biotechnol Adv 2022;54. https://doi.org/10.1016/j.biotechadv.2021.107788. [56] Amer B, Baidoo EEK. Omics-driven biotechnology for industrial applications. Front Bioeng Biotechnol 2021;9. https://doi.org/10.3389/fbioe.2021.613307. [57] Brands I. Challenges in multi-omics data integration., 2022, https://blog.biostrand.be/en/challenges-in-multiomics-data-integration. [58] Yan J, Risacher SL, Shen L, Saykin AJ. Network approaches to systems biology analysis of complex disease: integrative methods for multi-omics data. Brief Bioinform 2017;19(6):1370–81. https://doi.org/10.1093/bib/ bbx066. [59] Hasin Y, Seldin M, Lusis A. Multi-omics approaches to disease. Genome Biol 2017;18:83. https://doi.org/ 10.1186/s13059-017-1215-1.

References

201

[60] Graw S, Chappell K, Washam CL, Gies A, Bird J, Robeson MS, et al. Multi-omics data integration considerations and study design for biological systems and disease. Mol Omics 2021;17(2):170–85. https://doi.org/ 10.1039/D0MO00041H. [61] Subramanian I, Verma S, Kumar S, Jere A, Anamika K. Multi-omics data integration, interpretation, and its application. Bioinform Biol Insights 2020;14:1–24. https://doi.org/10.1177/1177932219899051. [62] Lazar Z, Liu N, Stephanopoulos G. Holistic approaches in lipid production by Yarrowia lipolytica. Trends Biotechnol 2018;36(11):1157–70. https://doi.org/10.1016/j.tibtech.2018.06.007. [63] Dekker WJC, Ortiz-Merino RA, Kaljouw A, Battjes J, Wiering FW, Mooiman C, et al. Engineering the thermotolerant industrial yeast Kluyveromyces marxianus for anaerobic growth. Metab Eng 2021;67:347– 64. https://doi.org/10.1016/j.ymben.2021.07.006. [64] Suh S-J, Invally K, Ju L-K. Rhamnolipids: pathways, productivities, and potential. In: Biobased surfactants. Elsevier; 2019. p. 169–203. https://doi.org/10.1016/B978-0-12-812705-6.00005-8. [65] Hewald S, Linne U, Scherer M, Marahiel MA, K€amper J, B€ olker M. Identification of a gene cluster for biosynthesis of mannosylerythritol lipids in the basidiomycetous fungus Ustilago maydis. Appl Environ Microbiol 2006;72(8):5469–77. https://doi.org/10.1128/AEM.00506-06. [66] Saika A, Koike H, Fukuoka T, Morita T. Tailor-made mannosylerythritol lipids: current state and perspectives. Appl Microbiol Biotechnol 2018;102(16):6877–84. https://doi.org/10.1007/s00253-018-9160-9. [67] Yu M, Liu Z, Zeng G, Zhong H, Liu Y, Jiang Y, et al. Characteristics of mannosylerythritol lipids and their environmental potential. Carbohydr Res 2015;407:63–72. https://doi.org/10.1016/j.carres.2014.12.012. [68] Hu F, Liu Y, Li S. Rational strain improvement for surfactin production: enhancing the yield and generating novel structures. Microb Cell Factories 2019;18:42. https://doi.org/10.1186/s12934-019-1089-x. [69] Wu Q, Zhi Y, Xu Y. Systematically engineering the biosynthesis of a green biosurfactant surfactin by Bacillus subtilis 168. Metab Eng 2019;52:87–97. https://doi.org/10.1016/j.ymben.2018.11.004. [70] Zhi Y, Wu Q, Xu Y. Genome and transcriptome analysis of surfactin biosynthesis in Bacillus amyloliquefaciens MT45. Sci Rep 2017;7. https://doi.org/10.1038/srep40976. [71] Zhou D, Hu F, Lin J, Wang W, Li S. Genome and transcriptome analysis of Bacillus velezensis BS-37, an efficient surfactin producer from glycerol, in response to D-/L-leucine. Microbiology 2019;8(8). https://doi. org/10.1002/mbo3.794. [72] Li W, Li J, Song X. Alkane utilization, the expression and function of cytochrome P450 in sophorolipid synthesis in Starmerella bombicola CGMCC 1576. J Microb Biotechnol 2016;5(2):58–63. [73] Hommel RK, Weber L, Weiss A, Himmelreich U, Rilke O, Kleber H-P. Production of sophorose lipid by Candida (Torulopsis) apicola grown on glucose. J Biotechnol 1994;33:147–55. https://doi.org/10.1016/ 0168-1656(94)90107-4. [74] de Waele S, Vandenberghe I, Laukens B, Planckaert S, Verweire S, van Bogaert INA, et al. Optimized expression of the Starmerella bombicola lactone esterase in Pichia pastoris through temperature adaptation, codon-optimization and co-expression with HAC1. Protein Expr Purif 2018;143:62–70. https://doi.org/ 10.1016/J.PEP.2017.10.016. [75] Gonc¸alves C, Wisecaver JH, Kominek J, Oom MS, Leandro MJ, Shen X-X, et al. Evidence for loss and reacquisition of alcoholic fermentation in a fructophilic yeast lineage. elife 2018;7. https://doi.org/10.7554/ eLife.33034.001. [76] Cabral S, Prista C, Loureiro-Dias MC, Leandro MJ. Occurrence of FFZ genes in yeasts and correlation with fructophilic behaviour. Microbiology (NY) 2015;161(10):2008–18. https://doi.org/10.1099/mic.0.000154. [77] Liu J, Zhao G, Zhang X, Song X. Identification of four secreted aspartic protease-like proteins associated with sophorolipids synthesis in Starmerella bombicola CGMCC 1576. Front Microbiol 2021;12. https://doi.org/ 10.3389/fmicb.2021.737244.

202

Chapter 9 Improved production of novel (bola) glycolipid biosurfactants

[78] Rispoli FJ, Badia D, Shah V. Optimization of the fermentation media for sophorolipid production from Candida bombicola ATCC 22214 using a simplex centroid design. Biotechnol Prog 2010;26(4):938–44. https:// doi.org/10.1002/BTPR.399. [79] Yang L, Li Y, Zhang X, Liu T, Chen J, Wei L, et al. Metabolic profiling and flux distributions reveal a key role of acetyl-CoA in sophorolipid synthesis by Candida bombicola. Biochem Eng J 2019;145:74–82. https:// doi.org/10.1016/J.BEJ.2019.02.013. [80] Ahn WS, Park SJ, Lee SY. Production of poly(3-hydroxybutyrate) from whey by cell recycle fed-batch culture of recombinant Escherichia coli. Biotechnol Lett 2001;23(3):235–40. https://doi.org/10.1023/ A:1005633418161. [81] Hoeks FWJMM, Kulla H, Meyer H-P. Continuous cell-recycle process for L-carnitine production: performance, engineering and downstream processing aspects compared with discontinuous processes. J Biotechnol 1992;22(1–2):117–27. https://doi.org/10.1016/0168-1656(92)90136-W. [82] Dierickx S, Maes K, Roelants SLKW, Pomian B, van Meulebroek L, de Maeseneire SL, et al. A multi-omics study to boost continuous bolaform sophorolipid production. New Biotechnol 2022;66:107–15. https://doi. org/10.1016/J.NBT.2021.11.002. [83] Dierickx S., Souvereyns M., Roelants S., van Meulebroek L., de Maeseneire S., Soetaert W., et al. Comprehensive metabolomics and lipidomics reveal correlation between sophorolipid biosynthesis and autophagy (unpublished manuscript, May 2022). [84] Vees CA, Veiter L, Sax F, Herwig C, Pfl€ugl S. A robust flow cytometry-based biomass monitoring tool enables rapid at-line characterization of S. cerevisiae physiology during continuous bioprocessing of spent sulfite liquor. Anal Bioanal Chem 2020;412(9):2137–49. https://doi.org/10.1007/s00216-020-02423-z. [85] Opel CF, Li J, Amanullah A. Quantitative modeling of viable cell density, cell size, intracellular conductivity, and membrane capacitance in batch and fed-batch CHO processes using dielectric spectroscopy. Biotechnol Prog 2010;26(4):1187–99. https://doi.org/10.1002/BTPR.425. [86] Alimagham F, Winterburn J, Dolman B, Domingues PM, Everest F, Platkov M, et al. Real-time bioprocess monitoring using a mid-infrared fibre-optic sensor. Biochem Eng J 2021;167. https://doi.org/10.1016/J. BEJ.2020.107889. [87] Mehendale N, Jenne F, Joshi C, Sharma S, Masakapalli SK, MacKinnon N. A nuclear magnetic resonance (NMR) platform for real-time metabolic monitoring of bioprocesses. Molecules 2020;25(20). https://doi.org/ 10.3390/MOLECULES25204675. [88] Zu TNK, Liu S, Gerlach ES, Germane KL, Servinsky MD, Mackie DM, et al. Real-time metabolite monitoring of glucose-fed Clostridium acetobutylicum fermentations using Raman assisted metabolomics. J Raman Spectrosc 2017;48(12):1852–62. https://doi.org/10.1002/JRS.5264. [89] Plekhova V, van Meulebroek L, de Graeve M, Perdones-Montero A, de Spiegeleer M, de Paepe E, et al. Rapid ex vivo molecular fingerprinting of biofluids using laser-assisted rapid evaporative ionization mass spectrometry. Nat Protoc 2021;16(9):4327–54. https://doi.org/10.1038/s41596-021-00580-8.

CHAPTER

Increasing the natural biodiversity of microbial lipopeptides using a synthetic biology approach

10

Alexis C.R. Hoste∗, Sigrid G€orgen∗, and Philippe Jacques Microbial Processes and Interactions Lab (MiPI), TERRA Teaching and Research Centre, Cross border Joint Research Unit (UMRt) BioEcoAgro, Gembloux Agro-Bio Tech/University of Lie`ge, Gembloux, Belgium

1. High natural biodiversity of lipopeptides Biosurfactants are surface-active agent that have a biological origin, which have many advantages compared to their chemical counterparts such as a lower toxicity and a better biodegradability. One of the main classes of biosurfactants are lipopeptides, which are composed of a hydrophobic fatty acid (FA) chain and a hydrophilic peptide moiety. The FA chain can be of different lengths, isomery, or saturation and can be β-hydroxylated, β-aminated, or guanylated (gFA). The peptide moiety consists of a sequence of amino acids of varying lengths that can be found in the L or D form. Different structure types of lipopeptides have been detected such as cyclic, partially cyclic, and linear lipopeptides. All lipopeptides are synthesized by nonribosomal peptide synthetases (NRPS) with the help of polyketide synthases (PKS) in some cases. These two types of biocatalyzers are multienzymatic proteins consisting of repeated modules, which function as assembly line machinery for the biosynthesis of a high set of bioactive microbial secondary metabolites [1]. Each NRPS is subdivided into modules, which contain the set of enzyme activities necessary to catalyze the incorporation of one specific amino acid into a peptide backbone. Four main catalytic domains are used in NRPSs responsible for the biosynthesis of lipopeptides. The adenylation (A) domain recognizes one amino acid residue and catalyzes its transformation into aminoacyl adenylate by a reaction with ATP. Several A domains show low specificity and can activate amino acid residues with structural similarities, such as the ones from the aliphatic group (e.g. valine, leucine, and isoleucine). This low specificity leads to an important biodiversity in lipopeptides synthesized by the same NRPS. In addition to 20 proteinogenic amino acids, nonproteinogenic amino acid residues can be incorporated, increasing the potential biodiversity of lipopeptides [2,3]. The peptidyl carrier protein (PCP) domain, also called the thiolation (T) domain, has to be transformed from apo-protein in active holo-protein by the addition on a serine residue of a phosphopantetheine arm, which is a part of coenzyme A. This transformation is catalyzed by a phosphopantetheinyl transferase encoded in Bacillus ∗

The two first authors equally contributed to the writing of this chapter.

Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00010-7 Copyright # 2023 Elsevier Inc. All rights reserved.

203

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Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

by the sfp gene [4,5]. It generates a sulfhydryl group that can react with an aminoacyl adenylate to create a thioester bond between the carboxylate group of an amino acid residue and the PCP domain. The third domain is the condensation (C) domain, which catalyzes (i) the formation of a peptide bond between an acceptor substrate (the nascent peptide) and a donor substrate (the amino acid carried by the adjacent module) and allows for (ii) the subsequent translocation of the growing chain to the following module. This C domain can be separated into two subdomains called C-donor (CD) and C-acceptor (CA), each of which is specific for the previous and current amino acid, respectively [6]. Functional subtypes of C domains have been characterized such as LCL domains (catalyze a peptide bond between two L-amino acids), DCL domains (catalyze a peptide bond between a donor D-amino acid and an acceptor L-amino acid), heterocyclization (Cyc) domains (catalyze both peptide bond formation and subsequent cyclization of Cys, Ser, and Thr residues), dual epimerization/condensation (E/C) domains (catalyze both epimerization and condensation), and C starter (CS) domains [7]. One main characteristic of NRPS responsible for the biosynthesis of lipopeptides is the presence of a CS domain that catalyzes the acylation of the amino acid residue activated by the first module [6]. This CS domain is known for its low specificity and ability to incorporate FA of different lengths and isomeries. The last domain (only present in the last module), a thioesterase (TE), is necessary for the release of the peptide and its cyclization. For some synthetases of lipopeptides, a second TE domain is present for which diverse roles have been identified. Type II TE domains are involved in the removal of nonreactive acyl residues, the removal of aberrant intermediates, the control of starter units, providing key intermediates, and releasing products [8]. Sometimes, optional domains such as methylation (M), epimerization (E), or hydroxylation domains can be present in the module [9]. An E domain will catalyze the conversion of the L-amino acid residue previously activated and fixed in the PCP domain of the module in a D-amino acid residue. Except for some lipopeptides such as locillomycin, NRPSs involved in lipopeptide synthesis operate according to a linear synthesis, with an initiation module (CS-A-T) able to recognize the first amino acid residue, followed by as many modules (C-A-T) as monomers required to complete the peptide and a last module (C-A-T-TE), which will incorporate the last monomer, release, and cyclize the lipopeptide. As previously mentioned, PKS is involved in the synthesis of some lipopeptides such as iturin. For iturin, six catalytic domains (malonyl-CoA transacylase, acyl-CoA ligase, two acyl carrier proteins, beta-keto acyl synthetase, and amino transferase) are responsible for the last steps of FA biosynthesis (last elongation and β-amination [10]) before its transfer to the first amino acid of the peptide moiety. Hansen et al. have shown that the acyl-CoA ligase domain can activate free FA through an acyl-adenylate intermediate and load it on the adjacent T domain independently of coenzyme A [11]. This mode of biosynthesis using multienzymatic proteins is responsible for the high biodiversity observed in the structures of lipopeptides. This mode of synthesis generates structural differences in the lipid moiety, such as length, isomery, saturation, β-hydroxylation, β-amination, guanylation, and in the peptide moiety, such as length, composition, amino acid residues configuration, primary sequence, and cyclisation (or not). Moreover, lipopeptides have been isolated not only from a wide variety of bacterial species but also from eukaryotes (mainly fungi), displaying the importance of these molecules (Table 1). The high biodiversity of lipopeptides induces a broad range of properties, which can be linked to multiple biological activities or various surface-active properties with many applications in different sectors (Table 2). To better understand how to develop new lipopeptides using a synthetic biology approach, it is first crucial to apprehend the natural biological diversity of lipopeptides.

1 High natural biodiversity of lipopeptides

205

Table 1 Biodiversity of lipopeptides isolated from different genera of bacteria, fungi, and animals with and without known surfactant properties. Animalia origin of lipopeptides has not yet been confirmed. Kingdom

Genus

Number of lipopeptide families isolated

Lipopeptide families described with known surfactant properties

Bacteria

Bacillus Paenibacillus Brevibacillus Aneurinibacillus Staphylococcus Actinoplanes Streptomyces Corynebacterium Rhodococcus Pseudomonas Burkholderia Achromobacter Ralstonia Janthinobacterium Serratia Anabaena Moorea Symploca Alcanivorax Lysobacter Pontibacter Aspergillus Beauveria Epichloe Fusarium Lecanicillium Microascus Mucor Penicillium Pochonia Scopulariopsis Trichoderma Aaptos Callipelta Neamphius Theonella Philinopsis Elysia

9 6 4 1 1 1 12 1 2 16 4 1 1 1 2 1 11 1 1 1 1 3 2 1 6 1 1 1 1 1 1 1 1 1 1 1 2 1

7 1 1 1 1 0 1 1 2 12 3 1 0 0 2 0 0 0 1 0 1 0 1 0 1 0 0 1 0 0 0 0 0 0 0 0 0 0

Fungi

Animalia

Table 2 Structural biodiversity and characteristics of surface-active lipopeptides. Surface tension (mN/m)

CMC (mg/L)

Reference

Kingdom

Genus

Lipopeptide

Structure type

Fatty acid

Amino acid sequence of the main representative

Charge

Activity

Bacteria

Bacillus sp.

Surfactin

Cyclic

C12–C17

Glu1-Leu2-Leu3-Val4-Asp5-Leu6-Leu/Val7

Anionic

Surfactant, antimicrobial, antiviral, anticancer, antiinflammatory, immunomodulatory, systemic resistance inducer in plant

27–37

9.2–226.4

[12–19]

Fengycin

Partially cyclic (D-Tyr3-L-Ile10)

C14–C18

L-Glu1-D-Orn2-D-Tyr3-D-allo-Thr4-L-Glu5-D-Ala/ Val6-L-Pro7-L-Gln8-L-Tyr9-L-Ile10

Anionic

Surfactant, antimicrobial, anticancer, antiviral, systemic resistance inducer in plant

42

9.1

[20–23]

Iturin

Cyclic

C14–C18

L-Asn1-D-Tyr2-D-Asn3-L-Gln4-L-Pro5-D-Asn6-L-Ser7

Nonionic

Surfactant, antimicrobial, anticancer, systemic resistance inducer in plant

46–56

45.7–85

[13,24– 27]

Kurstakin

Partially cyclic (Ser4-Gln7)

C11–C13

Thr1-Gly2-Ala3-Ser4-His5-Gln6-Gln7

Cationic

Surfactant, antimicrobial

33

144.5

[28,29]

Antiadhesin

Cyclic

C13–C17

L-Asp1-L-Leu2-L-Leu3-L-Val4-L-Val5-L-Glu6-L-Leu7

Anionic

Surfactant, antimicrobial

ND

ND

[30]

Bamylocin A

Cyclic

C13

L-Glu1-L-Leu2-Met3-Leu4-Pro5-D-Leu6-L-Leu7

Anionic

Surfactant, antimicrobial

25

ND

[31]

Licheniformin

Partially cyclic (Asp4-Tyr7)

C43

Gly1-Ala2-Val3-Asp4-Ser5-Gly6-Tyr7

Anionic

Surfactant

38

15

[32]

Paenibacillus sp.

Polymyxin

Partially cyclic (L-Dab4-Thr10)

C8–C9

L-Dab1-Thr2-L-Dab3-L-Dab4-L-Dab5-D-Phe/ Leu6-L-Leu7-L-Dab8-L-Dab9-Thr10

Cationic

Surfactant, antimicrobial

35–40

1733

[33,34]

Brevibacillus sp.

Brevifactin

Linear

C18

Pro1-Leu2-Gly3-Gly4

Nonionic

Surfactant, antimicrobial

28.5

100

[35]

Aneurinibacillus sp.

Aneurinifactin

Linear

C18

Thr1-Tyr2-Val3-Ser4-Tyr5-Thr6

Nonionic

Surfactant, antimicrobial

26

26

[36]

Staphylococcus sp.

Unnamed lipopeptide

Linear

C9

Pro1-Gly2-Gly3

Nonionic

Surfactant, antimicrobial

25

750

[37]

Corynebacterium sp.

Coryxin

Cyclic

C11

Asn1-Arg2-Asn3-Gln4-Pro5-Asn6-Ser7

Cationic

Surfactant, antimicrobial

31.4

25

[38]

Streptomyces sp.

Daptomycin

Partially cyclic (L-Thr4-L-Kyn13)

C10

L-Trp1-L-Asn2-L-Asp3-L-Thr4-Gly5-L-Orn6-L-Asp7D-Ala8-L-Asp9-Gly10-D-Ser11-3-MeGlu12-L-Kyn13

Anionic

Surfactant, antimicrobial

50

147.5– 194.5

[39–41]

Rhodococcus sp.

Rhodofactin

Linear

C14–C19

Ala1-Ile2-Asp3-Met4-Pro5

Anionic

Surfactant

30.7

23.7

[42]

Unnamed lipopeptide

Linear

ND

L-Ser1-L-Val2-L-Val/Leu3-L-Thr4-L-Thr5-D-Leu6-Ser7

Nonionic

Surfactant

30

ND

[43]

Pseudomonas sp.

Burkholderia sp.

Pseudofactin

Partially cyclic (Thr3-Leu/Val8)

C16

Gly1-Ser2-Thr3-Leu4-Leu5-Ser6-Leu7-Leu/Val8

Nonionic

Surfactant, antimicrobial, anticancer

31.5

72

[44–47]

Viscosin

Partially cyclic (D-allo-Thr3-LIle9)

C10–C12

L-Leu1-D-Glu2-D-allo-Thr3-L-Val4-L-Leu5 -D-Ser6-L-Leu7-D-Ser8-L-Ile9

Anionic

Surfactant, antimicrobial, antiviral

25–28

21.6–54

[48–53]

Orfamide

Partially cyclic (D-allo-Thr3L-Val10)

C12/C14

L-Leu1-D-Glu2-D-allo-Thr3-D-allo-Ile/D-Val4-LLeu5-D-Ser6-L-Leu7-L-Leu8-D-Ser9-L-Val10

Anionic

Surfactant, antimicrobial, insecticidal

35.7

10

[54,55]

Amphisin

Partially cyclic (D-allo-Thr3L-Asp11)

C8–C11

D-Leu1-D-Asp2-D-allo-Thr3-D-Leu4-D-Leu5D-Ser6-L-Leu7-D-Gln8-L-Leu9-L-Ile10-L-Asp11

Anionic

Surfactant, antimicrobial

24

14

[56,57]

Putisolvin

Partially cyclic (D-Ser6-L/ D-Ser12)

C6

D-Leu1-D-Glu2-D-Leu3-D-Ile4-D-Gln5-D-Ser6-DVal7-D-Ile8-D-Ser9-L-Leu10-L-Val/Xle11-L/D-Ser12

Anionic

Surfactant

ND

ND

[58,59]

Entolysin

Partially cyclic (D-Ser10-L-Ile14)

C10

D-Leu1-D-Glu2-D-Gln3-D-Val4-D-Leu5-D-Gln6-DVal7-D-Leu8-D-Gln9-D-Ser10-L-Val11-L-Leu12D-Ser13-L-Ile14

Anionic

Surfactant

ND

ND

[60]

Xantolysin

Partially cyclic (Ser7-Ile/Val14)

C10/C12

Leu1-Glu2-Gln3-Val4-Leu5-Gln6-Ser7-Val8-Leu9-Gln10Leu11-Leu12-Gln13-Ile/Val14

Anionic

Surfactant, antimicrobial, anticancer, antiviral

ND

ND

[61,62]

Tolaasin

Partially cyclic (D-allo-Thr14L-Lys18)

C8

Dhb1-D-Pro2-D-Ser3-D-Leu4-D-Val5-D-Ser6-D-Leu7D-Val8-D-Val9-D-Gln10-L-Leu11-D-Val12-Dhb13-D-alloThr14-L-Ile15-L-Hse/Gly16-D-Dab17-L-Lys18

Cationic

Surfactant, antimicrobial

38–41

420–460

[63–65]

Syringomycin

Cyclic

C10/C12/C14

L-Ser1-D-Ser2-L-Dab3-D-Dab4-L-Arg5-L-Phe6-L-Dhb7 -L-3(OH)Asp8-L-4(Cl)Thr9

Cationic

Surfactant, antimicrobial

33

1180

[66,67]

Syringopeptin 22

Partially cyclic (D-allo-Thr15L-Tyr22)

C10/C12

Dhb1-D-Pro2-D-Val3-L-Val4-D-Ala5-D-Ala6-D-Val7D-Val8-Dhb9-D-Ala10-D-Val11-L-Ala12-D-Ala13-Dhb14D-allo-Thr15-D-Ser16-D-Ala17-Dhb18-L-Ala19-L-Dab20D-Dab21-L-Tyr22

Cationic

Surfactant, antimicrobial

34.5–36

800

[68–70]

Syringopeptin 25

Partially cyclic (D-allo-Thr18L-Tyr/Phe25)

C10/C12

Dhb1-D-Pro2-D-Val3-L-Ala4-D-Ala5-L-Val6-D-Leu7D-Ala8-D-Ala9-Dhb10-D-Val11-Dhb12-D-Ala13D-Val14-D-Ala15-D-Ala16-Dhb17-D-allo-Thr18-D-Ser19D-Ala20-L-Val21-L-Ala22-L-Dab23-D-Dab24-L-Tyr/Phe25

Cationic

Surfactant, antimicrobial


50

2178

[68,71]

Syringafactin

Linear

C10/C12

L-Leu1-L-Leu2-D-Gln3-L-Leu4-D-Thr5-L-Val/Leu/ Ile6-D-Leu7-L-Leu8

Nonionic

Surfactant, antimicrobial

25

1200

[72,73]

Burriogladin

Linear

C10

Dhb1-Pro2-Gln3-Ala4-p-Hpg5-Phe6-Pro7-Thr8

Nonionic

Surfactant

ND

951.4

[74]

Haereogladin

Linear

C8

Dhb1-Dhb2-(β-OH)-Tyr3-p-Hpg4-PABA5-Thr6

Anionic

Surfactant

ND

549.8– 994.2

[74]

Holrhizin

Linear

C6-C8

L-Val1-L-Phe2-L-Glu3-L-Ile4-L-Ala5-L-Ile6

Anionic

Surfactant

ND

ND

[75,76]

Continued

Table 2 Structural biodiversity and characteristics of surface-active lipopeptides—cont’d Kingdom

Fungi

Genus

Lipopeptide

Structure type

Fatty acid

Amino acid sequence of the main representative

Charge

Activity

Surface tension (mN/m)

Achromobacter sp.

Unnamed lipopeptide

Cyclic

C21

Gly1-Gly2-Leu3-Met4-Leu5-Leu6

Nonionic

Surfactant

24.2

48

[77]

Serratia sp.

Serrawettin

Cyclic

C8–C14 (W1) or C8/C10 (W2) or C12 (W3)

L-Ser1-L-Ser2 (W1) or D-Leu/Ile1-L-Ser2-L-Thr3D-Phe4-L-Leu/Ile5 (W2)

Nonionic

Surfactant, antimicrobial, anticancer

28.8– 33.9

ND

[78–83]

Stephensiolide

Cyclic

C8/C10/C12-C14

L-Thr1-D-Ser2-L-Ser3-D-Val/Ile4-D-Ile/Val5

Nonionic

Surfactant, antimicrobial, antimalarial

ND

ND

[84]

Alcanivorax sp.

Unnamed lipopeptide

Linear

C14/C16/C18

Pro1

Nonionic

Surfactant

29.8– 32.8

40

[85]

Pontibacter sp.

Pontifactin

Linear

C16

Ser1-Asp2-Val3-Ser4-Ser5

Anionic

Surfactant, antimicrobial

25

25

[86]

Beauveria sp.

BBLP

ND

ND

ND

Cationic

Surfactant, antifungal

18

15

[87]

Fusarium sp.

Unnamed lipopeptide

ND

ND

ND

ND

Surfactant

32

1200

[88]

Mucor sp.

Unnamed lipopeptide

ND

C16-C18

ND

Cationic

Surfactant

26

15,000

[89]

Dab, 2,4-diaminobutyric acid; MeGlu, methylglutamic acid; Dhb, dehydrobutyrine; Hse, homoserine; p-Hpg, p-hydroxyphenyl glycine; PABA, p-amino benzoate; ND, not determined.

CMC (mg/L)

Reference

1 High natural biodiversity of lipopeptides

209

1.1 Bacterial lipopeptides 1.1.1 Lipopeptides produced by Bacillales Surfactin, first isolated from Bacillus sp. in 1969, was characterized as a lipopeptide and was found to be an exceptional surfactant. Since then, Bacillus sp. have been found to be an extensive source of lipopeptides with surfactant activities and this high biodiversity is divided into five main families: surfactin, iturin, fengycin, and the more recently discovered kurstakin and locillomycin (Table 2). Among these families, locillomycin is the only one that has not been described with surfactant properties. In addition to the lipopeptides isolated from the extensively studied Bacillus sp., other lipopeptides have been isolated from Bacillales such as Paenibacillus sp. and Brevibacillus sp., with some of these lipopeptides showing promising surfactant properties.

Bacillus-related lipopeptides Surfactin. Surfactins are heptapeptides, with the chiral sequence LLDLLDL, and have a FA linked to the peptide chain by lactone closure. Three main types of lipopeptides have been identified within the surfactin family. Surfactin is produced by Bacillus sp., including Bacillus subtilis, Bacillus amyloliquefaciens, and Bacillus velezensis, and has the following primary structure: Glu1-X2-Leu3-X4-Asp5-Leu6-X7. The main compound has a leucine residue in positions 2 and 7 and a valine in position 4. Lichenysin is produced by Bacillus licheniformis and has a glutamine residue at position 1 instead of a glutamic acid residue [90]. Finally, pumilacidin is produced by B. pumilus and has a leucine residue at position 4 instead of a valine residue and an isoleucine residue at position 7 instead of a leucine residue [91]. In the surfactin family, four genes encode the NRPS: srfAA, srfAB, srfAC, and srfAD. srfAA codes for the first three modules of the heptamodular NRPS, srfAB codes for the next three ones, srfAC codes for the seventh module containing a first TE domain, and srfAD codes for a second TE/acyltransferase domain that stimulates surfactin biosynthesis initiation. Within a single strain, different peptide chains can be produced. This structural diversity is due to the low specificity of some A domains in NRPS, with a high variability for the amino acids that can be accepted for positions 2, 4, and 7. At these positions, either leucine, valine, or isoleucine can be incorporated and alanine can also be incorporated at position 4 [12,92]. Furthermore, the length of the FA or its isomery can vary. For the surfactin family, the chain can vary from 12 to 17 carbon atoms and have a linear (n) or branched configuration (iso and anteiso). Surfactins have high surface-active properties, which depend on the structure of the surfactin tested. Many studies have reported the reduction of the surface tension of water in the presence of surfactin. Arima et al. first reported the reduction of surface tension by surfactin to 27 mN/m at a concentration of 0.005% [93]. Other studies have reported values ranging from 27 to 37 mN/m [12,94,95]. Within the surfactin family, surfactin and lichenysin are the most active when pumilacidin is found less active [96–98]. The FA chain (length and configuration) also has an influence on surface tension reduction, with a longer FA chain being more efficient at reducing the surface tension than a short one [99]. The critical micellar concentration (CMC) for surfactin was determined to range between 9 μM at 25°C in 0.1 M NaHCO3 solution [13,96] and 15 μM in water [94,95]. Iturin. Iturins are lipoheptapeptide interlinked with a β-amino FA of varying lengths [100]. The peptide cycle of the iturin family is characterized by a constant LDDLLDL chiral sequence of the amino acid residues. Within this family, the first three amino acids correspond to a conserved pattern (L-Asx1-D-Tyr2-D-Asn3) and the last four amino acids of the peptide moiety are variable [12]. In addition, the length of the FA varies from C14 to C18 [101].

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Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

Iturins are synthesized by a hybrid PKS-NRPS complex [14]. The iturin operon is synthesized by ituD, ituA, ituB, and ituC. ItuD is essential, specifically for iturin A, as it encodes malonyl coenzyme A transferase, which participates in the formation of the FA. ituA encodes the PKS modules responsible for the last elongation, β-amination, and incorporation of the acyl chain, and the first amino acid residue. ituB encodes an NRPS of four modules, incorporating the next four amino acids and ituC, encodes a two-module protein, incorporating the last two amino acids, and releasing the peptide. The iturin family includes a high diversity of lipopeptides with iturin A, whose peptide sequence is L-Asn-D-TyrD-Asn-L-Gln-L-Pro-D-Asn-L-Ser, linked to a FA from C14 to C18, being the most studied one [100]. The presence of a lactam ring obtained by the formation of an amide bond between the carboxylic group of the last amino acid residue and the amino group located on the β-carbon of the FA chain is one specificity of the iturin family. Overall, lipopeptides belonging to the iturin family have a lower surfactant activity than surfactin, with values ranging between 46 and 55 mN/m at an air-water interface [13]. The CMC of iturin A and C (with Asp1 instead of Asn1) were determined to be 43 and 80 μM, respectively, at 25°C in 0.1 M NaHCO3 solution, which is higher than the one obtained for surfactin [13]. Fengycin. Fengycins (also called plipastatins) are lipodecapeptides that are partially cyclic as they display an internal lactone ring in the peptide moiety between the carboxyl terminal amino acid (Ile) and the hydroxyl group in the side chain of the third amino acid residue (Tyr) [20,102]. The peptide moiety is linked to a β-hydroxy FA moiety that varies from C14 to C18. Fengycin NRPS are encoded by five genes: fenC, fenD, fenE, fenA, and fenB [103]. The first three enzymes contain two modules. fenC, fenD, and fenE, each activates two amino acids respectively, with the sixth module being less specific and able to incorporate two different amino acids [104,105]. The enzyme fenA contains three modules that activate the next three amino acids [106]. Finally, fenB activates the last amino acid, and a TE domain involved in the release of the peptide and the formation of an ester bond between the last and third amino acids of the peptide moiety [107]. The fengycin family encompasses a high diversity due to variations in both the peptide moiety and the FA chain. Fengycins A/B have the peptide sequence L-Glu-D-Orn-D-Tyr-D-allo-Thr-L-GluD-Ala/Val-L-Pro-L-Gln-L-Tyr-L-Ile linked to a β-hydroxy FA from C14 to C18. They are the two main classes of fengycins, differing in their sixth amino acid of the peptide chain [20]. A mixture of fengycin A and B reduces the surface tension of water (with 10% methanol (v/v)) to 42 mN/m, which is comprised between the surface activity of surfactin and iturin [21]. However, the CMC of fengycin is 6.25 μM in water with 10% methanol (v/v), lower than the one obtained for surfactin or iturin [21]. Kurstakin. Kurstakins are lipoheptapeptides produced by B. cereus and Bacillus thuringiensis [28]. Their peptide moiety consists of partially cyclic heptapeptides linked with an amide bond to a FA. Lactonization occurs between the carboxyl group of the terminal amino acid (Gln) and the hydroxyl group on the side chain of the fourth amino acid (Ser). Its common peptide sequence is Thr-Gly-Ala-Ser-HisGln-Gln, corresponding to a cationic peptide. Four different FA have been detected so far for kurstakin (isoC11, nC12, isoC12, and isoC13). The NRPSs involved in kurstakin synthesis are encoded by the krs locus, which consists of six genes: (i) krsE, whose product is involved in the efflux of kurstakin; (ii) the synthetase genes krsA, krsB, and krsC, which contain one, two, and four modules, respectively, with an E domain present in both the first and sixth modules; (iii) sfp, which codes for a phosphopantetheinyl transferase; and (iv) krsD, which mediates a type II TE.

1 High natural biodiversity of lipopeptides

211

Kurstakin has antifungal activity [108–110]. In addition, it has some surfactant properties, as a study by Diallo et al. showed that the surface tension of water by kurstakin was reduced to 33 mN/m, exhibiting a surface-active property comparable to the one of surfactin. However, the CMC value obtained for kurstakin (162 μM in ultrapure water) is higher than the one of surfactin [29]. Other lipopeptides from Bacillus sp. Several other lipopeptides have been isolated from Bacillus sp., with some of them showing promising surfactant characteristics and their structure and mode of biosynthesis have been unraveled. Antiadhesin: Antiadhesin is a cyclic lipoheptapeptide isolated from B. licheniformis 603. The heptapeptide (L-Asp-L-Leu-L-Leu-L-Val-L-Val-L-Glu-L-Leu) is N-acylated by a 3-hydroxy FA (from 13:0 to 17:0 with n-, iso-, and anteiso-chains). Antiadhesin showed antiadhesive activity, exhibiting its potential biosurfactant characteristics, as well as an antimicrobial activity [30]. Bamylocin A: Bamylocin A is a cyclic lipoheptapeptide isolated from B. amyloliquefaciens LP03. The heptapeptide (L-Glu-L-Leu-Met-Leu-Pro-D-Leu-L-Leu) is linked to a β-hydroxy-C13 FA. Bamylocin A exhibited antimicrobial and surfactant activities. Bamylocin reduces the surface tension of water from 72 to 25 mN/m, showing a slightly better surface tension reduction than surfactin [31]. Licheniformin: Licheniformin is a partially cyclic lipoheptapeptide isolated from B. licheniformis MS3 with an unusually long FA. The peptide moiety is formed by Gly-Ala-Val-Asp-Ser-Gly-Tyr, with a lactone linkage between the carboxyl group of Asp and hydroxyl group of Tyr residue. This lipopeptide consists of a C43H87 branched FA with a 13-fold repeated C3H6 unit linked by an amide bond to the heptapeptide. Licheniformin reduces the surface tension of water from 72 to 38 mN/m and has a CMC of 15 mg/L, showing the surfactant potential of this lipopeptide [32].

Paenibacillus-related lipopeptides In Paenibacillus sp., a high biodiversity of lipopeptides with varying activities have been isolated and characterized such as fusaricidin [111,112], heptadepsin [113], paenibacterin [114], octapeptin [115], and pelgipeptin [116]. Polymyxin, a partially cyclic FA-decapeptide, is the only Paenibacillusrelated lipopeptide already described with a surfactant activity, in addition to an antimicrobial one [33,117]. The peptide moiety of polymyxin is L-2,4-diaminobutyric acid (Dab1)-Thr2-L-Dab3-LDab4-L-Dab5-D-Phe/Leu6-L-Leu7-L-Dab8-L-Dab9-Thr10 with an amide linkage between the C-terminal threonine and the side-chain amino group of Dab4. The peptide moiety is linked to a C8-C9 FA chain. Polymyxin’s NRPS is encoded by three genes, pmxA, pmxB, and pmxE, and two genes, pmxC and pmxD, encode transporter-like proteins [118]. pmxE, pmxA, and pmxB code for five, four, and one (with a TE domain) modules, respectively. Colistin (polymyxin E) and its derivative colistin methanesulfonate reduce the surface tension of water to values between 35 and 40 mN/m and their corresponding CMCs are 1.5 and 3.5 mM for colistin and its derivative, respectively [119]. These CMCs values are higher than the ones obtained for Bacillus sp. lipopeptides, showing the lower surface activity of colistin compared to other biosurfactants.

Brevibacillus-related lipopeptides Lipopeptides isolated from Brevibacillus sp. are all linear lipopeptides that have shown antimicrobial and/or surfactant activities. Brevifactin is a linear lipopeptide with a short amino acid sequence (four amino acids, Pro-Leu-Gly-Gly) with a lactone linkage to a C18 FA. This lipopeptide was shown to have antimicrobial activities and surfactant characteristics. Brevifactin reduces the surface tension of water

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Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

from 72 to 28.5 mN/m, which is close to the surface tension reduction obtained with surfactin and a corresponding CMC of 0.1 g/L. In addition, brevifactin was highly effective to recover crude oil and was stable at high NaCl concentrations (up to 5%), displaying its potential use for bioremediation [35]. Other linear lipopeptides isolated from Brevibacillus sp. include bogorol [120], tauramamide [121], and tupuseleiamide [122].

Other lipopeptides produced by Bacillales In addition to the lipopeptides produced by bacteria from the Bacillales order described previously, two other lipopeptides have been described with surfactant activities (Table 1). Aneurinifactin is a linear hexapeptide, with the sequence Thr-Tyr-Val-Ser-Tyr-Thr, linked to a C18 FA chain isolated from Aneurinibacillus sp. This lipopeptide has a broad-spectrum antibacterial activity, in addition to surfactant activities. Aneurinifactin reduces the surface tension of water from 72 to 26 mN/m, which is comparable to the results obtained with Bamylocin A, and its CMC is estimated to be 26 mg/L in distilled water. Furthermore, aneurinifactin shows promising potential as a crude oil recovery agent as, in experimental settings, it could recover over 80% of the oil at a biosurfactant concentration of 200 mg/L [36]. Finally, an unnamed linear lipotripeptide, with the sequence Pro-Gly-Gly, was isolated from Staphylococcus sp. The peptide moiety was linked to a C9 FA chain. This lipopeptide has antimicrobial and surfactant properties. This lipopeptide reduces the surface tension of water from 72 to 25 mN/m and is stable at various pH, up to 55°C and at NaCl concentrations up to 0.3 mol/L. The CMC of this lipopeptide is 750 mg/L, which is higher than other biosurfactants [37].

1.1.2 Actinobacteria-related lipopeptides A wide variety of lipopeptides are produced by Actinobacteria, with most of them having an antimicrobial activity. That is the case of: friulimicin [123], A54145 [124], arylomycin [125], calcium dependent antibiotic (CDA) [126], enduracidin [127], laspartomycin [128], viennamycin [129], cadaside [130], malacidin [131], enamidonin [132], sarpeptin [133], and cystargamide [134]. In addition to these antimicrobial or antiviral lipopeptides, lipopeptides with surfactant activities have been isolated from Actinobacteria (Table 2). Coryxin is a cyclic heptapeptide, with the sequence Asn-Arg-Asn-Gln-Pro-Asn-Ser, linked to a C11 FA chain, isolated from Corynebacterium xerosis strain NS5 [38]. Coryxin has antibacterial activity against Gram-positive (S. aureus and Streptococcus mutans) and Gram-negative bacteria (Escherichia coli and Pseudomonas aeruginosa) and disrupts the biofilm of these bacteria. Furthermore, coryxin has surfactant properties, as coryxin lowers the surface tension of water to 31.4 mN/m, has a CMC of 25 mg/L in water, and is a strong emulsifier [38]. Daptomycin is a partially cyclic tridecapeptide linked to a C10 FA chain, isolated from Streptomyces sp. Its peptide sequence is L-Trp-L-Asn-L-Asp-L-Thr-Gly-L-Orn-L-Asp-D-AlaL-Asp-Gly-D-Ser-3-methylglutamic acid (MeGlu)-L-Kyn and the lactone ring is formed between the carboxyterminal residue of the peptide and the fourth amino acid. Daptomycin’s NRPS is coded by three genes: dptA, dptBC, and dptD, coding for five modules, six modules, and two modules and a TE domain, respectively [135]. Two additional genes found upstream of dptA, dptE, and dtpF, code for two enzymes involved in the initiation of daptomycin biosynthesis. Two nonproteinogenic amino acids, Orn6 and Kyn13, and a MeGlu residue are introduced in the peptide moiety. In addition, dtpI shares a high homology with a gene encoding for a glutamate 3-methyltransferase, which was identified in the daptomycin gene cluster and is involved in the MeGlu biosynthesis [136]. Daptomycin

1 High natural biodiversity of lipopeptides

213

is recognized for its antimicrobial activity against all clinically relevant Gram-positive bacteria, such as vancomycin-resistant enterococci, methicillin-resistant Staphylococcus aureus (MRSA), glycopeptide intermediately susceptible S. aureus, coagulase-negative staphylococci, and penicillin-resistant Streptococcus pneumoniae [39]. Daptomycin is an antibiotic approved for the treatment of MRSA and bacteremia. Daptomycin also reduces the surface tension of an aqueous buffer solution (10 mM HEPES; 100 mM potassium chloride, 2 mM calcium dichloride; pH ¼ 7.4) to
50 mN/m with a CMC of 91 μM in the same buffer [40] and a CMC of 120 μM in 0.1 M KCl [41]. Furthermore, daptomycin was shown to have a complex interaction with lipid membranes [40]. Rhodofactin is a linear pentapeptide, with the sequence Ala-Ile-Asp-Met-Pro, linked to a C14–C19 FA chain, isolated from Rhodococcus sp. TW53 [42]. This lipopeptide has surfactant characteristics as it reduces the surface tension of water to 30.7 mN/m with a CMC value of 23.7 mg/L in water. Rhodococcus sp. could be a valuable source of biosurfactants as a study from Habib et al. describes the production of a second lipopeptide in this genus, a linear heptalipopeptide with the presumable sequence L-Ser-L-Val-L-Val/Leu-L-Thr-L-Thr-D-Leu-Ser [43]. This lipopeptide reduces the surface tension of water to 29.7 mN/m with a maximal emulsification index at 45%.

1.1.3 Pseudomonas-related lipopeptides Lipopeptides produced by Pseudomonas sp. have been recently classified by Geudens and Martins into 14 families of cyclic lipopeptides, based on the length of the peptide moiety, the size of the cycle, or the length of the FA chain [137]. To these 14 families of cyclic lipopeptides, two families of linear lipopeptides can be added. Lipopeptides isolated from Pseudomonas sp. have a variety of activities such as surfactant, antimicrobial, antiviral, and anticancer (Table 2). Bananamide, fuscopeptin, corpeptin, and corrugatin are lipopeptide families isolated from Pseudomonas sp. that have not been described with surfactant properties.

Pseudofactin Pseudofactin is a partially cyclic octapeptide linked to a C16 FA chain. The peptide sequence of pseudofactin is Gly-Ser-Thr-Leu-Leu-Ser-Leu-Leu/Val and the C-terminal carboxylic group of the last amino acid forms a lactone linkage with the hydroxyl group of the third amino acid [44]. Pseudofactins have surfactant, antimicrobial, and anticancer activities [45–47]. Pseudofactin II (with a Leu8) reduces the surface tension of water from 72 to 31.5 mN/m, which is comparable to the activity of other Pseudomonad biosurfactants. The CMC of pseudofactin II is around 72 mg/L in water and pseudofactin II showed a better emulsification activity than chemical surfactants such as Tween 20 and Triton X100 [44].

Viscosin Viscosin is a family of partially cyclic nonapeptide linked to a C10–C12 FA encompassing a high biodiversity. This family includes viscosins, massetolides, White Line Inducing Principle, and pseudophomins. The amino acid sequence of viscosin is L-Leu-D-Glu-D-allo-Thr-L-Val-L-Leu-D-SerL-Leu-D-Ser-L-Ile, and the cycle occurs between the third amino acid and the last amino acid. Three genes encode viscosin: viscA, viscB, and viscC, with the particularity that viscA is located in a different locus than the other genes. viscA codes the first two modules, viscB codes the next four modules, and viscC codes the last three modules and two TE domains. The modules incorporating the D-form amino acids do not contain cognate E domains, but dual C/E domains, as seen in other Pseudomonad

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Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

lipopeptides. Lipopeptides from this family have exhibited antimicrobial activities against a range of different microorganisms and antiviral activities by providing a protective effect in embryonated eggs infected with infectious bronchitis virus, as well as an effect on influenza A virus [48,49,138]. In addition to these activities, viscosins have biosurfactant properties, as they are involved in swarming motility. The requirements for swarming are the presence of flagella, an increase in flagellar biosynthesis, cell-cell interactions, and the presence of a surfactant, which will allow the spread of bacteria over surfaces by reducing the surface tension between the bacteria and the substrate [139]. Two studies have reported the measurement of the surface tension of water in the presence of viscosin and the corresponding CMC. Saini et al. reported the reduction of the surface tension of water to 27–28 mN/m in the presence of viscosin isolated from P. libanensis strain M9-3, and its CMC was determined to be 54 mg/L in water [50]. Renard et al. determined that the surface tension of water was reduced to 25 mN/m in the presence of viscosin isolated from Pseudomonas sp. PDD-14b-2 and a corresponding CMC of 21.6 mg/L [140]. The surface activity of viscosin is comparable to the one obtained with surfactin, produced by Bacillus sp.

Orfamide Orfamide is a partially cyclic decapeptide, with the amino acid sequence L-Leu-D-Glu-D-allo-Thr-Dallo-Ile/D-Val-L-Leu-D-Ser-L-Leu-L-Leu-D-Ser-L-Val, connected to a C12 or C14 FA. The Cterminal carboxylic group of the last amino acid forms a lactone linkage with the hydroxyl of the third amino acid to form a partially cyclic lipopeptide. Three genes encode orfamide’s NRPS: ofaA, ofaB, and ofaC. ofaA codes for the first two modules of the decamodular NRPS but lacks a typical initiation module, which was reported for other NRPSs [54]. ofaB codes for the next four modules and ofaC codes for the last four modules and two TE domains, which was already reported in the biosynthesis of other Pseudomonad lipopeptides [141–143]. As described for other NRPSs such as the one of viscosin, ofaA, ofaB, and ofaC do not contain E domains, however, six dual C/E domains are found, but only five amino acids are in the D-form [54]. Orfamide A has antifungal effects on the amphotericin B-resistant strain of Candida albicans, insecticidal activity against aphids in greenhouse biocontrol trials, and surfactant properties [54,55]. Orfamide is involved in bacterial surface motility and can reduce the surface tension of water from 72 to 35.7 mN/m, which is a lower surface activity than other Pseudomonad lipopeptides, with a CMC around 10 μg/mL [54,55].

Amphisin Amphisin is a partially cyclic undecapeptide linked to a C8–C11 FA chain. The amino acid sequence of amphisin is D-Leu-D-Asp-D-allo-Thr-D-Leu-D-Leu-D-Ser-L-Leu-D-Gln-L-Leu-L-Ile-L-Asp and the cycle is formed by a lactone linkage between the C-terminal amino acid and the fourth amino acid [56]. The amphisin synthetase gene, amsY, has not been characterized yet, but it is controlled by the regulatory system GacS/GacA [144]. Amphisin has antifungal and surfactant properties and could be used in bioremediation as amphisin was effective in releasing polycyclic aromatic hydrocarbons adsorbed to sediments [56,145]. Arthrofactin, a member of the amphisin family, also has surfactant activities; it reduces the surface tension of water to 24 mN/m and has a CMC of 10 μM in 10 mM phosphate buffer (pH 8.0), which is more effective than surfactin [57]. Arthrofactin was also shown to be a better oil remover than synthetic surfactants such as Triton X-100 and sodium dodecyl sulfate in an oil displacement assay [57].

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215

Putisolvin Putisolvin is a partially cyclic dodecapeptide, with the amino acid sequence D-Leu-D-Glu-D-LeuD-Ile-D-Gln-D-Ser-D-Val-D-Ile-D-Ser-L-Leu-L-Val/Xle-L/D-Ser, for putisolvin I and II, respectively [58]. The C-terminal carboxylic acid group forms an ester with the hydroxyl sidechain of Ser9 and the peptide is connected to a C6 FA chain. Three genes encode putisolvin’s NRPS: psoA, psoB, and psoC, which code for two modules, seven modules, and three modules and two TE domains, respectively [59]. In addition, one gene (psoR), located upstream of psoA, is required for the expression of the pso cluster and two genes (macA and macB), located downstream of psoC, are involved in the production or in the export of putisolvin [59]. Putisolvin has surfactant activities, as it reduces the surface tension of the culture medium by 25 mN/m when compared to sterile culture medium [58]. In addition, putisolvin inhibited biofilm formation by other Pseudomonas spp. [58].

Entolysin Entolysin is a partially cyclic tetradecapeptide, with the amino acid sequence D-Leu-D-Glu-D-GlnD-Val-D-Leu-D-Gln-D-Val-D-Leu-D-Gln-D-Ser-L-Val-L-Leu-D-Ser-L-Ile with a lactone ring formed between the C-terminal carboxyl group (Ile) and the tenth amino acid (Ser) [60,146]. The peptide is linked to a C10 FA. Entolysin’s NRPS is encoded by three genes, etlA, etlB, and etlC, which code for two, eight, and four modules of the NRPS, respectively, with etlC also coding for two TE domains [60]. The genes encoding the NRPS are located in two different loci of the genome. In addition to these three genes, other genes are involved in entolysin’s production such as etlR, a transcriptional regulator. GacS/GacA, a two-component system, regulates entolysin’s production at the biosynthetic gene level, as described for amphisin. As for other Pseudomonad lipopeptides, no E domains are present in the NRPS. Entolysin was tested in a “drop-collapsing assay,” and it was shown that entolysin decreases the surface tension of the culture medium and is required for swarming motility and hemolytic activity [60].

Xantolysin Xantolysin is a partially cyclic tetradecapeptide linked to a C10 or C12 FA. The sequence of the peptide moiety is Leu-Glu-Gln-Val-Leu-Gln-Ser-Val-Leu-Gln-Leu-Leu-Gln-Ile/Val, with a lactone ring formed between the carboxyterminal residue of the peptide and the seventh amino acid [61]. Three genes encode xantolysin’s NRPS: xtlA, xtlB, and xtlC. This NRPS is similar to the one producing entolysin; however, a shift in the eighth module, incorporating Ser instead of Val in entolysin, generates a different amino acid sequence from entolysins and also changes xantholysin’s macrocyclization into an octacyclic structure, distinct from the pentacyclic closure in entolysin [61]. Xantholysin has antifungal activity and antibacterial activity against Gram-positive and Gram-negative bacteria, an antiviral activity against hepatitis C virus, a supportive role in biofilm formation, and facilitation of surface colonization through swarming [61,62].

Tolaasin Tolaasin is a partially cyclic octadecapeptide linked to a C8 FA. The sequence of the peptide moiety is Dhb-D-Pro-D-Ser-D-Leu-D-Val-D-Ser-D-Leu-D-Val-D-Val-D-Gln-L-Leu-D-Val-Dhb-Dallo-Thr-L-Ile-L-homoserine (Hse)/Gly-D-Dab-L-Lys for tolaasin I/II, with a lactone ring formed between the carboxyterminal residue of the peptide and the 14th amino acid [63]. The large NRPS gene cluster includes five genes: taaA, taaB, taaC, taaD, and taaE. taaA, taaB, taaC, and taaD each codes

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three modules and taaE codes the last six modules with two TE domains [147]. Tolaasin has antimicrobial activity, in addition to surfactant characteristics [63]. Tolaasin has been reported to decrease the surface tension of water to 38–41 mN/m and to have a CMC between 0.42 and 0.46 g/L in water, which is a lower surfactant activity than other Pseudomonad biosurfactants [64].

Syringomycin Syringomycin is the only lipopeptide family isolated from Pseudomonas sp. to be fully cyclic and is one of the four biosurfactant lipopeptides produced by P. syringae. The amino acid sequence of the peptide is L-Ser-D-Ser-L-Dab-D-Dab-L-Arg-L-Phe-L-dehydrobutyrine (Dhb)-L-3(OH)Asp-L-4(Cl) Thr with the β-carboxy group of the C-terminal residue closing a macrocyclic ring on the OH group of the N-terminal Ser [66]. The N-terminal Ser is N-acylated by a C10, C12, or C14 FA, depending on the syringomycin isoform [67]. Five genes are involved in the production of syringomycin, two encoding the NRPS (SyrB1, SyrE) and three modifying protein systems (SyrB2, SyrC, SyrP). The first eight modules are coded by SyrE and the last module by SyrB1. However, the rule of colinearity is not followed as the ninth module, responsible for the last amino acid incorporation is located in the upstream region [142]. L-Thr9 is then chlorinated to 4-Cl-L-Thr by the nonheme Fe(II) halogenase SyrB2 [148]. The aminoacyl-transferase SyrC shuttles the Thr/Cl-Thr moiety between the pantetheinyl arms of the thiolation domain of SyrB1 and the thiolation domain in module nine of SyrE [149]. Finally, the hydroxylation of Asp8 is catalyzed by SyrP [150]. Syringomycin has surface active properties, as well as antimicrobial activities with a broad antifungal action [151]. A mixture of syringomycin isoforms has a CMC of 1.18 g/L in water and reduces the surface tension of water from 72 to 33 mN/m, which is comparable to the results obtained with other Pseudomonad biosurfactants, such as pseudofactin [67]. The CMC obtained for syringomycin is significantly higher than the CMCs obtained for other Pseudomonads biosurfactants, showing its lower surfactant activity.

Syringopeptin 22 Syringopeptin 22 is a partially cyclic lipopeptide with a 22 amino acid peptide, with the sequence DhbD-Pro-D-Val-L-Val-D-Ala-D-Ala-D-Val-D-Val-Dhb-D-Ala-D-Val-L-Ala-D-Ala-Dhb-D-allo-ThrD-Ser-D-Ala-Dhb-L-Ala-L-Dab-D-Dab-L-Tyr linked to a C10 or C12 FA chain isolated from P. syringae [68]. A lactone ring is formed between the carboxyterminal residue of the peptide and the 15th amino acid. Syringopeptin 220 s NRPS is coded by three genes: sypA, sypB, and sypC. sypA and sypB code for 5 modules and sypC codes for 12 modules and 2 TE domains [143]. Syringopeptin 22 has antimicrobial activity against Gram-positive bacteria and yeasts, as well as surfactant activities [69]. Syringopeptin 22 has a CMC of 0.8 g/L in water and lowers the surface tension of water to 34.5–36 mN/m, close to the surface activity of other Pseudomonad lipopeptides such as syringomycin [70].

Syringopeptin 25 Syringopeptin 25 is a partially cyclic lipopeptide with a 25 amino acid peptide, with the sequence DhbD-Pro-D-Val-L-Ala-D-Ala-L-Val-D-Leu-D-Ala-D-Ala-Dhb-D-Val-Dhb-D-Ala-D-Val-D-AlaD-Ala-Dhb-D-allo-Thr-D-Ser-D-Ala-L-Val-L-Ala-L-Dab-D-Dab-L-Tyr/Phe linked to a C10 or C12 FA chain isolated from P. syringae [68]. A lactone ring is formed between the carboxyterminal residue of the peptide and the 18th amino acid. Syringopeptin 25 has an antibacterial activity against Gram-positive bacteria and an antifungal activity [71]. Syringopeptin 25 also

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exhibits surface activity properties, as it reduces the surface tension of water to
50 mN/m with a CMC of 0.9 mM [152].

Syringafactin Syringafactin is a linear octapeptide linked to a C10 or C12 FA chain isolated from P. syringae [72]. The composition of the peptide sequence is L-Leu-L-Leu-D-Gln-L-Leu-D-Thr-L-Val/Leu/Ile-D-LeuL-Leu. In the biosynthetic gene cluster, five genes are involved in the production of syringafactin: syfR, syfA, syfB, syfC, and syfD. syfA codes for the first three modules of the NRPS and syfB codes for the last five modules and two TE domains [72]. syfR is a LuxR homolog that could play a role in the regulation of the production of syringafactin, while syfC and syfD show a sequence similarity to subunits of a multidrug efflux pump [72]. Syringafactin is involved in the swarming motility of the producing strain, and syringafactin-producing strains can degrade diesel in an artificially contaminated sand, exhibiting the surfactant properties of syringafactin [72,153]. Syringafactin reduces the surface tension of water to 25 mN/m and its CMC was determined to be 1.2 g/L [140]. This high CMC is comparable with the CMC obtained for other Pseudomonad lipopeptide such as syringopeptin 22 or syringomycin. These lipopeptides have a lower surfactant activity than other Pseudomonad biosurfactants.

1.1.4 Burkholderiales-related lipopeptides Lipopeptides produced by Burkholderiales include lipopeptides isolated from Burkholderia sp., Achromobacter sp., Ralstonia sp., and Janthinobacterium sp. These lipopeptides have surfactant or antimicrobial properties or are involved in the virulence of their producing strain (Table 2). Three linear lipopeptides have been isolated from Burkholderia sp. with surfactant characteristics: burriogladin, haereogladin, and holrhizin. Burriogladin is an octapeptide and haereogladin is a hexapeptide, linked to C10 or C8 FA chains, respectively [74]. Burriogladin’s peptide sequence is Dhb-Pro-Gln-Ala-p-hydroxyphenyl glycine (p-Hpg)-Phe-Pro-Thr and haereogladin’s is Dhb-Dhb(β-OH)-Tyr-p-Hpg-p-amino benzoate (PABA)-Thr. Burriogladin’s NRPS is coded by two genes, bgdA and bgdB, coding for three and four modules, respectively. Haereogladin’s NRPS is coded by one gene, hgdA, coding for the five modules. Interestingly, both lipopeptides have an additional amino acid in their peptide moiety compared to the number of modules of their NRPS. The most plausible explanation is that a Thr residue is introduced during offloading [74]. Haereogladin and burriogladin both have surfactant properties as they decreased the contact angle of a water droplet on a hydrophobic surface. A haereogladin that lacks the threonine moiety did not reduce the contact angle, exhibiting the critical role of the threonine residue for the surfactant properties of haereogladin. The surface activity of haereogladin and burriogladin is close to the one of Tween 80, used as positive control in the study [74]. The CMC values of haereogladin and burriogladin in water were determined to be 0.64–1.12 and 0.99 mM, respectively. Finally, it was shown that burriogladin plays a pivotal role in swarming motility of Burkholderia sp. and that haereogladin promotes biofilm formation [74]. Holrhizin A is a linear hexapeptide, with the sequence L-Val-L-Phe-L-Glu-L-Ile-L-Ala-L-Ile, linked to a C8 FA chain [75]. Other holrhizin isoforms were later isolated, varying in their amino acid length (from 4 to 6 amino acids), composition and FA chain length (from C6 to C8) [76]. Holrhizin’s NRPS is coded by one gene, holA, coding for six modules. The biodiversity observed in holrhizin is due to the substrate flexibility of the fourth, fifth and sixth adenylation domains and from the starter C domain [76]. Holrhizin is a virulence factor and a biosurfactant as it reduces surface tension and is involved in the formation of biofilms and cell motility [75]. The measurement of the contact angle of

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an aqueous holrhizin solution on a hydrophobic surface found that holrhizin A exhibits strong surfactant activities comparable to Tween 80 [75]. An unnamed lipopeptide has been isolated from Achromobacter sp. HZ01 with biosurfactant properties [77]. This lipopeptide is a cyclic hexapeptide, with the sequence Gly-Gly-Leu-Met-Leu-Leu, linked to a C21 FA chain. This biosurfactant reduces the surface tension of water to 24.2 mN/m, lower than the one obtained with surfactin, and has a CMC of 48 mg/L in water. In addition to the biosurfactant lipopeptides isolated from Burkholderiales, three other lipopeptides not characterized as biosurfactants have been described: icosalide, ralsolamycin, and jargacin.

1.1.5 Serratia-related lipopeptides Two lipopeptides with biosurfactant activities have been isolated from Serratia sp.: serrawettin, which was first referred to as serratamolide and the recently isolated stephensiolide (Table 2). The high diversity of lipopeptides that have been characterized within these two families is detailed later.

Serrawettin Serrawettin is a family of cyclic lipopeptides that were first isolated from a Serratia marcescens strain and considered to be identical to serratamolide A [78]. Three different lipopeptides are gathered into this family: serrawettin W1, W2, and W3, isolated from S. marcescens with serrawettin W2 also isolated from S. surfactantfaciens [78–80]. Serrawettin W1 is a symmetrical dilactone molecule with two L-serine amino acids linked to two βhydroxy FA that can be of varying length (C8–C14), contributing to the diversity of serrawettin W1 isoforms [78]. The gene encoding the serrawettin W1 NRPS is swrW, which encodes an uncommon uni-modular NRPS incorporating only L-Ser [154]. Serrawettin W2 is a pentapeptide, with the amino acid sequence D-Leu/Ile-L-Ser-L-Thr-D-PheL-Leu/Ile, linked to a β-hydroxy FA moiety of varying length (C8 or C10) [79]. Serrawettin W2 is produced by a hybrid PKS-NRPS complex [81]. The serrawettin W2 operon is synthesized by swrE, swrF, swrG, and swrA. The PKS genes swrE, swrF, and swrG, encode for an acyltransferase, a ketosynthase, and a keto reductase domain. swrA encodes an NRPS of five modules with a TE domain, incorporating the five amino acids. Serrawettins W2 with varying amino acid compositions have been isolated [155]. Serrawettin W3 is a pentapeptide containing the amino acids Thr, Ser, Val, and Leu/Ile linked to a C12 FA chain [79]. The exact structure of serrawettin W3 has not been elucidated yet. Serrawettin W1 and W2 have been shown to have antibacterial activity against pathogenic Grampositive and Gram-negative bacteria including drug-resistant S. aureus and antifungal activity [81–83]. Serrawettin W2 exhibited a higher cytotoxicity against Hela and Caco2 cells compared to nonmalignant cells (Vero and HEK293 cells), showing the potential anticancer activity of this lipopeptide [81]. In addition to these activities, serrawettins are involved in swarming motility and showed biosurfactant characteristics as serrawettin W1, W2, and W3 reduce the surface tension of water to 32.2, 33.9, and 28.8 mN/m, respectively [80,156].

Stephensiolide Stephensiolide is a cyclic pentapeptide, with the amino acid sequence L-Thr-D-Ser-L-Ser-D-Val/IleD-Ile/Val linked to a FA chain [84]. The FA chain can be constituted of 8, 10, 12, 13, or 14 carbons, leading to different isoforms, from A to K. There is a unique gene, sphA, encoding for the five modules

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of the NRPS with a TE domain [84]. Stephensiolide showed antibacterial activity against B. subtilis and antimalarial activity against Plasmodium falciparum [84]. Finally, stephensiolide promotes swarming motility of the producing strain, acting as a biosurfactant [84].

1.1.6 Cyanobacteria-related lipopeptides A wide variety of lipopeptides have been isolated from Cyanobacteria, mostly from Moorea producens (formely Lyngbya majuscula), with different activities. Despite this biodiversity, to our knowledge, no characterized lipopeptide isolated from Cyanobacteria has been described for its biosurfactant activities. Lipopeptides from Cyanobacteria include anabaenolysin [157], antillatoxin [158], apramide [159], barbamide [160], carmabin [161], curacin [162], hectochlorin [163], laxaphycin [164], majusculamide [165], malyngamide [166], microcolin [167], somamide [168], and symplostatin [169]. More research is needed as the wide variety of this group of lipopeptides could include potential biosurfactants with interesting characteristics.

1.1.7 Other bacterial lipopeptides Lipopeptides have also been isolated from other bacteria, with some of these lipopeptides having surfactant characteristics (Table 1). An unnamed linear lipopeptide has been isolated from Alcanivorax dieselolei B-5 by Qiao and Shao [85]. The lipopeptide was identified as a long FA chain (C14, C16, or C18) linked to a unique proline. This lipopeptide reduced the surface tension of water to 29.8–32.8 mN/m and its CMC was determined to be 40 mg/L in water, exhibiting the surfactant characteristics of this lipopeptide. Additionally, the lipopeptide was stable and retained its surfactant activity over a wide range of pH [3–10] and temperature (40–100°C). Pontifactin, a linear pentapeptide linked to a C16 FA chain has been isolated from Pontibacter korlensis [86]. The peptide sequence of pontifactin is Ser-Asp-Val-Ser-Ser. Pontifactin has antibacterial activity against Gram-positive (S. mutans and Micrococcus luteus) and Gram-negative bacteria (Salmonella typhi and Klebsiella oxytoca) and an antibiofilm activity. Pontifactin has surfactant characteristics as it reduces the surface tension of water to 25 mN/m, which is better than for surfactin, has a CMC of 25 mg/L in water, and an emulsification activity. These activities were stable over a broad range of pH [4–10] and temperature (up to 100°C).

1.2 Eukaryotic lipopeptides 1.2.1 Yeast and fungi-related lipopeptides Most of the lipopeptides described in the literature are lipopeptides that have been isolated from bacteria (Table 1). However, the fungi kingdom is also a reservoir of lipopeptide-producing organisms with various biological activities. From the characterized fungi-lipopeptides, only a few of them have shown surfactant activities, with most of the other lipopeptides having antimicrobial, antifungal, or anticancer activities. Three lipopeptides, isolated from Beauveria sp., Fusarium sp., and Mucor sp., have shown surfactant characteristics (Table 2). BBLP, isolated from Beauveria sp., has an unknown structure but has an antifungal activity against Microsporum canis and was able to reduce the surface tension of water to 18 mN/m with a CMC of 15 mg/L in water, exhibiting the high biosurfactant potential of this lipopeptide [87]. The lipopeptide isolated from Fusarium sp. by Qazi et al. reduced the surface tension of water

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to 32 mN/m and had a CMC of 1.2 g/L in water [88]. The CMC obtained for this lipopeptide shows its lower surface activity compared to other lipopeptides, but it is comparable to the CMC obtained for some Pseudomonad lipopeptides (such as syringomycin and syringafactin). However, the structure of this lipopeptide has not been elucidated yet. The third fungi-lipopeptide with surfactant activity was isolated from a Mucor circinelloides strain by Sa´ Alencar do Amaral Marques et al. [89]. The peptide sequence of the lipopeptide was not determined, but it is a cationic peptide linked to a C16 or C18 FA chain. The lipopeptide reduced the surface tension of water to 26 mN/m, with a CMC of 15 g/L in water. This lipopeptide showed promising bioremediation properties as it could recover 95.9% of motor oil adsorbed on a clay soil sample and the lipopeptide remained stable at varying pH, high temperatures, and high NaCl concentrations. However, the high CMC obtained for this lipopeptide (15 g/L) exhibits the weak surfactant activity of this lipopeptide. Other lipopeptides have been isolated from fungi, including the following: aspochracin [170], echinocandin [171], beauverolide [172], fusaristatin [173], apicidin [174], fusaoctaxin A [175], gramillin [176], W493 [177], alveolaride [178], emericellamide [179], epichlicin [180], fellutamide [181], scopularide [182], trichoderin [183], and verlamelin [184]. For a more detailed listing of nonribosomal peptides and lipopeptides produced by fungi, see the reviews of Zhao et al. [185] and Bills et al. [186].

1.2.2 Animal-related lipopeptide A few lipopeptides have been isolated from deep-sea sponges or mollusks; however, none of them have been tested for their potential surfactant characteristics. Lipopeptides isolated from sponges include callipeltin [187], ciliatamide [188], neamphamide [189], and papuamide [190]. Three lipopeptides have been isolated from two distinct mollusks: kulolide, kulomo’opunalide [191] and kahalalide [192]. There is still a doubt about the animal origin of these compounds which could be produced by symbiotic microorganisms.

1.3 Uncharacterized biosurfactant lipopeptides Studies have reported the discovery of potentially new lipopeptides from a wide variety of organisms showing surfactant properties, without a structural characterization of the biosurfactant. These studies describe the isolation of promising biosurfactant lipopeptides, isolated from not yet reported biosurfactant-producing genus, demonstrating the wide biodiversity of biosurfactant lipopeptides [193–198]. Other studies report the isolation of biosurfactant lipopeptides from known lipopeptide-producing genus, which could correspond to already characterized lipopeptides or new lipopeptides [199–204].

2. Production of novel lipopeptides Lipopeptides present two main advantages compared to chemical surfactants: their biodegradability and, for some of them such as surfactin or lichenysin, their high surfactant activity at low CMC [205]. Their complex mode of biosynthesis has two consequences: (i) in average they are produced in lower yields leading to higher costs than other biosurfactants such as sophorolipids, for example [206] and (ii) their structures and thus their properties exhibit a very high biodiversity as demonstrated in the first part of this chapter. This biodiversity could be increased in the future by using metabolic

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engineering and synthetic biology approaches [207]. This should lead to novel compounds, more performant and especially adapted to some surfactant niche markets. These approaches could also improve the lipopeptide yield and reduce their cost price. The second part of the chapter will thus focus on these methods used to generate (a) new lipopeptides, (b) to improve the production of lipopeptides in the native host, and (c) to allow a heterologous production through synthetic biology.

2.1 Change in the composition of amino acids The diverse strategies developed to modify the composition of the amino acids to change the structure of lipopeptides and thus their properties are presented in this following sections.

2.1.1 Precursor directed biosynthesis A common strategy used is called precursor directed biosynthesis. This strategy is based on the feeding with a specific amino acid or the increase by metabolic engineering of the intracellular concentration of a specific amino acid residue and the flexibility of an A domain to activate different amino acid residues with relatively close structure [208]. For example, the knock-out of codY in B. subtilis increases the valine intracellular pool and the production of [Val7] surfactin [209]. This method also allows the incorporation of nonnatural or natural amino acids into the final product, for example, with the incorporation of fluorinated nonproteinogenic amino acid into iturin [210–212]. This strategy allows the production of a diversity of molecules, but, in some cases, with a low yield, because of the competition of the feeding amino acid and the natural amino acid fixed by the A domain. Therefore this strategy must be coupled with another one using mutagenesis to enhance the incorporation of new precursors. This consists in deleting genes responsible for the synthesis of the natural amino acid to reduce or suppress the synthesis of the targeted amino acid and avoiding competition between the natural amino acid and the one fed. Thus only the fed one will be incorporated into the lipopeptide [208,213,214]. This method was successfully applied to produce new analogues of CDA from Streptomyces coelicolor. CDA production was abolished by deleting the gene hmaS, a putative 4-hydroxymandelic acid synthase-encoding gene. The feeding with exogenous 4-hydroxymandelate, 4-hydroxyphenylglyoxylate, or 4-hydroxyphenylglycine reestablished CDA production by the ΔhmaS mutant leading to the production of novel lipopeptides with modified arylglycine residues [215].

2.1.2 Substrate recognition domain The A domain controls which substrate enters the assembly line and gets attached to the adjacent T or PCP domains [216]. Thus a second method to change the composition of the final lipopeptides produced is by mutation of the A domain. The site-directed mutagenesis technique was successfully applied on the A domain of surfactin nonribosomal synthetases in order to change the specific recognition of the amino acid incorporated, leading to the shift from L-Glu to L-Gln in module 1 and from L-Asp to L-Asn in module 5 [217]. In fusaricidin synthetase, the adenylation domain of the third module was mutated to mimic the Adomains of gramicidin S synthetase 1 and tyrocidine synthetase 1, both of which predominantly activate phenylalanine. This leads to the production of an analogue-LI-F07 fusaricidin that showed more potent antimicrobial activity than other analogues [218].

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2.1.3 Domain and module exchange As previously explained, each module of NRPS is composed of three domains: C domain, A domain, and T (¼ PCP) domain. These three domains are globally responsible for the incorporation of one amino acid [219]. The strategy described here relies on an exchange between two domains or two modules, leading to the production of new lipopeptides (Fig. 1). However, this exchange is more successful if the domains exchanged come from the same (or closely related) biosynthetic gene cluster [208]. Some studies successfully produced new lipopeptides, by domain substitution between D-Ala8 and D-Ser11 by exchanging the biosynthetic gene belonging to daptomycin and CDA, which led to the production of novel lipopeptides [220]. Additionally, Bode’s team revolutionized the module swapping strategy with the development of the concept of the exchange unit condensation domain (XUC). They have shown that the C domain is subdivided into two subdomains surrounding a catalytic center: a N-terminus subdomain donor of amino acid and a C-terminus subdomain acceptor of amino acid. Thus an XUC is ideally designed as Cdonor-A-T-Cacceptor [221].

2.2 Altering the number of monomers This strategy consists in changing the number of amino acids in the produced lipopeptides, by increasing or reducing the numbers of modules [208,219] (Fig. 1). Some attempts were made to reduce the number of modules, e.g., L-Leu2 of surfactin was removed by deleting the C, A, and PCP domains of module 2 of the gene srfA-A. The new surfactin exhibited lower hemolytic bioactivities [222].

FIG. 1 Representation of NRPS domains and the possible modifications that can be made to modify the amino acids moiety by: changing the composition of amino acids, deletion/insertion of modules, alteration of monomer connectivity, insertion of tailoring enzyme, or combination of different biosynthetic gene clusters. C, condensation domain; AA, amino acid; T, thiolation domain, E, epimerization domain; TE, thioesterase domain.

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Other attempts produced three novel surfactins individually lacking Leu3, Asp5, and Leu6 by deleting the whole respective module composed of C-A-T and/or E domain. ΔLeu3 and ΔLeu6 showed reduced toxicity and ΔAsp5 showed a stronger inhibition against B. pumilus and M. luteus. However, the final production yield was low, with 0.82, 1.35, and 0.96 mg/L, respectively for ΔLeu3, ΔAsp5, and ΔLeu6 [223]. This low production could be overcome by using the XUC concept previously described, where the deletion should be done between the two subdomains of the C domain to form a Cdonor-A-TCacceptor, preventing a loss in production [221]. Another method has been used to decrease the number of amino acids in the final lipopeptide by adding a TE domain between two modules. A TE domain, responsible for macrocyclization, was placed after a thiolation domain of different modules of plipastatin synthetase: domain 7 (L-Pro), domain 8 (L-Gln), and domain 9 (L-Tyr). The TE was capable of recognizing and catalyzing the lactone formation between L-Tyr9 with the last few residues L-Pro7 and L-Gln8 at the C-terminus, producing circular plipastatin lipopeptides with a reduced number of amino acid residues [224].

2.3 Altering monomer connectivity In the NRPS system, the last TE domain releases the linear lipopeptide using a hydrolytic cleavage, then cyclizes the lipopeptide, via, usually, an ester bond (e.g., surfactin, daptomycin, fengycin, kurstakin, etc.) or in a few cases an amide bond (e.g., iturin, mycosubtilin, bacillomycin, etc.) between the FA chain and the last amino acid or an ester bond between the hydroxyl group of an amino acid residue and the last amino acid or an amide bond between the amine group of an amino acid residue and the last amino acid [14,225]. Modifying the ring closure could have a huge positive impact on the resistance of bacteria to lipopeptide antibiotics. For example, daptomycin, a commonly used antibiotic against different bacteria-related diseases [39], can be easily inactivated by hydrolysis of the ester linkage [226]. The replacement of the ester linkage by a more hydrolytically stable linkage will likely result in a lipopeptide less susceptible to ring hydrolysis.

2.4 Tailoring enzyme—Change conformation A variety of tailoring enzyme are involved in lipopeptide biosynthesis, as epimerase and methyltransferase [9]. Deleting or adding tailoring enzymes could lead to the biosynthesis of new lipopeptides (Fig. 1). Instead of modifying the composition of amino acids in the final lipopeptide product, it is possible to keep the same amino acids but change their stereoisomery, thus the structure backbone of the lipopeptide, by deleting the epimerization domain, leading to the incorporation of a L-amino acid instead of a D-amino acid [227]. In A21978C lipopeptides, a substitution of L-Asn2 to D-Asn2 changed the chemical properties and increased the minimal inhibitory concentration (MIC) against S. aureus, Enterococcus faecium, and Enterococcus faecalis by 10-fold compared to daptomycin [228,229]. In Streptomyces fradiae NRRL 18160, the deletion of lptI, a gene encoding a methyltransferase involved in the conversion of Glu12 to 3mGlu12, to generate A54145 lipopeptide with Glu12, enhanced the production titer of the lipopeptide, as Glu12 is more easily inserted into the peptide backbone than 3mGlu12 [230]. However, the deletion of the gene dptI coding a methytransferase in Streptomyces roseosporus showed that the mutant producing a daptomycin with a substitution of 3mGlu12 by Glu12 has a reduced antibiotic activity [136].

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2.5 Remodeling/assembling of biosynthetic gene clusters One way to create new nonnatural lipopeptides is by remodeling genes involved in the biosynthesis of different NRPS (Fig. 1). Myxochromides are lipopeptides produced by hybrid megasynthetase PKS/ NRPS and are produced by myxobacteria [231]. In order to produce new myxochromides molecules, different biosynthetic gene clusters of myxochromides were mixed and assembled by using type II endonuclease strategies. They formed more than 30 different artificial myxochromides biosynthetic gene clusters encoding multifunctional PKS-NRPS megasynthetases, each of around 30 kb and expressed in Myxococcus xanthus DK1622 as a host. This remodeling led to the inactivation, deletion, duplication of catalytic domains, and the production of new lipopeptides [232]. Another example of this strategy is that of NRPS genes encoding subunits from two related lipopeptide biosynthetic pathways that were cloned into S. roseosporus to create a daptomycin hybrid NRPS system [233]. In addition, two biosynthetic gene clusters of daptomycin and A54145 were combined to create new lipopeptides structures [228,234]. The authors showed that bidomain exchange, module exchange, and multidomain exchanges are possible, leading to the production of 40 daptomycin-related lipopeptides with some of them displaying different properties. The antibacterial activities of these new lipopeptides against S. aureus were determined with or without the addition of 1% bovine surfactant (used to mimic pulmonary biosurfactant, a surface-active complex of phospholipids, and proteins formed by type II alveolar cells), known to decrease antibacterial activity of daptomycin and thus to mimic the in vivo situation and help to identify candidates that might be efficacious against S. pneumoniae pulmonary infection. They observed that the in vitro antibacterial activities varied by 256-fold when the core peptides of daptomycin and A54145 were modified [235].

2.6 Structure modification by modification of the FA moiety The FA moiety of CDA was modified in order to increase its biological activity. New analogues of CDA were produced by introducing alternative FA groups using the mutasynthesis method to mutate serine into an alanine residue in the module 1 PCP domain of CdaPS1, preventing the phosphopantetheinylation of the PCP domain [236]. This mutation led to the incorporation of different N-acylL-serinyl N-acetylcysteamine thioester analogues, leading to the production of CDA products with pentanoyls as well as hexanoyl side chains [236].

3. Improving the homologous production of lipopeptides Although it is possible to produce new lipopeptides, the quantity produced is frequently too low for an industrial application. In addition, the production of lipopeptides in wild-type strains are sometimes low, e.g., in shake-flask cultures, the production is around 30 mg/L for daptomycin in S. roseosporus [237], 17 mg/L for mycosubtilin in B. subtilis [238], and 8 mg/L for myxochromide in Stigmatella aurantiaca [239]. Therefore it is necessary to improve the production of lipopeptides in the natural hosts [240]. This part will focus on the improvement of the production of lipopeptides at different biosynthesis levels in natural or modified hosts.

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3.1 Targeting gene regulation Production of lipopeptides relies on many factors, therefore deleting negative and/or overexpressing positive, direct, or indirect, regulators of transcription of genes involved in the biosynthesis of lipopeptides could increase the production of these molecules (Fig. 2). Many studies have already identified negative/positive regulators involved in lipopeptide production. For example, in Bacillus, the expression of srfA is under the control of a complex cascade of regulation linked to the quorum sensing metabolism [241] and involving pheromones such as ComX and Phr. Phr peptides inhibit the activity of RapC, RapF, and RapK, negative regulators of srfA. ComX activates ComP that will then phosphorylate ComA that will directly initiates the transcription of both operon srfA and myc. Additional transcription factors, DegU and PepR, act as positive regulators of srfA transcription. CodY represses the transcription of srfA when a high concentration of Ile, Leu, and Val is present in the cells. Likewise, AbrB and the RNA polymerase-binding protein Spx inhibit srfA transcription [242–246]. DegQ positively controls the regulation of fengycin, plipaspatin, and iturin A [247,248]. Finally, myc operon is also influenced by the sigma H factor, Spo0H [242]. Other examples include the transcription of kurstakin, that is, indirectly and negatively affected by Spo0A and could also be affected by a small antisense RNA or a riboswitch [110]. Details about other regulator factors for lipopeptide biosynthesis in Bacillus and Pseudomonas can be found in the review of Roongsawang, Washio, and Morikawa [242]. Likewise, for daptomycin, description of regulators can be found in the reviews of Mao, Luo, and Li and Mao et al. [249,250].

3.2 Increase in transcription Another common strategy to increase the transcription of NRPS biosynthesis is to directly change the promoter of the operon (Fig. 2). It has been observed that transcription is a critical and limiting step in the production of lipopeptides [240,248]. Several studies exchanged the promoter of the operon, by stronger inducer-specific or constitutive promoters, to increase the transcription level of biosynthetic gene clusters and thus the lipopeptide production. Often these genetic modifications are coupled with an optimized fermentation process [238,240,241,248,251–254]. For daptomycin, low transcription of the biosynthetic gene cluster is responsible for a low production, and, by using top-down synthetic biology approach, Ji et al. increased the transcription level and daptomycin production. This strategy relied on codon reprogramming to eliminate large sequence repeats present in the biosynthetic gene cluster of daptomycin and promoter engineering. These modifications led to a 2,300% improvement in total lipopeptide titers compared to the wild-type strain [255].

3.3 Targeting the FA metabolism Lipopeptides are composed of FA chains, therefore increasing FA biosynthesis could lead to an increase in lipopeptide biosynthesis [256] (Fig. 2). Different metabolic engineering approaches were applied to improve FA biosynthesis by: (1) enhancing the branched-chain α-ketoacyl CoA supply, (2) enhancing malonyl-ACP synthesis, (3) overexpressing the whole FA synthase complex, and (4) diverting precursors pathway used for cell growth toward surfactin synthesis by enhancing srfA transcription [209,254,257]. These modifications resulted in a significant increase in surfactin production. Recently, the production of fengycin was improved by overexpressing acs, birA, and accACD, which are relevant genes of the malonyl-CoA synthesis pathway and a limiting precursor of the FA biosynthesis [256].

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FIG. 2 Representation of the complete mechanism involved in the biosynthesis of lipopeptides in a natural producer. The import of precursors as carbon source is used by the metabolic pathway which then synthesizes fatty acids (FA) and amino acids (AA) and possess genes of regulation, which downregulate or upregulate the transcription of the operon of biosynthetic gene clusters. Then, NRPS genes are transcribed and translated into apoproteins. The activation of thiolation domains by the phosphopenteteinate transferase encoded by sfp gene transformed the apo-protein in holo-protein able to synthetize the lipopeptide. Then, specifics transporters are used by the cells to export lipopeptides outside the cells. In certain conditions, lipopeptides may undergo degradation and FA and AA then may be used by the cells.

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3.4 Targeting the amino acid metabolism With the same logic as mentioned previously, increasing precursors of amino acids could lead to an increase of lipopeptide biosynthesis. It can be carried out either by improving anabolic pathways or reducing catabolic pathways or by increasing positive gene regulators or decreasing negative gene regulators that would influence the intracellular pool of amino acids in the cell [209,258]. For example, the deletion of murC, yrpC, and racE, negative regulators involved in the metabolism of glutamate, increased the production of surfactin [257]. Moreover, amino acids can be incorporated into cells via transporters, which could be overexpressed as well (Fig. 2).

3.5 Targeting the genome A new strategy emerged to improve lipopeptide production by using a whole secondary metabolic engineering approach and some experiments have showed promising results. Surfactin and plipastatin biosynthesis share several precursors such as those involved in FA chain as well as glutamic acid. Each biosynthesis requires high level of ATP for the activation of different amino acids to be incorporated into the peptide. The strategy used to improve surfactin production was to delete ppsA gene (involved in plipastatin production), to increase the availability of precursors for the biosynthesis of surfactin [241]. Therefore the knock-out of competitive pathways redirect the precursors incorporated/produced toward the biosynthesis of wanted lipopeptides. Same results were obtained by deletion of the pathway involved in biofilm formation-related genes and NRPSs/PKS pathways (3.8% of the total genome of B. subtilis 168), which increased the surfactin titer by 3.3-fold [254]. A reduction of 4.18% of the genome of B. amyloliquefaciens LL3, with disruption of the iturin and fengycin biosynthesis cluster, led to an increase of surfactin production [259]. However, the reduction of 10% of the genome of B. subtilis, deleting genes synthesizing the plipastatin, the antibiotic bacilysin, toxins, and prophages, as well as genes involved in sporulation, led to a decrease of lipopeptide production [260]. Finally, repetitive random mutation [261] and genome shuffling [262] were conducted on S. roseosporus, and these modifications improved daptomycin production.

3.6 Increase in transporters and toxicity resistance genes Following their biosynthesis, lipopeptides are excreted outside the cells. Various transporters have been identified in Bacillus such as YerP, YcxA, KrsE, AcrB, and their overexpression increased surfactin production [254,263,264] (Fig. 2). Moreover, different lipopeptide resistance genes have been identified [265]. The combination of overexpression of transporters and lipopeptide resistance proteins increased surfactin production [254]. The biosynthesis of daptomycin is initiated by the condensation of decanoic acid and the N-terminal L-tryptophan. Thus the addition of decanoic acid in the culture is essential for the daptomycin synthesis, but this precursor is toxic for the cells. Increasing resistance to decanoic acid improved its production [266].

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3.7 Degradation Some studies have observed a degradation of surfactin in Bacillus species. Two main hypotheses have been proposed: (a) in some natural producer of surfactin, when a certain level of production is reached, a degradation occurs to avoid a decrease of growth rate or cellular death and (b) after carbon depletion, surfactin can be used as carbon and energy sources. However, no specific proteases or lactonases have been yet identified [267,268]. With these observations, degradation of lipopeptides should be further investigated to increase their production.

4. Heterologous production Although it is possible to improve the lipopeptide production in some natural host, the yield is frequently low and not all native hosts are suitable for genetic manipulations or large-scale growth because of their complex cultivation [269]. To overcome the problems encountered with native host, a heterologous production may be considered. Heterologous recombinant expression of lipopeptides has already been carried out and showed a better production than observed in the native host, e.g., daptomycin production in Streptomyces lividans, but the enhancement was not enough for an industrial process [270]. For the optimization of heterologous production, three main steps must be considered: the complete comprehension of the biosynthesis mechanism, the choice of the host strain, and the cloning strategies.

4.1 Deciphering the complete biosynthesis mechanism The first step of heterologous production is the full identification of genes, their interactions, and understanding the whole complex mechanism of NRPS biosynthesis, from the first substrate/precursor to the complete product formed and its localization inside or outside the cells. Molecules from secondary metabolisms are tightly regulated at different levels and the factors involved in these regulations are multiple [271,272]. As seen in the previous sections, various precursors from primary (enough incorporation of carbon, nitrogen, phosphate, etc.) and secondary metabolisms are involved in the biosynthesis of lipopeptides; thus several diverse positive/negative regulators or metabolisms such as quorum sensing may influence this biosynthesis [241]. During the biosynthesis, different posttranslational modifications are added to synthetize the functional produced lipopeptide by enzymes such as methytransferase, phosphopantetheinyl-transferase, etc. Finally, specific transporters (such as YerP) are involved in the export of the lipopeptides. Therefore it is crucial to fully understand the complete mechanism of biosynthesis of NRPS.

4.2 Choice of the host strain To produce lipopeptides in a host strain, the producer must have metabolic capabilities for precursors, cofactor incorporation and synthesis, and should not be affected by the molecules produced. In this case, the choice of the host for heterologous lipopeptide production is critical. The strain has to present desirable characteristics such as fast growth rate, high cell density and simple cultivation, very good knowledge of the entire genome, easily genetically manipulable, tolerance to expression of foreign proteins, well understood metabolic pathways, and codon compatibility [273] (Fig. 3). If the host does not have naturally these characteristics, these capabilities must be implemented.

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FIG. 3 Critical steps for heterologous production of NRPS molecules. These include the choice of the strain, the de novo synthesis and codon optimization, and finally the expression of the NRPS biosynthetic gene clusters via a replicative plasmid or after its integration inside the genome.

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Insertion of one biosynthesis gene cluster of NRPS may be not sufficient because the complete synthesis and activities require other proteins such as transporters for incorporation of precursors or excretion of the produced molecules. Precursors can be added by feeding the strains or by cloning transporters for specific precursors or increasing the efficiency of already present transporters. Studies have already shown that increasing metabolisms involved in carbon, nitrogen, and phosphorous enhanced bacitracin production in B. licheniformis [274]. Closely related strains may facilitate the production of lipopeptides. In some examples, successful heterologous productions were achieved. A 42-kb gene containing the complete 38-kb iturin A operon from B. subtilis RB14, as well as the sfp gene, was transferred into nonproducer B. subtilis 168, leading to the production of iturin A in the natural nonproducer. The insertion of the pleiotropic regulator degQ into B. subtilis 168, in which the gene degQ is mutated and non-functional, further increased the production of iturin A, to reach half of the one obtained in the natural producer B. subtilis RB14. The authors showed that to produce iturin A, the complete operon is the only cluster required for the conversion of a non-iturin A producer into an iturin A producer, with the essential presence of the sfp gene. Moreover, no additional regulators, self-resistance, or efflux pump genes were added, showing that the heterologous host naturally possesses some genes to produce iturin A [275]. On the contrary, phylogenetically distant strains may lead to the production of different molecules. The gene encoding serrawettin synthetase swrW from S. marcescens DSM12481 was cloned into an IPTG-induced expression plasmid and transformed into E. coli. This heterologous host presented a 20-fold increase of serrawettin W1 compared to the natural host. However, the serrawettin was produced with a FA chain length of 10 carbons instead of 13 and 14 carbons, suggesting that E. coli used a different pathway involved in FA synthesis [276]. With phylogenetically distant bacteria, some other steps must be taken into account. For example, the presence of protease can degrade heterologous proteins; thus to overcome this problem, specific proteases can be deleted [277]. As another example, protein/enzyme activities can be low or inexistant due to a partial or complete misfolding. To overcome this problem, chaperones can be coexpressed [278]. Compared to proteins which are constituted of the 20 proteinogenic amino acids residues, NRPSdependent molecules can mobilize more than 500 different monomers [3]. Thus if the heterologous host cannot synthesize some of them, they must be incorporated through feeding, or metabolic pathways must be changed, to allow the synthesis of these monomers. Moreover, it can be essential to express specific tailoring enzymes or transporters in the heterologous host. For example, enniatin molecules, encoded by the esyn gene, from Fusarium oxysporum, and cloned into bacteria were successfully exported outside the cells in B. subtilis but accumulated intracellularly in E. coli [279]. In addition, the production of heterologous molecules may be toxic for the surrogate bacteria. This toxicity can be due to the feeding of precursors (e.g., decanoic acid) or the molecules produced. To solve this, different solutions exist, such as expressing resistance proteins [254,266] or expressing NRPS under the control of inducible promoters to control the period of production [277]. Lipopeptide synthesis, in addition to transcription and translation, requires posttranslational modifications. One of the most studied and crucial posttranslational gene is sfp that encodes a phosphopantetheinyl-transferase, essential for lipopeptide production, as it mediates the conversion of the NRPS from the apo form to the holo form of the PCP domain [254,280]. Sfp is also involved in PKS synthesis and FA synthesis [281]. This gene must be present or incorporated into the recipient strain.

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4.3 Cloning strategy To express the NRPS into the host, the first step is to clone the biosynthetic gene cluster, which can be over 40 kilobases (Fig. 3). Genes can be obtained from the strain by polymerase chain reaction, but in that case codon optimization is not possible. However, contrary to bacteria, to take genes from eukaryotes, it is necessary to target the mRNA, intron-free open reading frame, and then perform a reverse transcription. Other techniques can be found in the review of Vassaux et al. [277]. Otherwise, genes may be de novo synthesized and codon optimized (Fig. 3). In the case of heterologous production, the transcription of the gene of interest can be low due to a lack or bad codon optimization. If the natural producer and host are phylogenetically distant, the different codon usage can slow down the expression of the NRPS. Phylogenetically, closer strains share common codon usage, increasing the NRPS synthesis efficiency [282]. By using codon optimization, it is possible to produce NRPS-dependent molecules from two phylogenetically distant strains, even between prokaryotes and eukaryotes. Studies have shown that heterologous production from prokaryotes to eukaryotes or vice versa was possible despite their genomic differences. As an example, the heterologous expression of the NRPS gene bpsA that converts glutamine into indigoidine, a non-lipopeptide NRPS blue pigment, from Streptomyces lavendulae into Saccharomyces cerevisiae was successful. The gene was codonoptimized and cloned into the genome and a maximum titer of 980 mg/L of indigoidine was obtained [283]. Additionally, the cloning of the gene esyn, encoding the enniatin synthetase, from F. oxysporum into B. subtilis lead to the production of enniatin [279]. For the cloning of large DNA sequences inside a plasmid, different vectors can be used such as cosmids or fosmids for the cloning of 30–35 kb regions or bacterial artificial chromosomes that can contain over 100 kb [284]. To insert DNA sequences into a plasmid, various methods can be used such as the Gibson technique [285], overlap extension PCR technique [286], and restriction/ligation method [279]. When many genes are cloned into a replicative plasmid, allowing a stable expression, the cloning strategy elected may influence the final production titer. Cyclic dipeptide D-Phe-Pro-diketopiperazine produced by the TycA/TycB1 system, from B. brevis, was heterologously expressed in E. coli, with different strategies. The two genes TycA and TycB1 were (a) cloned in one plasmid where genes are under control of same promotor; (b) fused into one open reading frame, under the control of IPTG-inducible T5 promoter; and (c) cloned into two different plasmids with each respective gene under the control of the IPTG-inducible promoters lac and trc. The best strategy was found to be using the two plasmids system, followed by the one plasmid strategy and then the fusion strategy. With the two plasmids strategy, an increase of 400% was observed and the production titer reached 9 mg/L [287]. Moreover, genes may be cloned and expressed from a replicative plasmid or can be integrated in the heterologous host chromosome. For example, the cloning of the complete myxochromide biosynthetic gene cluster from S. aurantiaca and its insertion inside the chromosome GT2 of the heterologous strain Corallococcus macrosporus GT-2 was successful. After promoter exchange, the heterologous production was higher than the natural producer (600 vs. 8 mg/L) [269]. Another study has investigated the production of cyclodepsipeptide enniatin in B. subtilis after cloning of the gene esyn inside a multicopy plasmid versus single chromosomal integration inside the gene amyE. The highest production of enniatin was observed with the use of the multicopy plasmid [279].

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Finally, if several genetic modifications must be done to produce lipopeptides in a heterologous host, it is essential to use marker-less methods such as the Cre-Lox recombination [288]. In conclusion, heterologous production seems to be promising, but the final molecules produced, and their yield is not a certainty. To achieve the best results possible, a whole comprehension of the mechanism involved in the biosynthesis of the molecule, the choice of host strain, the genetic modifications, and the strategy of cloning must be considered as these steps will influence the final production.

5. Conclusion Lipopeptides are a class of biosurfactant molecules with a very high biodiversity. Indeed, they can vary in the size, isoform, and the saturation of their FA chain as well as in the size, composition, and sequence of its peptide moiety leading to several thousand of biomolecules already described in nature. This list is for sure nonexhaustive. Some lipopeptides are encoded by cryptic genes [289,290] or by microorganisms which are not yet described. The modular character of their mode of biosynthesis which can be modified at the level of domains, modules, operons, or clusters of genes is a wonderful tool to further increase this biodiversity. The main challenge in the future is to get cell factories able to produce highly surface-active lipopeptides in high yields.

Funding This work was supported by the European ERACoBioTech program (BestBioSurf project), the European INTERREG Va SmartBioControl project, the MaisMisVal project (Walloon Region, Agriculture department DG03), and the FNRS-funded SURFACOVID project.

Acknowledgments We thank Andrew Zicler for his help in the figure design.

Conflict of interest PJ is a co-founder of Lipofabrik and Lipofabrik Belgium and a member of the scientific advisory board of both companies.

References [1] S€ussmuth RD, Mainz A. Nonribosomal peptide synthesis—principles and prospects reviews. Angew Chem Int Ed 2017;56:3770–823. [2] Walsh CT, O’Brien RV, Kholsa C. Nonproteinogenic amino acid building blocks for nonribosomal peptide and hybrid polyketide scaffolds. Angew Chem Int Ed Engl 2013;52(28):7098–124.

References

233

[3] Caboche S, Lecle`re V, Pupin M, Kucherov G, Jacques P. Diversity of monomers in nonribosomal peptides: towards the prediction of origin and biological activity. J Bacteriol 2010;192(19):5143–50. [4] Mootz HD, Finking R, Marahiel MA. 40 -Phosphopantetheine transfer in primary and secondary metabolism of Bacillus subtilis. J Biol Chem 2001;276(40):37289–98. [5] Quadri LEN, Weinreb PH, Lei M, Nakano MM, Zuber P, Walsh CT. Characterization of Sfp, a Bacillus subtilis phosphopantetheinyl transferase for peptidyl carder protein domains in peptide synthetases. Biochemistry 1998;37(6):1585–95. [6] Bloudoff K, Schmeing TM. Structural and functional aspects of the nonribosomal peptide synthetase condensation domain superfamily: discovery, dissection and diversity. Biochim Biophys Acta Proteins Proteomics 2017;1865:1587–604. Elsevier B.V. [7] Rausch C, Hoof I, Weber T, Wohlleben W, Huson DH. Phylogenetic analysis of condensation domains in NRPS sheds light on their functional evolution. BMC Evol Biol 2007;7(78). [8] Kotowska M, Pawlik K. Roles of type II thioesterases and their application for secondary metabolite yield improvement. Appl Microbiol Biotechnol 2014;98:7735–46. [9] Walsh CT, Chen H, Keating TA, Hubbard BK, Losey HC, Luo L, et al. Tailoring enzymes that modify nonribosomal peptides during and after chain elongation on NRPS assembly lines. Curr Opin Chem Biol 2001;5:525–34. [10] Aron ZD, Dorrestein PC, Blackhall JR, Kelleher NL, Walsh CT. Characterization of a new tailoring domain in polyketide biogenesis: the amine transferase domain of MycA in the mycosubtilin gene cluster. J Am Chem Soc 2005;127:14986–7. [11] Hansen DB, Bumpus SB, Aron ZD, Kelleher NL, Walsh CT. The leading module of mycosubtilin: an adenylation domain with fatty acid selectivity. J Am Chem Soc 2007;129:6366–7. [12] Bonmatin JM, Laprevote O, Peypoux F. Diversity among microbial cyclic lipopeptides: iturins and surfactins. activity-structure relationships to design new bioactive agents. Comb Chem High Throughput Screen 2003;6:541–56. [13] Thimon L, Peypoux F, Marget-Dana R, Michel G. Surface-active properties of antifungal lipopeptides produced by Bacillus subtilis. J Am Oil Chem Soc 1992;69(1):92–3. [14] Jacques P. Surfactin and other lipopeptides from Bacillus spp. In: Sobero´n-Cha´vez G, editor. Biosurfactants. microbiology monographs, vol. 20. Berlin, Heidelberg: Springer; 2011. p. 57–91. [15] Raaijmakers JM, de Bruijn I, Nybroe O, Ongena M. Natural functions of lipopeptides from Bacillus and Pseudomonas: more than surfactants and antibiotics. FEMS Microbiol Rev 2010;34(6):1037–62. € [16] Kracht M, Rokos H, Ozel M, Kowall M, Pauli G, Vater J. Antiviral and hemolytic activities of surfactin isoforms and their methyl ester derivatives. J Antibiot (Tokyo) 1999;52(7):613–9. [17] Vollenbroich D, Muhsin O, Vater J, Kamp RM, Pauli G. Mechanism of inactivation of enveloped viruses by the biosurfactant surfactin from Bacillus subtilis. Biologicals 1997;25:289–97. [18] Liu X, Tao X, Zou A, Yang S, Zhang L, Mu B. Effect of themicrobial lipopeptide on tumor cell lines: apoptosis induced by disturbing the fatty acid composition of cell membrane. Protein Cell 2010;1(6):584–94. [19] Park SY, Kim YH. Surfactin inhibits immunostimulatory function of macrophages through blocking NK-κ B, MAPK and Akt pathway. Int Immunopharmacol 2009;9(7–8):886–93. [20] Vanittanakom N, Loeffler W, Koch U, Jung G. Fengycin—a novel antifungal lipopeptide antibiotic produced by Bacillus subtilis F-29-3. J Antibiot (Tokyo) 1986;39(7):888–901. [21] Shakerifard P, Gancel F, Jacques P, Faille C. Effect of different Bacillus subtilis lipopeptides on surface hydrophobicity and adhesion of Bacillus cereus 98/4 spores to stainless steel and Teflon. Biofouling 2009;25(6):533–41. [22] Cheng W, Feng YQ, Ren J, Jing D, Wang C. Anti-tumor role of Bacillus subtilis fmbJ-derived fengycin on human colon cancer HT29 cell line. Neoplasma 2016;63(2):215–22. [23] Kang BR, Park JS, Jung WJ. Antiviral activity by lecithin-induced fengycin lipopeptides as a potent key substrate against Cucumber mosaic virus. Microb Pathog 2021;155, 104910.

234

Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

[24] Besson F, Peypoux F, Michel G, Delcambe L. Characterization of iturin a in antibiotics from various strains of Bacillus subtilis. J Antibiot (Tokyo) 1976;29(10):1043–9. [25] Dey G, Bharti R, Dhanarajan G, Das S, Dey KK, Kumar BNP, et al. Marine lipopeptide Iturin A inhibits Akt mediated GSK3β and FoxO3a signaling and triggers apoptosis in breast cancer. Sci Rep 2015;5 (January):1–14. [26] Zhao H, Li J, Zhang Y, Lei S, Zhao X, Shao D, et al. Potential of iturins as functional agents: safe, probiotic, and cytotoxic to cancer cells. Food Funct 2018;9:5580–7. [27] Razafindralambo H. Contribution a` l’etude des proprietes tensioactives de lipopeptides de Bacillus subtilis. ULie`ge. GxABT - Lie`ge Universite. Gembloux Agro-Bio Tech.; 1996 [Unpublished doctoral thesis]. [28] Hathout Y, Ho YP, Ryzhov V, Demirev P, Fenselau C. Kurstakins: a new class of lipopeptides isolated from Bacillus thuringiensis. J Nat Prod 2000;63(11):1492–6. [29] Diallo MM, Vural C, Şahar U, Ozdemir G. Kurstakin molecules facilitate diesel oil assimilation by Acinetobacter haemolyticus strain 2SA through overexpression of alkane hydroxylase genes. Environ Technol 2021;42(13):2031–45. [30] Batrakov SG, Rodionova TA, Esipov SE, Polyakov NB, Sheichenko VI, Shekhovtsova NV, et al. A novel lipopeptide, an inhibitor of bacterial adhesion, from the thermophilic and halotolerant subsurface Bacillus licheniformis strain 603. Biochim Biophys Acta Mol Cell Biol Lipids 2003;1634(3):107–15. [31] Lee SC, Kim SH, Park IH, Chung SY, Choi YL. Isolation and structural analysis of bamylocin A, novel lipopeptide from Bacillus amyloliquefaciens LP03 having antagonistic and crude oil-emulsifying activity. Arch Microbiol 2007;188(4):307–12. [32] Biria D, Maghsoudi E, Roostaazad R, Dadafarin H, Lotfi SS, Amoozegar MA. Purification and characterization of a novel biosurfactant produced by Bacillus licheniformis MS3. World J Microbiol Biotechnol 2010;26(5):871–8. [33] Poirel L, Jayol A, Nordmann P. Polymyxins: antibacterial activity, susceptibility testing, and resistance mechanisms encoded by plasmids or chromosomes. Clin Microbiol Infect 2017;30(2):557–96. [34] Few AV, Schulman JH. The absorption of polymyxin E by bacteria and bacterial cell walls and its bactericidal action. J Gen Microbiol 1953;9(3):454–66. [35] Seghal Kiran G, Anto Thomas T, Selvin J, Sabarathnam B, Lipton AP. Optimization and characterization of a new lipopeptide biosurfactant produced by marine Brevibacterium aureum MSA13 in solid state culture. Bioresour Technol 2010;101:2389–96. [36] Senthil Balan S, Ganesh Kumar C, Jayalakshmi S. Aneurinifactin, a new lipopeptide biosurfactant produced by a marine Aneurinibacillus aneurinilyticus SBP-11 isolated from Gulf of Mannar: purification, characterization and its biological evaluation. Microbiol Res 2017;194:1–9. [37] Eddouaouda K, Mnif S, Badis A, Ben Younes S, Cherif S, Ferhat S, et al. Characterization of a novel biosurfactant produced by Staphylococcus sp. strain 1E with potential application on hydrocarbon bioremediation. J Basic Microbiol 2011;51:1–11. [38] Dalili D, Amini M, Faramarzi MA, Fazeli MR, Khoshayand MR, Samadi N. Isolation and structural characterization of Coryxin, a novel cyclic lipopeptide from Corynebacterium xerosis NS5 having emulsifying and anti-biofilm activity. Colloids Surf B Biointerfaces 2015;135:425–32. [39] Tally FP, Zeckel M, Wasilewski MM, Carini C, Berman CL, Drusano GL, et al. Daptomycin: a novel agent for Gram-positive infections. Expert Opin Investig Drugs 1999;8(8):1223–38. [40] Juhaniewicz-De¸binska J, Dziubak D, Se¸k S. Physicochemical characterization of daptomycin interaction with negatively charged lipid membranes. Langmuir 2020;36:5324–35. [41] Kirkham S, Castelletto V, Hamley IW, Inoue K, Rambo R, Reza M, et al. Self-assembly of the cyclic lipopeptide daptomycin: spherical micelle formation does not depend on the presence of calcium chloride. ChemPhysChem 2016;17:2118–22. [42] Peng F, Wang Y, Sun F, Liu Z, Lai Q, Shao Z. A novel lipopeptide produced by a Pacific Ocean deep-sea bacterium, Rhodococcus sp. TW53. J Appl Microbiol 2008;105(3):698–705.

References

235

[43] Habib S, Ahmad SA, Johari WLW, Shukor MYA, Alias SA, Smykla J, et al. Production of lipopeptide biosurfactant by a hydrocarbon-degrading Antarctic Rhodococcus. Int J Mol Sci 2020;21(6138). [44] Janek T, Łukaszewicz M, Rezanka T, Krasowska A. Isolation and characterization of two new lipopeptide biosurfactants produced by Pseudomonas fluorescens BD5 isolated from water from the Arctic Archipelago of Svalbard. Bioresour Technol 2010;101(15):6118–23. [45] Janek T, Krasowska A, Radwanska A, Łukaszewicz M. Lipopeptide biosurfactant pseudofactin II induced apoptosis of melanoma A 375 cells by specific interaction with the plasma membrane. PLoS One 2013;8 (3):1–9. [46] Biniarz P, Baranowska G, Feder-Kubis J, Krasowska A. The lipopeptides pseudofactin II and surfactin effectively decrease Candida albicans adhesion and hydrophobicity. Antonie Van Leeuwenhoek 2015;108(2):343–53. [47] Janek T, Łukaszewicz M, Krasowska A. Antiadhesive activity of the biosurfactant pseudofactin II secreted by the Arctic bacterium Pseudomonas fluorescens BD5. BMC Microbiol 2012;12(24). [48] Groupe V, Pugh LH, Weiss D, Kochi M. Observations on antiviral activity of viscosin. Proc Soc Exp Biol Med 1951;78(1):354–8. [49] Gerard J, Lloyd R, Barsby T, Haden P, Kelly MT, Andersen RJ. Massetolides A-H, antimycobacterial cyclic depsipeptides produced by two pseudomonads isolated from marine habitats. J Nat Prod 1997;60(3):223–9. [50] Saini HS, Barragan-Huerta BE, Lebron-Paler A, Pemberton JE, Vazquez RR, Burns AM, et al. Efficient purification of the biosurfactant viscosin from Pseudomonas libanensis Strain M9-3 and its physicochemical and biological properties. J Nat Prod 2008;71:1011–5. [51] Mortishire-Smith RJ, Nutkins JC, Packman LC, Brodey CL, Rainey PB, Johnstone K, et al. Determination of the structure of an extracellular peptide produced by the mushroom saprotroph Pseudomonas reactans. Tetrahedron 1991;47(22):3645–54. [52] Laycock MV, Hildebrand PD, Thibault P, Walter JA, Wright JLC. Viscosin, a potent peptidolipid biosurfactant and phytopathogenic mediator produced by a pectolytic strain of Pseudomonas fluorescens. J Agric Food Chem 1991;39(3):483–9. [53] De Bruijn I, De Kock MJD, De Waard P, Van Beek TA, Raaijmakers JM. Massetolide A biosynthesis in Pseudomonas fluorescens. J Bacteriol 2008;190(8):2777–89. [54] Gross H, Stockwell VO, Henkels MD, Nowak-Thompson B, Loper JE, Gerwick WH. The genomisotopic approach: a systematic method to isolate products of orphan biosynthetic gene clusters. Chem Biol 2007;14:53–63. [55] Jang JY, Yang SY, Kim YC, Lee CW, Park MS, Kim JC, et al. Identification of orfamide A as an insecticidal metabolite produced by Pseudomonas protegens F6. J Agric Food Chem 2013;61:6786–91. [56] Sorensen D, Nielsen TH, Christophersen C, Sorensen J, Gajhede M. Cyclic lipoundecapeptide amphisin from Pseudomonas sp. strain DSS73. Acta Crystallogr C 2001;57:1123–4. [57] Morikawa M, Daido H, Takao T, Murata S, Shimonishi Y, Imanaka T. A new lipopeptide biosurfactant produced by Arthrobacter sp. Strain MIS38. J Bacteriol 1993;175(20):6459–66. [58] Kuiper I, Lagendijk EL, Pickford R, Derrick JP, Lamers GEM, Thomas-Oates JE, et al. Characterization of two Pseudomonas putida lipopeptide biosurfactants, putisolvin I and II, which inhibit biofilm formation and break down existing biofilm. Mol Microbiol 2004;51(1):97–113. [59] Dubern JF, Coppoolse ER, Stiekema WJ, Bloemberg GV. Genetic and functional characterization of the gene cluster directing the biosynthesis of putisolvin I and II in Pseudomonas putida strain PCL1445. Microbiology 2008;154:2070–83. [60] Vallet-Gely I, Novikov A, Augusto L, Liehl P, Bolbach G, Pechy-Tarr M, et al. Association of hemolytic activity of Pseudomonas entomophila, a versatile soil bacterium, with cyclic lipopeptide production. Appl Environ Microbiol 2010;76(3):910–21. [61] Li W, Rokni-Zadeh H, De Vleeschouwer M, Ghequire MGK, Sinnaeve D, Xie GL, et al. The antimicrobial compound xantholysin defines a new group of Pseudomonas cyclic lipopeptides. PLoS One 2013;8(5).

236

Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

[62] Shimura S, Ishima M, Nakajima S, Fujii T, Himeno N, Ikeda K, et al. Total synthesis and anti-hepatitis C virus activity of MA026. J Am Chem Soc 2013;135(50):18949–56. [63] Jourdan F, Lazzaroni S, Lopez Mendez B, Lo Cantore P, de Julio M, Amodeo P, et al. A left-handed alphahelix containing both L- and D-amino acids: the solution structure of the antimicrobial lipodepsipeptide tolaasin. Proteins Struct Funct Genet 2003;52:534–43. [64] Hutchison ML, Johnstone K. Evidence for the involvement of the surface-active properties of the extracellular toxin tolaasin in the manifestation of brown blotch disease symptoms by Pseudomonas tolaasii on Agaricus bisporus. Physiol Mol Plant Pathol 1993;42:373–84. [65] Bassarello C, Lazzaroni S, Bifulco G, Lo Cantore P, Iacobellis NS, Riccio R, et al. Tolaasins A-E, five new lipodepsipeptides produced by Pseudomonas tolaasii. J Nat Prod 2004;67(5):811–6. [66] Segre A, Bachmann RC, Ballio A, Bossa F, Grgurina I, Iacobellis NS, et al. The structure of syringomycins A1, E and G. FEBS Lett 1989;255(1):27–31. [67] Hutchison ML, Tester MA, Gross DC. Role of biosurfactant and ion channel-forming activities of syringomycin in transmembrane ion flux: a model for the mechanism of action in the plant-pathogen interaction. Mol Plant Microbe Interact 1994;8(4):610–20. [68] Ballio A, Barra D, Bossa F, Collina A, Grgurina I, Marino G, et al. Syringopeptins, new phytotoxic lipodepsipeptides of Pseudomonas syringae pv. syringae. FEBS Lett 1991;291(1):109–12. [69] Grgurina I, Bensaci M, Pocsfalvi G, Mannina L, Cruciani O, Fiore A, et al. Novel cyclic lipodepsipeptide from Pseudomonas syringae pv. lachrymans Strain 508 and syringopeptin antimicrobial activities. Antimicrob Agents Chemother 2005;49(12):5037–45. [70] Hutchison ML, Gross DC. Lipopeptide phytotoxins produced by Pseudomonas syringae pv. syringae: comparison of the biosurfactant and ion channel-forming activities of syringopeptin and syringomycin. Mol Plant Microbe Interact 1997;10(3):347–54. [71] Lavermicocca P, Sante Iacobellis N, Simmaco M, Graniti A. Biological properties and spectrum of activity of Pseudomonas syringae pv. syringae toxins. Physiol Mol Plant Pathol 1997;50(2):129–40. [72] Berti AD, Greve NJ, Christensen QH, Thomas MG. Identification of a biosynthetic gene cluster and the six associated lipopeptides involved in swarming motility of Pseudomonas syringae pv. tomato DC3000. J Bacteriol 2007;189(17):6312–23. [73] Pauwelyn E, Huang CJ, Ongena M, Lecle`re V, Jacques P, Bleyaert P, et al. New linear lipopeptides produced by pseudomonas cichorii SF1-54 are involved in virulence, swarming motility, and biofilm formation. Mol Plant Microbe Interact 2013;26(5):585–98. [74] Thongkongkaew T, Ding W, Bratovanov E, Oueis E, Garcia-Altares M, Zaburannyi N, et al. Two types of threonine-tagged lipopeptides synergize in host colonization by pathogenic Burkholderia species. ACS Chem Biol 2018;13:1370–9. [75] Niehs SP, Scherlach K, Hertweck C. Genomics-driven discovery of a linear lipopeptide promoting host colonization by endofungal bacteria. Org Biomol Chem 2018;16:8345–52. [76] Chen H, Zhou H, Sun T, Xu J, Tu Q, Yang J, et al. Identification of Holrhizins E-Q reveals the diversity of nonribosomal lipopeptides in Paraburkholderia rhizoxinica. J Nat Prod 2020;83:537–41. [77] Deng MC, Li J, Hong YH, Xu XM, Chen WX, Yuan JP, et al. Characterization of a novel biosurfactant produced by marine hydrocarbon-degrading bacterium Achromobacter sp. HZ01. J Appl Microbiol 2016;120:889–99. [78] Matsuyama T, Fujita M, Yano I. Wetting agent produced by Serratia marcescens. FEMS Microbiol Lett 1985;28:125–9. [79] Matsuyama T, Kaneda K, Nakagawa Y, Isa K, Hara-Hotta H, Yano I. A novel extracellular cyclic lipopeptide which promotes flagellum-dependent and -independent spreading growth of Serratia marcescens. J Bacteriol 1992;174(6):1769–76. [80] Matsuyama T, Murakami T, Fujita M, Fujita S, Yano I. Extracellular vesicle formation and biosurfactant production by Serratia marcescens. J Gen Microbiol 1986;132(4):865–75.

References

237

[81] Su C, Xiang Z, Liu Y, Zhao X, Sun Y, Li Z, et al. Analysis of the genomic sequences and metabolites of Serratia surfactantfaciens sp. nov. YD25T that simultaneously produces prodigiosin and serrawettin W2. BMC Genomics 2016;17(1):1–19. [82] Kadouri DE, Shanks RMQ. Identification of a methicillin-resistant Staphylococcus aureus inhibitory compound isolated from Serratia marcescens. Res Microbiol 2013;164(8):821–6. [83] Clements T, Ndlovu T, Khan W. Broad-spectrum antimicrobial activity of secondary metabolites produced by Serratia marcescens strains. Microbiol Res 2019;229, 126329. [84] Ganley JG, Carr G, Ioerger TR, Sacchettini JC, Clardy J, Derbyshire ER. Discovery of antimicrobial lipodepsipeptides produced by a Serratia sp. within mosquito microbiomes. ChemBioChem 2018;19:1590–4. [85] Qiao N, Shao Z. Isolation and characterization of a novel biosurfactant produced by hydrocarbon-degrading bacterium Alcanivorax dieselolei B-5. J Appl Microbiol 2010;108:1207–16. [86] Senthil Balan S, Ganesh Kumar C, Jayalakshmi S. Pontifactin, a new lipopeptide biosurfactant produced by a marine Pontibacter korlensis strain SBK-47: purification, characterization and its biological evaluation. Process Biochem 2016;51:2198–207. [87] Abdel-Aziz MM, Al-Omar MS, Mohammed HA, Emam TA. In vitro and ex vivo antibiofilm activity of a lipopeptide biosurfactant produced by the entomopathogenic Beauveria bassiana strain against Microsporum canis. Microorganisms 2020;8(232):1–16. [88] Qazi MA, Kanwal T, Jadoon M, Ahmed S, Fatima N. Isolation and characterization of a biosurfactantproducing Fusarium sp. BS-8 from oil contaminated soil. Biotechnol Prog 2014;30(5):1065–75. [89] Alencar S, do Amaral Marques N, Alves Lima e Silva T, Fontenele da Silva Andrade R, Fabiann Branco Junior J, Okada K, Maria Campos Takaki G. Lipopeptide biosurfactant produced by Mucor circinelloides UCP/WFCC 0001 applied in the removal of crude oil and engine oil from soil. Acta Sci Technol 2019;41:1–9. [90] Horowitz S, Gilbert JN, Griffin WM. Isolation and characterization of a surfactant produced by Bacillus licheniformis 86. J Ind Microbiol 1990;6:243–8. [91] Naruse N, Tenmyo O, Kobaru S, Kamei H, Miyaki T, Konishi M, et al. Pumilacidin, a complex of new antiviral antibiotics production, isolation, chemical properties, structure and biological activity. J Antibiot (Tokyo) 1990;43(3):267–80. [92] Peypoux F, Bonmatin JM, Labbe H, Das BC, Ptak M, Michel G. Isolation and characterization of a new variant of surfactin, the [Val7]surfactin. Eur J Biochem 1991;202:101–6. [93] Arima K, Kakinuma A, Tamura G. Surfactin, a crystalline peptidelipid surfactant produced by Bacillus subtilis: isolation, characterization and its inhibition of fibrin clot formation. Biochem Biophys Res Commun 1968;31(3):488–94. [94] Long X, He N, He Y, Jiang J, Wu T. Biosurfactant surfactin with pH-regulated emulsification activity for efficient oil separation when used as emulsifier. Bioresour Technol 2017;241:200–6. [95] Abdel-Mawgoud AM, Hassouna NA. Characterization of surfactin produced by Bacillus subtilis isolate BS5. Appl Biochem Biotechnol 2008;150:289–303. [96] Ishigami Y, Osman M, Nakahara H, Sano Y, Ishiguro R, Matsumoto M. Significance of beta-sheet formation for micellization and surface adsorption of surfactin. Colloids Surf B Biointerfaces 1995;4:341–8. [97] Grangemard I, Wallach J, Maget-Dana R, Peypoux F. Lichenysin: a more efficient cation chelator than surfactin. Appl Biochem Biotechnol 2001;90(3):199–210. [98] de Araujo LLGC, Sodre LGP, Brasil LR, Domingos DF, de Oliveira VM, da Cruz GF. Microbial enhanced oil recovery using a biosurfactant produced by Bacillus safensis isolated from mangrove microbiota—part I biosurfactant characterization and oil displacement test. J Petrol Sci Eng 2019;180:950–7. [99] Razafindralambo H, Thonart P, Paquot M. Dynamic and equilibrium surface tensions of surfactin aqueous solutions. J Surfactant Deterg 2004;7:41–6. [100] Peypoux F, Guinand M, Michel G, Delcambe L, Das BC, Lederer E. Structure of Iturine A, a Peptidolipid Antibiotic from Bacillus subtilis. Biochemistry 1978;17(19):3992–6.

238

Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

[101] Maget-Dana R, Peypoux F. Iturins, a special class of pore-forming lipopeptides: biological and physicochemical properties. Toxicology 1994;87(1–3):151–74. [102] Nishikiori T, Naganawa H, Muraoka Y, Aoyagi T, Umezawa H. Plipastatins: new inhibitors of phospholipase A2 produced by Bacillus Cereus BMG302-fF67 III. Structural elucidation of plipastatins. J Antibiot (Tokyo) 1986;39(6):755–61. [103] Steller S, Vollenbroich D, Leenders F, Stein T, Conrad B, Hofemeister J, et al. Structural and functional organization of the fengycin synthetase multienzyme system from Bacillus subtilis b213 and A1/3. Chem Biol 1999;6:31–41. [104] Lin TP, Chen CL, Chang LK, Tschen JSM, Liu ST. Functional and transcriptional analyses of a fengycin synthetase gene, fenC, from Bacillus subtilis. J Bacteriol 1999;181(16):5060–7. [105] Lin TP, Chen CL, Fu HC, Wu CY, Lin GH, Huang SH, et al. Functional analysis of fengycin synthetase FenD. Biochim Biophys Acta Gene Struct Expr 2005;1730(2):159–64. [106] Schneider J, Taraz K, Budzikiewicz H, Deleu M, Thonart P, Jacques P. The structure of two fengycins from Bacillus subtilis S499. Z Naturforsch C J Biosci 1999;54(11):859–66. [107] Lin GH, Chen CL, Tschen JSM, Tsay SS, Chang YS, Liu ST. Molecular cloning and characterization of fengycin synthetase gene fenB from Bacillus subtilis. J Bacteriol 1998;180(5):1338–41. [108] Abderrahmani A, Tapi A, Nateche F, Chollet M, Lecle`re V, Wathelet B, et al. Bioinformatics and molecular approaches to detect NRPS genes involved in the biosynthesis of kurstakin from Bacillus thuringiensis. Appl Microbiol Biotechnol 2011;92(3):571–81. [109] Bechet M, Caradec T, Hussein W, Abderrahmani A, Chollet M, Leclere V, et al. Structure, biosynthesis, and properties of kurstakins, nonribosomal lipopeptides from Bacillus spp. Appl Microbiol Biotechnol 2012;95 (3):593–600. [110] Gelis-Jeanvoine S, Canette A, Gohar M, Caradec T, Lemy C, Gominet M, et al. Genetic and functional analyses of krs, a locus encoding kurstakin, a lipopeptide produced by Bacillus thuringiensis. Res Microbiol 2017;168(4):356–68. [111] Kajimura Y, Kaneda M, Fusaricidin A. a new depsipeptide antibiotic produced by Bacillus polymyxa KT-8 taxonomy, fermentation, isolation, structure elucidation and biological activity. J Antibiot (Tokyo) 1996;49 (2):129–35. [112] Kajimura Y, Kaneda M, Fusaricidins B. C and D, new depsipeptide antibiotics produced by Bacillus polymyxa KT-8: isolation, structure elucidation and biological activity. J Antibiot (Tokyo) 1997;50 (3):220–8. [113] Ohno O, Ikeda Y, Sawa R, Igarashi M, Kinoshita N, Suzuki Y, et al. Isolation of heptadepsin, a novel bacterial cyclic depsipeptide that inhibits lipopolysaccharide activity. Chem Biol 2004;11:1059–70. [114] Guo Y, Huang E, Yuan C, Zhang L, Yousef AE. Isolation of a Paenibacillus sp. strain and structural elucidation of its broad-spectrum lipopeptide antibiotic. Appl Environ Microbiol 2012;78(9):3156–65. [115] Velkov T, Gallardo-Godoy A, Swarbrick JD, Blaskovich MAT, Elliott AG, Han M, et al. Structure, function and biosynthetic origin of octapeptin antibiotics active against extensively drug-resistant Gram-negative bacteria. Cell Chem Biol 2018;25(4):380–91. [116] Kim J, Le KD, Yu NH, Kim JI, Kim JC, Lee CW. Structure and antifungal activity of pelgipeptins from Paenibacillus elgii against phytopathogenic fungi. Pestic Biochem Physiol 2020;163:154–63. [117] Storm DR, Rosenthal KS, Swanson PE. Polymyxin and related peptide antibiotics. Annu Rev Biochem 1977;46:723–63. [118] Choi SK, Park SY, Kim R, Kim SB, Lee CH, Kim JF, et al. Identification of a polymyxin synthetase gene cluster of Paenibacillus polymyxa and heterologous expression of the gene in Bacillus subtilis. J Bacteriol 2009;191(10):3350–8. [119] Wallace SJ, Li J, Nation RL, Prankerd RJ, Velkov T, Boyd BJ. Self-assembly behaviour of colistin and its prodrug colistin methanesulfonate: implications for solution stability and solubilization. J Phys Chem B 2010;114(14):4836–40.

References

239

[120] Barsby T, Warabi K, Sørensen D, Zimmerman WT, Kelly MT, Andersen RJ. The bogorol family of antibiotics: template-based structure elucidation and a new approach to positioning enantiomeric pairs of amino acids. J Org Chem 2006;71:6031–7. [121] Desjardine K, Pereira A, Wright H, Matainaho T, Kelly M, Andersen RJ. Tauramamide, a lipopeptide antibiotic produced in culture by Brevibacillus laterosporus isolated from a marine habitat: structure elucidation and synthesis. J Nat Prod 2007;70:1850–3. [122] Barsby T, Kelly MT, Andersen RJ. Tupuseleiamides and basiliskamides, new acyldipeptides and antifungal polyketides produced in culture by a Bacillus laterosporus isolate obtained from a tropical marine habitat. J Nat Prod 2002;65:1447–51. [123] Aretz W, Meiwes J, Seibert G, Vobis G, Wink J. Friulimicins: Novel lipopeptide antibiotics with peptidoglycan synthesis inhibiting activity from Actinoplanes friuliensis sp. nov. I. Taxonomic studies of the producing microorganism and fermentation. J Antibiot (Tokyo) 2000;53(8):807–15. [124] Boeck LD, Papiska HR, Wetzel RW, Mynderse JS, Fukuda DS, Mertz FP, et al. A54145, a new lipopeptide antibiotic complex: discovery, taxonomy, fermentation and HPLC. J Antibiot (Tokyo) 1989;43(6):587–93. [125] Schimana J, Gebhardt K, H€oltzel A, Schmid DG, S€ ussmuth R, M€ uller J, et al. Arylomycins A and B, new biaryl-bridged lipopeptide antibiotics produced by Streptomyces sp. T€ u 6075. I. Taxonomy, fermentation, isolation and biological activities. J Antibiot (Tokyo) 2002;55(6):565–70. [126] Lakey J, Lea E, Rudd B, Wright H, Hopwood D. A new channel-forming antibiotic from Streptomyces coelicolor A3(2) which requires calcium for its activity. J Gen Microbiol 1983;129(12):3565–73. [127] Higashide E, Hatano K, Shibata M, Nakazawa K. Enduracidin, a new antibiotic. I. Streptomyces fungicidicus No. B5477, an enduracidin producing organism. J Antibiot (Tokyo) 1968;21(2):78–86. [128] Borders DB, Leese RA, Jarolmen H, Francis ND, Fantini AA, Falla T, et al. Laspartomycin, an acidic lipopeptide antibiotic with a unique peptide core. J Nat Prod 2007;70(3):443–6. [129] Bekiesch P, Zehl M, Domingo-Contreras E, Martı´n J, Perez-Victoria I, Reyes F, et al. Viennamycins: lipopeptides produced by a Streptomyces sp. J Nat Prod 2020;83(8):2381–9. [130] Wu C, Shang Z, Lemetre C, Ternei MA, Brady SF. Cadasides, calcium-dependent acidic lipopeptides from the soil metagenome that are active against multidrug-resistant bacteria. J Am Chem Soc 2019;141 (9):3910–9. [131] Hover BM, Kim SH, Katz M, Charlop-Powers Z, Owen JG, Ternei MA, et al. Culture-independent discovery of the malacidins as calcium-dependent antibiotics with activity against multidrug-resistant Grampositive pathogens. Nat Microbiol 2018;3(4):415–22. [132] Son S, Ko SK, Kim SM, Kim E, Kim GS, Lee B, et al. Antibacterial cyclic lipopeptide enamidonins with an enamide-linked acyl chain from a Streptomyces Species. J Nat Prod 2018;81:2462–9. ¯ mura S, et al. Sarpeptins A and B, [133] Koomsiri W, Inahashi Y, Leetanasaksakul K, Shiomi K, Takahashi YK, O lipopeptides produced by Streptomyces sp. KO-7888 overexpressing a specific SARP regulator. J Nat Prod 2019;82(8):2144–51. [134] Kitani S, Yoshida M, Boonlucksanawong O, Panbangred W, Anuegoonpipat A, Kurosu T, et al. Cystargamide B, a cyclic lipodepsipeptide with protease inhibitory activity from Streptomyces sp. J Antibiot (Tokyo) 2018;71:662–6. [135] Robbel L, Marahiel MA. Daptomycin, a bacterial lipopeptide synthesized by a nonribosomal machinery. J Biol Chem 2010;285(36):27501–8. [136] Nguyen KT, Kau D, Gu JQ, Brian P, Wrigley SK, Baltz RH, et al. A glutamic acid 3-methyltransferase encoded by an accessory gene locus important for daptomycin biosynthesis in Streptomyces roseosporus. Mol Microbiol 2006;61(5):1294–307. [137] Geudens N, Martins JC. Cyclic lipodepsipeptides from Pseudomonas spp.—biological Swiss-Army knives. Front Microbiol 2018;9(AUG):1–18. [138] Hiramoto M, Okada K, Nagai S, Kawamoto H. The structure of viscosin, a peptide antibiotic. I. Syntheses of D- and L-3-hydroxyacyl-L-leucine hydrazides related to viscosin. Chem Pharm Bull 1985;19(7):1308–14.

240

Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

[139] Kearns DB. A field guide to bacterial swarming motility. Nat Rev Microbiol 2010;8(9):634–44. [140] Renard P, Canet I, Sancelme M, Matulova M, Uhliarikova I, Eyheraguibel B, et al. Cloud microorganisms, an interesting source of biosurfactants. IntechOpen; 2019. p. 1–17. [141] Roongsawang N, Hase K, Haruki M, Imanaka T, Morikawa M, Kanaya S. Cloning and characterization of the gene cluster encoding arthrofactin synthetase from Pseudomonas sp. MIS38. Chem Biol 2003;10:869–80. [142] Guenzi E, Galli G, Grgurina I, Gross DC, Grandi G. Characterization of the syringomycin synthetase gene cluster: a link between prokaryotic and eukaryotic peptide synthetases. J Biol Chem 1998;273(49):32857–63. [143] Scholz-Schroeder BK, Soule JD, Gross DC. The sypA, sypB, and sypC synthetase genes encode twenty-two modules involved in the nonribosomal peptide synthesis of syringopeptin by Pseudomonas syringae pv. syringae B301D. Mol Plant Microbe Interact 2003;16(4):271–80. [144] Koch B, Nielsen TH, Sørensen D, Andersen JB, Christophersen C, Molin S, et al. Lipopeptide production in Pseudomonas sp. strain DSS73 is regulated by components of sugar beet seed exudate via the Gac twocomponent regulatory system. Appl Environ Microbiol 2002;68(9):4509–16. [145] Groboillot A, Portet-Koltalo F, Le Derf F, Feuilloley MJG, Orange N, Duclairoir PC. Novel application of cyclolipopeptide amphisin: feasibility study as additive to remediate polycyclic aromatic hydrocarbon (PAH) contaminated sediments. Int J Mol Sci 2011;12:1787–806. [146] Bode HB, Reimer D, Fuchs SW, Kirchner F, Dauth C, Kegler C, et al. Determination of the absolute configuration of peptide natural products by using stable isotope labeling and mass spectrometry. Chem A Eur J 2012;18(8):2342–8. [147] Scherlach K, Lackner G, Graupner K, Pidot S, Bretschneider T, Hertweck C. Biosynthesis and mass spectrometric imaging of tolaasin, the virulence factor of brown blotch mushroom disease. ChemBioChem 2013;14(18):2439–43. [148] Vaillancourt FH, Yin J, Walsh CT. SyrB2 in syringomycin E biosynthesis is a nonheme FeII α-ketoglutarate and O2-dependent halogenase. Proc Natl Acad Sci U S A 2005;102(29):10111–6. [149] Singh GM, Vaillancourt FH, Yin J, Walsh CT. Characterization of SyrC, an aminoacyltransferase shuttling threonyl and chlorothreonyl residues in the syringomycin biosynthetic assembly line. Chem Biol 2007;14:31–40. [150] Singh GM, Fortin PD, Koglin A, Walsh CT. β-hydroxylation of the aspartyl residue in the phytotoxin syringomycin E: characterization of two candidate hydroxylases AspH and SyrP in Pseudomonas syringae. Biochemistry 2008;47(43):11310–20. [151] Sorensen KN, Kim KH, Takemoto JY. In vitro antifungal and fungicidal activities and erythrocyte toxicities of cyclic lipodepsinonapeptides produced by Pseudomonas syringae pv. syringae. Antimicrob Agents Chemother 1996;40(12):2710–3. [152] Dalla Serra M, Fagiuoli G, Nordera P, Bernhart I, Della Volpe C, Di Giorgio D, et al. The interaction of lipodepsipeptide toxins from Pseudomonas syringae pv. syringae with biological and model membranes: a comparison of syringotoxin, syringomycin, and two syringopeptins. Mol Plant Microbe Interact 1999;12 (5):391–400. [153] Zouari O, Lecouturier D, Rochex A, Chataigne G, Dhulster P, Jacques P, et al. Bio-emulsifying and biodegradation activities of syringafactin producing Pseudomonas spp. strains isolated from oil contaminated soils. Biodegradation 2019;30:259–72. [154] Li H, Tanikawa T, Sato Y, Nakagawa Y, Matsuyama T. Serratia marcescens gene required for surfactant serrawettin W1 production encodes putative aminolipid synthetase belonging to nonribosomal peptide synthetase family. Microbiol Immunol 2005;49(4):303–10. [155] Motley JL, Stamps BW, Mitchell CA, Thompson AT, Cross J, You J, et al. Opportunistic sampling of roadkill as an entry point to accessing natural products assembled by bacteria associated with non-anthropoidal mammalian microbiomes. J Nat Prod 2017;80(3):598–608. [156] Matsuyama T, Bhasin A, Harshey RM. Mutational analysis of flagellum-independent surface spreading of Serratia marcescens 274 on a low-agar medium. J Bacteriol 1995;177(4):987–91.

References

241

[157] Jokela J, Oftedal L, Herfindal L, Permi P, Wahlsten M, Ove Døskeland S, et al. Anabaenolysins, novel cytolytic lipopeptides from benthic Anabaena cyanobacteria. PLoS One 2012;7(7). [158] Orjala J, Nagle DG, Hsu VL, Genvick WH. Antillatoxin: an exceptionally ihthyotoxic cyclic lipopeptide from the tropical cyanobacterium Lyngbya majuscula. J Am Chem Soc 1995;117:8281–2. [159] Luesch H, Yoshida WY, Moore RE, Paul VJ. Apramides A-G, novel lipopeptides from the marine cyanobacterium Lyngbya majuscula. J Nat Prod 2000;63:1106–12. [160] Orjala J, Gerwick WH. Barbamide, a chlorinated metabolite with molluscicidal activity from the Caribbean cyanobacterium Lyngbya majuscula. J Nat Prod 1996;59:427–30. [161] Hooper GJ, Orjala J, Schatzman RC, Gerwick WH. Carmabins A and B, new lipopeptides from the Caribbean cyanobacterium Lyngbya majuscula. J Nat Prod 1998;61:529–33. [162] Gerwick WH, Proteau PJ, Nagle DG, Hamel E, Blokhin A, Slate DL. Structure of Curacin A, a novel antimitotic, antiproliferative and brine shrimp toxic natural product from the marine cyanobacterium Lyngbya majuscula. J Org Chem 1994;59:1243–5. [163] Marquez BL, Watts KS, Yokochi A, Roberts MA, Verdier-Pinard P, Jimenez JI, et al. Structure and absolute stereochemistry of hectochlorin, a potent stimulator of actin assembly. J Nat Prod 2002;65:866–71. [164] Bonnard I, Rolland M, Francisco C, Banaigs B. Total structure and biological properties of laxaphycins A and B, cyclic lipopeptides from the marine cyanobacterium Lyngbya majuscula. Lett Pept Sci 1997;4:289–92. [165] Natsume N, Ozaki K, Nakajima D, Yokoshima S, Teruya T. Structure-activity relationship study of majusculamides A and B and their analogues on osteogenic activity. J Nat Prod 2020;83(8):2477–82. [166] Cardellina II JH, Marner FJ, Moore RE. Malyngamide A, a novel chlorinated metabolite of the marine cyanophyte Lyngbya Majuscula. J Am Chem Soc 1979;101(1):240–2. [167] Koehn FE, Longley RE, Reed JK. Microcolins A and B, new immunosuppressive peptides from the bluegreen alga Lyngbya majuscula. J Nat Prod 1992;55(5):613–9. [168] Nogle LM, Williamson RT, Gerwick WH. Somamides A and B, two new depsipeptide analogues of dolastatin 13 from a Fijian cyanobacterial assemblage of Lyngbya majuscula and Schizothrix species. J Nat Prod 2001;64:716–9. [169] Harrigan GG, Luesch H, Yoshida WY, Moore RE, Nagle DG, Paul VJ, et al. Symplostatin 1: a dolastatin 10 analogue from the marine cyanobacterium Symploca hydnoides. J Nat Prod 1998;61(9):1075–7. [170] Myokei R, Sakurai A, Chang CF, Kodaira Y, Takahashi N, Tamura S. Aspochracin, a new insecticidal metabolite of Aspergillus ochraceus. Agric Biol Chem 1969;33(10):1491–500. [171] Yue Q, Chen L, Zhang X, Li K, Sun J, Liu X, et al. Evolution of chemical diversity in echinocandin lipopeptide antifungal metabolites. Eukaryot Cell 2015;14(7):698–718. [172] Kuzma M, Jegorov A, Kacer P, Havlicek V. Sequencing of new beauverolides by high-performance liquid chromatography and mass spectrometry. J Mass Spectrom 2001;36:1108–15. [173] Shiono Y, Tsuchinari M, Shimanuki K, Miyajima T, Murayama T, Koseki T, et al. Fusaristatins A and B, two new cyclic lipopeptides from an endophytic Fusarium sp. J Antibiot (Tokyo) 2007;60(5):309–16. [174] Darkin-Rattray SJ, Gurnett AM, Myers RW, Dulski PM, Crumley TM, Allocco JJ, et al. Apicidin: a novel antiprotozoal agent that inhibits parasite histone deacetylase. Proc Natl Acad Sci U S A 1996;93 (23):13143–7. [175] Westphal KR, Nielsen KAH, Wollenberg RD, Møllehøj MB, Bachleitner S, Studt L, et al. Fusaoctaxin a, an example of a two-step mechanism for non-ribosomal peptide assembly and maturation in fungi. Toxins 2019;11(5). [176] Bahadoor A, Brauer EK, Bosnich W, Schneiderman D, Johnston A, Aubin Y, et al. Gramillin A and B: cyclic lipopeptides identified as the nonribosomal biosynthetic products of Fusarium graminearum. J Am Chem Soc 2018;140(48):16783–91. [177] Nihei K, Itoh H, Hashimoto K, Miyairi K, Okuno T. Antifungal cyclodepsipeptides, W493 A and B, from Fusarium sp.: isolation and structural determination. Biosci Biotechnol Biochem 1998;62(5):858–63.

242

Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

[178] Fotso S, Graupner P, Xiong Q, Gilbert JR, Hahn D, Avila-Adame C, et al. Alveolarides: antifungal peptides from Microascus alveolaris active against phytopathogenic fungi. J Nat Prod 2018;81:10–5. [179] Oh DC, Kauffman CA, Jensen PR, Fenical W. Induced production of emericellamides A and B from the marine-derived fungus Emericella sp. in competing co-culture. J Nat Prod 2007;70(4):515–20. [180] Seto Y, Takahashi K, Matsuura H, Kogami Y, Yada H, Yoshihara T, et al. Novel cyclic peptide, epichlicin, from the endophytic fungus, Epichloe typhina. Biosci Biotechnol Biochem 2007;71(6):1470–5. [181] Shigemori H, Wakuri S, Yazawa K, Nakamura T, Sasaki T, Kobayashi J. Fellutamides A and B, cytotoxic peptides from a marine fish-possessing fungus Penicillium fellutanum. Tetrahedron 1991;47(40):8529–34. [182] Yu Z, Lang G, Kajahn I, Schmaljohann R, Imhoff JF. Scopularides A and B, cyclodepsipeptides from a marine sponge-derived fungus, Scopulariopsis brevicaulis. J Nat Prod 2008;71:1052–4. [183] Pruksakorn P, Arai M, Kotoku N, Vilcheze C, Baughn AD, Moodley P, et al. Trichoderins, novel aminolipopeptides from a marine sponge-derived Trichoderma sp., are active against dormant mycobacteria. Bioorg Med Chem Lett 2010;20(12):3658–63. [184] Ishidoh K, Kinoshita H, Igarashi Y, Ihara F, Nihira T. Cyclic lipodepsipeptides verlamelin A and B, isolated from entomopathogenic fungus Lecanicillium sp. J Antibiot (Tokyo) 2014;67(6):459–63. [185] Zhao P, Xue Y, Li X, Li J, Zhao Z, Quan C, et al. Fungi-derived lipopeptide antibiotics developed since 2000. Peptides 2019;113:52–65. [186] Bills G, Li Y, Chen L, Yue Q, Niu XM, An Z. New insights into the echinocandins and other fungal nonribosomal peptides and peptaibiotics. Nat Prod Rep 2014;31(10):1348–75. [187] Zampella A, D’Auria MV, Gomez Paloma L, Casapullo A, Minale L, Debitus C, et al. Callipeltin A, an antiHIV cyclic depsipeptide from the new caledonian lithistida sponge Callipelta sp. J Am Chem Soc 1996;118:6202–9. [188] Nakao Y, Kawatsu S, Okamoto C, Okamoto M, Matsumoto Y, Matsunaga S, et al. Ciliatamides A-C, bioactive lipopeptides from the deep-Sea Sponge Aaptos ciliata. J Nat Prod 2008;71:469–72. [189] Oku N, Gustafson KR, Cartner LK, Wilson JA, Shigematsu N, Hess S, et al. Neamphamide A, a New HIVinhibitory depsipeptide from the Papua New Guinea marine sponge Neamphius huxleyi. J Nat Prod 2004;67:1407–11. [190] Ford PW, Gustafson KR, McKee TC, Shigematsu N, Maurizi LK, Pannell LK, et al. Papuamides A-D, HIVinhibitory and cytotoxic depsipeptides from the sponges Theonella mirabilis and Theonella swinhoei collected in Papua New Guinea. J Am Chem Soc 1999;13:5899–909. [191] Reese MT, Gulavita NK, Nakao Y, Hamann MT, Yoshida WY, Coval SJ, et al. Kulolide: a cytotoxic depsipeptide from a cephalaspidean mollusk, Philinopsis speciosa. J Am Chem Soc 1996;118:11081–4. [192] Hamann MT, Otto CS, Scheuer PJ, Dunbar DC. Kahalalides: bioactive peptides from a marine mollusk Elysia rufescens and its algal diet Bryopsis sp. J Org Chem 1996;61:6594–600. [[193] Lourenc¸o LA, Alberton Magina MD, Ballod Tavares LB, Guelli Ulson de Souza SMA, Garcı´a Roman M, Altmajer Vaz D. Biosurfactant production by Trametes versicolor grown on two-phase olive mill waste in solid-state fermentation. Environ Technol 2018;39(23):3066–76. [194] Bao M, Pi Y, Wang L, Sun P, Li Y, Cao L. Lipopeptide biosurfactant production bacteria Acinetobacter sp. D3-2 and its biodegradation of crude oil. Environ Sci Process Impacts 2014;16:897–903. [195] Piegza M, Pietrzykowska J, Trojan-Piegza J, Laba W. Biosurfactants from Trichoderma filamentous fungi—a preliminary study. Biomolecules 2021;11(519):1–13. [196] Burgos-Dı´az C, Pons R, Espuny MJ, Aranda FJ, Teruel JA, Manresa A, et al. Isolation and partial characterization of a biosurfactant mixture produced by Sphingobacterium sp. isolated from soil. J Colloid Interface Sci 2011;361:195–204. [197] Hajfarajollah H, Mokhtarani B, Akbari NK. Newly antibacterial and antiadhesive lipopeptide biosurfactant secreted by a probiotic strain, Propionibacterium freudenreichii. Appl Biochem Biotechnol 2014;174:2725–40.

References

243

[198] Gandhimathi R, Seghal Kiran G, Hema TA, Selvin J, Rajeetha Raviji T, Shanmughapriya S. Production and characterization of lipopeptide biosurfactant by a sponge-associated marine actinomycetes Nocardiopsis alba MSA10. Bioprocess Biosyst Eng 2009;32:825–35. [199] Femina Carolin C, Senthil Kumar P, Janet Joshiba G, Madhesh P, Ramamurthy R. Sustainable strategy for the enhancement of hazardous aromatic amine degradation using lipopeptide biosurfactant isolated from Brevibacterium casei. J Hazard Mater 2021;408, 124943. [200] Chaves Martins P, Garcia Bastos C, Afonso Granjeiro P, Guimara˜es MV. New lipopeptide produced by Corynebacterium aquaticum from a low-cost substrate. Bioprocess Biosyst Eng 2018;41(8):1177–83. [201] Huang Y, Zhou H, Zheng G, Li Y, Xie Q, You S, et al. Isolation and characterization of biosurfactantproducing Serratia marcescens ZCF25 from oil sludge and application to bioremediation. Environ Sci Pollut Res 2020;27:27762–72. [202] Vilela WFD, Fonseca SG, Fantinatti-Garboggini F, Oliveira VM, Nitschke M. Production and properties of a surface-active lipopeptide produced by a new marine Brevibacterium luteolum strain. Appl Biochem Biotechnol 2014;174:2245–56. [203] Chebbi A, Hentati D, Cheffi M, Bouabdallah R, Choura C, Sayadi S, et al. Promising abilities of mercaptodegrading Staphylococcus capitis strain SH6 in both crude oil and waste motor oil as sole carbon and energy sources: its biosurfactant production and preliminary characterization. J Chem Technol Biotechnol 2018;93:1401–12. [204] Najafi AR, Rahimpour MR, Jahanmiri AH, Roostaazad R, Arabian D, Soleimani M, et al. Interactive optimization of biosurfactant production by Paenibacillus alvei ARN63 isolated from an Iranian oil well. Colloids Surf B Biointerfaces 2011;82:33–9. [205] Marchant R, Banat IM. Biosurfactants: a sustainable replacement for chemical surfactants? Biotechnol Lett 2012;34(9):1597–605. [206] Santos D, Rufino R, Luna J, Santos V, Sarubbo L. Biosurfactants: multifunctional biomolecules of the 21st century. Int J Mol Sci 2016;17(3):401. [207] De Almeida DG, Soares Da Silva RCF, Luna JM, Rufino RD, Santos VA, Banat IM, et al. Biosurfactants: promising molecules for petroleum biotechnology advances. Front Microbiol 2016;7:1718. [208] Alanjary M, Cano-Prieto C, Gross H, Medema MH. Computer-aided re-engineering of nonribosomal peptide and polyketide biosynthetic assembly lines. Nat Prod Rep 2019;36(9):1249–61. [209] Dhali D, Coutte F, Arias AA, Auger S, Bidnenko V, Chataigne G, et al. Genetic engineering of the branched fatty acid metabolic pathway of Bacillus subtilis for the overproduction of surfactin C14 isoform. Biotechnol J 2017;12(7):1600574. [210] O’Connor NK, Rai DK, Clark BR, Murphy CD. Production of the novel lipopeptide antibiotic trifluorosurfactin via precursor-directed biosynthesis. J Fluor Chem 2012;143:210–5. [211] Moran S, Rai DK, Clark BR, Murphy CD. Precursor-directed biosynthesis of fluorinated iturin A in Bacillus spp. Org Biomol Chem 2009;7(4):644. [212] Micklefield J. Biosynthesis and biosynthetic engineering of calcium-dependent lipopeptide antibiotics. Pure Appl Chem 2009;81(6):1065–74. [213] Hug JJ, Krug D, M€uller R. Bacteria as genetically programmable producers of bioactive natural products. Nat Rev Chem 2020;4(4):172–93. [214] Weissman KJ. Mutasynthesis—uniting chemistry and genetics for drug discovery. Trends Biotechnol 2007;25(4):139–42. [215] Hojati Z, Milne C, Harvey B, Gordon L, Borg M, Flett F, et al. Structure, biosynthetic origin, and engineered biosynthesis of calcium-dependent antibiotics from Streptomyces coelicolor. Chem Biol 2002;9 (11):1175–87. [216] Stanisˇic A, Kries H. Adenylation domains in nonribosomal peptide engineering. ChemBioChem 2019;20 (11):1347–56.

244

Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

[217] Eppelmann K, Stachelhaus T, Marahiel MA. Exploitation of the selectivity-conferring code of nonribosomal peptide synthetases for the rational design of novel peptide antibiotics. Biochemistry 2002;41 (30):9718–26. [218] Han JW, Kim EY, Lee JM, Kim YS, Bang E, Kim BS. Site-directed modification of the adenylation domain of the fusaricidin nonribosomal peptide synthetase for enhanced production of fusaricidin analogs. Biotechnol Lett 2012;34(7):1327–34. [219] Theatre A, Cano-Prieto C, Bartolini M, Laurin Y, Deleu M, Niehren J, et al. The surfactin-like lipopeptides From Bacillus spp.: natural biodiversity and synthetic biology for a broader application range. Front Bioeng Biotechnol 2021;9:623701. [220] Baltz RH, Brian P, Miao V, Wrigley SK. Combinatorial biosynthesis of lipopeptide antibiotics in Streptomyces roseosporus. J Ind Microbiol Biotechnol 2006;33(2):66–74. [221] Bozh€uy€uk KAJ, Linck A, Tietze A, Kranz J, Wesche F, Nowak S, et al. Modification and de novo design of non-ribosomal peptide synthetases using specific assembly points within condensation domains. Nat Chem 2019;11(7):653–61. [222] Mootz HD, Kessler N, Linne U, Eppelmann K, Schwarzer D, Marahiel MA. Decreasing the ring size of a cyclic nonribosomal peptide antibiotic by in-frame module deletion in the biosynthetic genes. J Am Chem Soc 2002;124(37):10980–1. [223] Jiang J, Gao L, Bie X, Lu Z, Liu H, Zhang C, et al. Identification of novel surfactin derivatives from NRPS modification of Bacillus subtilis and its antifungal activity against Fusarium moniliforme. BMC Microbiol 2016;16(1):31. [224] Gao L, Liu H, Ma Z, Han J, Lu Z, Dai C, et al. Translocation of the thioesterase domain for the redesign of plipastatin synthetase. Sci Rep 2016;6(1):38467. [225] Giessen TW, Marahiel MA. Ribosome-independent biosynthesis of biologically active peptides: application of synthetic biology to generate structural diversity. FEBS Lett 2012;586(15):2065–75. [226] D’Costa VM, Mukhtar TA, Patel T, Koteva K, Waglechner N, Hughes DW, et al. Inactivation of the lipopeptide antibiotic daptomycin by hydrolytic mechanisms. Antimicrob Agents Chemother 2012;56(2):757– 64. [227] Koglin A, Doetsch V, Bernhard F. Molecular engineering aspects for the production of new and modified biosurfactants. In: Sen R, editor. Biosurfactants. Biosurfactants. Advances in experimental medicine and biology, vol 672. New York, NY: Springer; 2010. p. 158–69. [228] Baltz RH. Combinatorial biosynthesis of cyclic lipopeptide antibiotics: a model for synthetic biology to accelerate the evolution of secondary metabolite biosynthetic pathways. ACS Synth Biol 2014;3 (10):748–58. [229] Miao V, Coe¨ffet-LeGal MF, Brian P, Brost R, Penn J, Whiting A, et al. Daptomycin biosynthesis in Streptomyces roseosporus: cloning and analysis of the gene cluster and revision of peptide stereochemistry. Microbiology 2005;151(5):1507–23. [230] Alexander DC, Rock J, Gu JQ, Mascio C, Chu M, Brian P, et al. Production of novel lipopeptide antibiotics related to A54145 by Streptomyces fradiae mutants blocked in biosynthesis of modified amino acids and assignment of lptJ, lptK and lptL gene functions. J Antibiot (Tokyo) 2011;64(1):79–87. [231] Ohlendorf B, Kehraus S, K€onig GM. Myxochromide B3, a new member of the myxochromide family of secondary metabolites. J Nat Prod 2008;71(10):1708–13. [232] Yan F, Burgard C, Popoff A, Zaburannyi N, Zipf G, Maier J, et al. Synthetic biology approaches and combinatorial biosynthesis towards heterologous lipopeptide production. Chem Sci 2018;9(38):7510–9. [233] Miao V, Coe¨ffet-Le Gal MF, Nguyen K, Brian P, Penn J, Whiting A, et al. Genetic engineering in Streptomyces roseosporus to produce hybrid lipopeptide antibiotics. Chem Biol 2006;13(3):269–76. [234] Baltz RH. Biosynthesis and genetic engineering of lipopeptides in Streptomyces roseosporus. In: Methods in enzymology. Academic Press; 2009. p. 511–31. Complex Enzymes in Microbial Natural Product Biosynthesis, Part A: Overview Articles and Peptides; vol. 458.

References

245

[235] Baltz RH, Nguyen KT, Alexander DC. Genetic engineering of acidic lipopeptide antibiotics. In: Bull AT, Junker B, Katz L, Lynd LR, Masurekar P, Reeves CD, et al., editors. Manual of industrial microbiology and biotechnology. Washington, DC, USA: ASM Press; 2010. p. 391–410. [236] Powell A, Borg M, Amir-Heidari B, Neary JM, Thirlway J, Wilkinson B, et al. Engineered biosynthesis of nonribosomal lipopeptides with modified fatty acid side chains. J Am Chem Soc 2007;129 (49):15182–91. [237] Ye C, Ng IS, Jing K, Lu Y. Direct proteomic mapping of Streptomyces roseosporus NRRL 11379 with precursor and insights into daptomycin biosynthesis. J Biosci Bioeng 2014;117(5):591–7. [238] Lecle`re V, Bechet M, Adam A, Guez JS, Wathelet B, Ongena M, et al. Mycosubtilin overproduction by Bacillus subtilis BBG100 enhances the organism’s antagonistic and biocontrol activities. Appl Environ Microbiol 2005;71(8):4577–84. [239] Wenzel SC, Gross F, Zhang Y, Fu J, Stewart AF, M€ uller R. Heterologous expression of a myxobacterial natural products assembly line in pseudomonads via red/ET recombineering. Chem Biol 2005;12 (3):349–56. [240] Qiu Y, Xiao F, Wei X, Wen Z, Chen S. Improvement of lichenysin production in Bacillus licheniformis by replacement of native promoter of lichenysin biosynthesis operon and medium optimization. Appl Microbiol Biotechnol 2014;98(21):8895–903. [241] Coutte F, Lecle`re V, Bechet M, Guez JS, Lecouturier D, Chollet-Imbert M, et al. Effect of pps disruption and constitutive expression of srfA on surfactin productivity, spreading and antagonistic properties of Bacillus subtilis 168 derivatives. J Appl Microbiol 2010;109(2):480–91. [242] Roongsawang N, Washio K, Morikawa M. Diversity of nonribosomal peptide synthetases involved in the biosynthesis of lipopeptide biosurfactants. Int J Mol Sci 2010;12(1):141–72. [243] Auchtung JM, Lee CA, Grossman AD. Modulation of the ComA-dependent quorum response in Bacillus subtilis by multiple Rap proteins and Phr peptides. J Bacteriol 2006;188(14):5273–85. [244] Nakano S, Nakano MM, Zhang Y, Leelakriangsak M, Zuber P. A regulatory protein that interferes with activator-stimulated transcription in bacteria. Proc Natl Acad Sci 2003;100(7):4233–8. [245] Hayashi K, Ohsawa T, Kobayashi K, Ogasawara N, Ogura M. The H2O2 stress-responsive regulator PerR positively regulates srfA expression in Bacillus subtilis. J Bacteriol 2005;187(19):6659–67. [246] M€ader U, Antelmann H, Buder T, Dahl M, Hecker M, Homuth G. Bacillus subtilis functional genomics: genome-wide analysis of the DegS-DegU regulon by transcriptomics and proteomics. Mol Genet Genomics 2002;268(4):455–67. [247] Tsuge K, Ano T, Hirai M, Nakamura Y, Shoda M. The genes degQ, pps, and lpa-8(sfp) are responsible for conversion of Bacillus subtilis 168 to plipastatin production. Antimicrob Agents Chemother 1999;43 (9):2183–92. [248] Dang Y, Zhao F, Liu X, Fan X, Huang R, Gao W, et al. Enhanced production of antifungal lipopeptide iturin A by Bacillus amyloliquefaciens LL3 through metabolic engineering and culture conditions optimization. Microb Cell Fact 2019;18(68). [249] Mao XM, Luo S, Li YQ. Negative regulation of daptomycin production by DepR2, an ArsR-family transcriptional factor. J Ind Microbiol Biotechnol 2017;44(12):1653–8. [250] Mao XM, Luo S, Zhou RC, Wang F, Yu P, Sun N, et al. Transcriptional regulation of the daptomycin gene cluster in Streptomyces roseosporus by an autoregulator, AtrA. J Biol Chem 2015;290(12):7992–8001. [251] Sun H, Bie X, Lu F, Lu Y, Wu Y, Lu Z. Enhancement of surfactin production of Bacillus subtilis fmbR by replacement of the native promoter with the Pspac promoter. Can J Microbiol 2009;55(8):1003–6. [252] Willenbacher J, Mohr T, Henkel M, Gebhard S, Mascher T, Syldatk C, et al. Substitution of the native srfA promoter by constitutive Pveg in two B. subtilis strains and evaluation of the effect on Surfactin production. J Biotechnol 2016;224:14–7. [253] Jiao S, Li X, Yu H, Yang H, Li X, Shen Z. In situ enhancement of surfactin biosynthesis in Bacillus subtilis using novel artificial inducible promoters. Biotechnol Bioeng 2017;114(4):832–42.

246

Chapter 10 Increasing the natural biodiversity of microbial lipopeptides

[254] Wu Q, Zhi Y, Xu Y. Systematically engineering the biosynthesis of a green biosurfactant surfactin by Bacillus subtilis 168. Metab Eng 2019;52:87–97. [255] Ji CH, Kim H, Je HW, Kwon H, Lee D, Kang HS. Top-down synthetic biology approach for titer improvement of clinically important antibiotic daptomycin in Streptomyces roseosporus. Metab Eng 2022;69:40–9. [256] Tan W, Yin Y, Wen J. Increasing fengycin production by strengthening the fatty acid synthesis pathway and optimizing fermentation conditions. Biochem Eng J 2022;177:108235. [257] Wang C, Cao Y, Wang Y, Sun L, Song H. Enhancing surfactin production by using systematic CRISPRi repression to screen amino acid biosynthesis genes in Bacillus subtilis. Microb Cell Fact 2019;18(1):90. [258] Coutte F, Niehren J, Dhali D, John M, Versari C, Jacques P. Modeling leucine’s metabolic pathway and knockout prediction improving the production of surfactin, a biosurfactant from Bacillus subtilis. Biotechnol J 2015;10(8):1216–34. [259] Zhang F, Huo K, Song X, Quan Y, Wang S, Zhang Z, et al. Engineering of a genome-reduced strain Bacillus amyloliquefaciens for enhancing surfactin production. Microb Cell Fact 2020;19(1):223. [260] Geissler M, K€uhle I, Morabbi Heravi K, Altenbuchner J, Henkel M, Hausmann R. Evaluation of surfactin synthesis in a genome reduced Bacillus subtilis strain. AMB Express 2019;9(1):84. [261] Yu G, Jia X, Wen J, Lu W, Wang G, Caiyin Q, et al. Strain Improvement of Streptomyces roseosporus for daptomycin production by rational screening of He–Ne laser and NTG induced mutants and kinetic modeling. Appl Biochem Biotechnol 2011;163(6):729–43. [262] Yu G, Hu Y, Hui M, Chen L, Wang L, Liu N, et al. Genome shuffling of Streptomyces roseosporus for improving daptomycin production. Appl Biochem Biotechnol 2014;172(5):2661–9. [263] Tsuge K, Ohata Y, Shoda M. Gene yerP, involved in surfactin self-resistance in Bacillus subtilis. Antimicrob Agents Chemother 2001;45(12):3566–73. [264] Li X, Yang H, Zhang D, Li X, Yu H, Shen Z. Overexpression of specific proton motive force-dependent transporters facilitate the export of surfactin in Bacillus subtilis. J Ind Microbiol Biotechnol 2015;42(1):93– 103. [265] Zhi Y, Wu Q, Xu Y. Genome and transcriptome analysis of surfactin biosynthesis in Bacillus amyloliquefaciens MT45. Sci Rep 2017;7(1):40976. [266] Lee SK, Kim HR, Jin YY, Yang SH, Suh JW. Improvement of daptomycin production via increased resistance to decanoic acid in Streptomyces roseosporus. J Biosci Bioeng 2016;122(4):427–33. [267] Nitschke M, Pastore GM. Biosurfactant production by Bacillus subtilis using cassava-processing effluent. Appl Biochem Biotechnol 2004;112(3):163–72. [268] Maass D, Ramı´rez IM, Roma´n MG, Alameda EJ, de Souza AAU, Valle JAB, et al. Two-phase olive mill waste (alpeorujo) as carbon source for biosurfactant production. J Chem Technol Biotechnol 2016;91 (7):1990–7. [269] Perlova O, Gerth K, Kuhlmann S, Zhang Y, M€ uller R. Novel expression hosts for complex secondary metabolite megasynthetases: production of myxochromide in the thermopilic isolate Corallococcus macrosporus GT-2. Microb Cell Fact 2009;8(1):1. [270] Penn J, Li X, Whiting A, Latif M, Gibson T, Silva CJ, et al. Heterologous production of daptomycin in Streptomyces lividans. J Ind Microbiol Biotechnol 2006;33(2):121–8. [271] Ke J, Yoshikuni Y. Multi-chassis engineering for heterologous production of microbial natural products. Curr Opin Biotechnol 2020;62:88–97. [272] Lee Y, Lee N, Hwang S, Kim K, Kim W, Kim J, et al. System-level understanding of gene expression and regulation for engineering secondary metabolite production in Streptomyces. J Ind Microbiol Biotechnol 2020;47(9–10):739–52. [273] Li J, Neubauer P. Escherichia coli as a cell factory for heterologous production of nonribosomal peptides and polyketides. N Biotechnol 2014;31(6):579–85. [274] Cai D, Zhu J, Zhu S, Lu Y, Zhang B, Lu K, et al. Metabolic engineering of main transcription factors in carbon, nitrogen, and phosphorus metabolisms for enhanced production of bacitracin in Bacillus licheniformis. ACS Synth Biol 2019;8(4):866–75.

References

247

[275] Tsuge K, Inoue S, Ano T, Itaya M, Shoda M. Horizontal transfer of iturin A operon, itu, to Bacillus subtilis 168 and conversion into an iturin A producer. Antimicrob Agents Chemother 2005;49(11):4641–8. [276] Thies S, Santiago-Sch€ubel B, Kovacic F, Rosenau F, Hausmann R, Jaeger KE. Heterologous production of the lipopeptide biosurfactant serrawettin W1 in Escherichia coli. J Biotechnol 2014;181:27–30. [277] Vassaux A, Meunier L, Vandenbol M, Baurain D, Fickers P, Jacques P, et al. Nonribosomal peptides in fungal cell factories: from genome mining to optimized heterologous production. Biotechnol Adv 2019;37(8), 107449. [278] Mutka SC, Carney JR, Liu Y, Kennedy J. Heterologous production of epothilone C and D in Escherichia coli. Biochemistry 2006;45(4):1321–30. [279] Zobel S, Kumpfm€uller J, S€ussmuth RD, Schweder T. Bacillus subtilis as heterologous host for the secretory production of the non-ribosomal cyclodepsipeptide enniatin. Appl Microbiol Biotechnol 2015;99(2):681– 91. [280] Jimoh AA, Lin J. Heterologous expression of Sfp-type phosphopantetheinyl transferase is Indispensable in the biosynthesis of lipopeptide biosurfactant. Mol Biotechnol 2019;61(11):836–51. [281] Bunet R, Riclea R, Laureti L, H^otel L, Paris C, Girardet JM, et al. A single Sfp-type phosphopantetheinyl transferase plays a major role in the biosynthesis of PKS and NRPS derived metabolites in Streptomyces ambofaciens ATCC23877. PLoS One 2014;9(1):e87607. [282] Gustafsson C, Govindarajan S, Minshull J. Codon bias and heterologous protein expression. Trends Biotechnol 2004;22(7):346–53. [283] Wehrs M, Prahl JP, Moon J, Li Y, Tanjore D, Keasling JD, et al. Production efficiency of the bacterial nonribosomal peptide indigoidine relies on the respiratory metabolic state in S. cerevisiae. Microb Cell Fact 2018;17(1):193. [284] Ongley SE, Bian X, Neilan BA, M€uller R. Recent advances in the heterologous expression of microbial natural product biosynthetic pathways. Nat Prod Rep 2013;30(8):1121. [285] Gibson DG, Young L, Chuang RY, Venter JC, Hutchison CA, Smith HO. Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 2009;6(5):343–5. [286] Bryksin AV, Matsumura I. Overlap extension PCR cloning: a simple and reliable way to create recombinant plasmids. Biotechniques 2010;48(6):463–5. [287] Gruenewald S, Mootz HD, Stehmeier P, Stachelhaus T. In vivo production of artificial nonribosomal peptide products in the heterologous host Escherichia coli. Appl Environ Microbiol 2004;70:3282–91. [288] Yan X, Yu HJ, Hong Q, Li SP. Cre/lox system and PCR-based genome engineering in Bacillus subtilis. Appl Environ Microbiol 2008;74(17):5556–62. [289] Bode HB, M€uller R. The impact of bacterial genomics on natural product research. Angew Chem Int Ed 2005;44(42):6828–46. [290] Ochi K, Hosaka T. New strategies for drug discovery: activation of silent or weakly expressed microbial gene clusters. Appl Microbiol Biotechnol 2013;97(1):87–98.

CHAPTER

Synthetic approaches to production of rhamnolipid and related glycolipids

11

Chett J. Boxleya, David E. Hoganb, Ryan M. Stolleya, and Raina M. Maierb a

GlycoSurf, Inc., Salt Lake City, UT, United States, bDepartment of Environmental Science, University of Arizona, Tucson, AZ, United States

1. Introduction Demand for green surfactants is increasing, as are advances in the production of sugar-based biosurfactants such as rhamnolipid (Fig. 1). The ability to produce biosurfactants at a large scale is causing significant growth in the popularity of these materials for a wide variety of applications. As a result, over the past several years the industry has witnessed accelerated growth in the commercialization of biosurfactants, especially sophorolipids and rhamnolipids. This growth has occurred alongside the explosive growth of “bio-based” surfactants where the term bio-based is defined as “…a commercial or industrial product, other than food or feed that is composed of, in whole or part, biological products, renewable domestic agricultural materials, or forestry materials” [1]. Such bio-based surfactants include the alkyl-poly glucosides (APGs), sugar esters, and phospholipid-based surfactants, among others. Likewise, biosurfactants may also be considered ‘bio-based surfactants’, but can further be defined as any microbially produced surfactant secreted by microorganisms into the culture medium when grown using hydrophobic carbon sources [2]. Examples of biosynthetic surfactants, both sugar-based and nonsugar-based include: rhamnolipids, sophorolipids, mannosylerythritol lipids, lipopeptides, and flavolipids—this chapter focuses on one type of sugar-based biosurfactant, rhamnolipid.

2. Rhamnolipids—Biosynthetic versus chemically synthesized 2.1 Congener distribution It is well known that for biosynthetic rhamnolipid, more than 100 unique structural congeners have been identified in the literature [3,4]. Biosynthetic rhamnolipids are produced by Pseudomonas aeruginosa and some Burkholderia species, and they are primarily categorized according to the number of sugar head groups present, e.g., mono- or di-rhamnolipid (Fig. 1). However, the structural diversity of rhamnolipids is far greater than simply the number of sugar headgroups present. In fact, research over the last 20 years shows the considerable diversity in the lipid tail group structure, primarily in the length of the β-hydroxy fatty acids that comprise this tail group [5–12]. In most cases, the literature Biosurfactants. https://doi.org/10.1016/B978-0-323-91697-4.00011-9 Copyright # 2023 Elsevier Inc. All rights reserved.

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Chapter 11 Synthetic approaches to production of rhamnolipid

FIG. 1 Examples of rhamnolipid structures formed biosynthetically. Monosaccharide rhamnolipid with single, double, and triple C10 chains are shown in the top row. Examples of disaccharide rhamnolipid with single, double, and triple C10 chains are shown in the bottom row.

shows two β-hydroxy fatty acid units; however, small percentages of single chain or even triple chain β-hydroxy fatty acids have also been reported [12]. Consequently, the use of biosynthetic production methods typically results in the formation of anywhere from 15 to 60 unique congeners in the final rhamnolipid mixture that is collected from any single bacterium following growth in culture. This complex mixture, of nearly inseparable chemical structures, is difficult to characterize and explore in a systematic way [5]. This is especially true when considering bacterium growth conditions cause frequent batch-to-batch variability in the resultant rhamnolipid congener mixture. This is true despite any attempt to control the growth conditions. Even modest shifts in the congener mixture can impart changes in surfactant characteristics [e.g., surface tension and critical micelle concentration (CMC)]. Further, the presence of even small minor amounts of congeners, which may seem trivial at first, can have remarkable impacts on the overall physicochemical properties of these materials and can thus impact their use in precision applications. For example, Zhang et al. [5], examined the impact of carbon source, particularly different length fatty acids, on monorhamnolipid produced by P. aeruginosa ATCC 9027. They found that the fatty acid substrate used influenced the mix of the four major monorhamnolipid congeners produced including the C10C10 monorhamnolipid (79 to 86% of the mixture) the C8C10 monorhamnolipid (3%–17% of the mixture), the C10C12 monorhamnolipid (5%–15% of the mixture) and the C10C12:1 monorhamnolipid (1%–2% of the mixture). The impact of this difference was rhamnolipid mixtures with a higher molar ratio of Rha-C10C8/Rha-C10C12 that were more effective at reducing surface tension. In contrast to biosynthetic methods, synthetic rhamnolipid methods produce a single congener that is structurally defined by the starting materials used. In particular, the choice of starting β-hydroxy acid chain length is specified at the outset of the synthesis process, as is the decision to produce single or

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double chain rhamnolipid. For instance, if single chain rhamnolipid C10 is desired, then only C10 β-hydroxy ester is used in the rhamnose glycosylation step (see description below). Whereas if a double chain C14C14 is desired, then enough C14 β-hydroxy ester is first produced, followed by a rhamnose glycosylation step (to form an intermediate single chain), and lastly, a coupling reaction step to form the final double chain product.

2.2 Stereochemistry One important difference in congener structure for synthetic and biosynthetic rhamnolipids is the stereochemistry of the molecule. Biosynthetic rhamnolipids are produced only as the R,R diastereomer [13]. For example, Fig. 2 shows a biosynthetic rhamnolipid C10C10 molecule, where the two stereocenters are both shown in the R conformation (the stereocenter location is labelled with an R). However, for synthetic rhamnolipids, all possible diastereomers are produced (in roughly equal percentages) as shown in Fig. 3 for a rhamnolipid C10C10 with four possible diastereomers. This difference might be

FIG. 2 Biosynthetic rhamnolipid C10C10 molecule with two stereocenters, both in the R conformation.

FIG. 3 The four possible congeners formed during the synthetic process.

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Chapter 11 Synthetic approaches to production of rhamnolipid

important for some applications that are focused on (or from a regulatory perspective require the use of) only “natural” products. Thus, it is noted that three of the four congeners present have not been observed in nature. Also, apart from whether or not these can be considered natural molecules, it is important to consider whether there are differences in chemical characteristics, toxicity, or biodegradability among diastereomers that could affect their use or environmental impact. These differences are discussed further in following sections. It should be noted that there are possible approaches to synthetically making only the R,R diastereomer. This includes the use of biologically derived β-hydroxy esters as starting materials and the separation of diastereomers following synthesis. Each of these approaches has advantages and disadvantages which will be discussed in the following section.

2.3 Purity As already mentioned, biosynthetic pathways for rhamnolipid production result in numerous congeners that are difficult to separate. In addition, the fermentation medium contains numerous secondary fermentation products that can be difficult to remove. Thus, purity levels that exceed 80%, while feasible, are labor-intensive, solvent-intensive, and generally cost-prohibitive [14,15]. In contrast, the synthetic process has built-in purification steps that produce high-purity single congener molecules (>95% pure).

2.4 Tailorability Biosynthetic rhamnolipids are congener mixtures that can be grossly tailored by: (1) choice of producing organism (e.g., there are natural mutants that produce only monorhamnolipid in contrast to the normally produced mono-di-mixture [16]); (2) choice of carbon and nitrogen sources for growth; and (3) genetic engineering of the producing organism [15]. However, a major advantage of synthetic approaches to making rhamnolipid is that the starting materials (type of carbohydrate, type and number of lipid tails) dictate the structure of the resulting glycolipid. This allows tailoring of the glycolipid with novel sugar head groups and/or novel tail groups. In fact, through designed choice of the starting materials, in the future it will be possible to tune the performance of the resulting glycolipid for a particular application. Choice of starting materials can also be made to influence the cost of the glycolipid produced. These ramifications are discussed more fully in the section below.

3. Chemical synthesis of rhamnolipids The need for reproducibility and the opportunity to be able to “tune” rhamnolipid molecular structure motivated the development of the synthetic production of these materials. Synthetically, there are three major approaches that depend on the type of rhamnolipid. Recall that there are four major structure types (Fig. 1): monolipid/monorhamnose, monolipid/polyrhamnose, polylipid/monorhamnose, and polylipid/polyrhamnose. In each of these molecules, the major synthetic concerns are stereospecific synthesis of the β-hydroxy lipid moieties, stereoselective formation of polyrhamnose moieties, and selective glycosylation of the lipid to the saccharide. In the simplest case, synthesis of the monolipid/monorhamnose structure, the synthesis typically begins with either use of biosynthetic or synthetically derived β-hydroxy esters. Using biologically derived esters has a distinct advantage as they typically produce the single chain R,R diastereomer;

3 Chemical synthesis of rhamnolipids

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however, selection of specific chain lengths is limited and the degree of saturation can vary. Alternatively, by synthetically producing the β-hydroxy esters one can overcome these limitations. Although it is possible to produce single diastereomers of β-hydroxy esters synthetically, it can be challenging and quite expensive due to the use of chiral reagents, enzymes, and additional synthetic steps. Therefore, it is most common to produce the β-hydroxy esters synthetically without diastereomer control with the understanding that nonnatural stereocenter will be present in the final molecular structure. The three major approaches in the chain synthesis are: (1) β-ketoester formation via Meldrum’s acid and a long-chain acid chloride; (2) cross-aldol with long chain aldehyde and an ester enolate; and (3) the Reformatsky reaction. In the former, many methods for the stereoselective reduction of the β-ketoester are possible; whereas, in the latter two approaches, stereospecificity can be accomplished via a chiral auxiliary ligand or the racemic mixtures can be separated postsynthesis. However, for many applications, racemic mixtures of lipids suffice, and therefore do not require the use of stereospecific synthesis. Following the formation of the β-hydroxy lipid moieties, glycosylation is the next synthetic step and it is accomplished via multiple methods. Typically, glycosylation occurs via standard Lewis-acid carbohydrate chemistry. In all cases, the necessity of protecting groups is required for efficient synthesis, regio- and stereoselectivity, and it reduces the overall atom efficiency. For polyrhamnose molecules, adding protecting groups to the sugar is particularly burdensome considering the bioidentical glycosidic linkages and stereoselectivity. As such, multigram synthesis of polyrhamnose species are currently not feasible. The introduction of a process to synthetically produce rhamnolipid was first published in 1988 by van Boom et al. and later updated by the same group in 1998 [17,18]. Although the process outlined was successful, it required more than 18 steps to complete the synthesis of the rhamnolipid, and it was not ideally suited for cost-effectively scaling the process to any significant quantities—the authors reported the possibility of scaling to multigram quantities. Interestingly, this research highlighted the fact that rhamnolipid biosynthetically has only one diastereomer while those produced synthetically have four. Further expansion of the synthetic platform to include additional rhamnolipid molecular structures was described using a novel method known as hydrophobically assisted switching phase (HASP) synthesis, which was demonstrated shortly after by Bauer et al. [19]. The HASP synthesis methodology enables flexible switching between solution-phase steps and solid-supported reaction steps using a hydrophobic silica support. Perhaps the biggest improvement using the HASP method was its ability to synthesize 3-hydroxy fatty acid chains that were diastereomer-selective, thus, overcoming the potential objection to having a mixture of the four diastereomers [19]. The challenge with these previous glycosylation chemistry methods is that they are consistently plagued by low reaction efficiencies ( proteases > glycoside hydrolases. They also differ in their tolerance to reduced water in the media (see subsection reaction media), the substrates each enzyme prefers (see subsection substrate ratio). Another difference arises from the shape of the active site. iCALB binds carbohydrates with at least two different orientations, giving two isomers as products [76]. As for the thermolysin from B. thermoproteolyticus rokko, evidence exists for a restricted cavity accepting only the palmitic acid vinyl esters and no other derivatives (palmitic anhydride, methyl palmitate, tripalmitin) [102]. Differences within an enzyme family also arise. The specificity toward the amidation or hydroxylation with phenylglycinol was tested for four commercially available immobilized lipases: (1) iCALB, (2) Candida sp. 99–125, (3) lipase from Thermomyces lanuginosus, and (4) lipase from Rhizomucor miehei. The best amide/ester product ratio was obtained with iCALB (ratio of wt%, 27) and the worst with lipase from Rhizomucor miehei (ratio of wt%, 0.8). Molecular docking

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simulations pointed to differences in the catalytic site resulting in differential binding of the phenylglycinol acceptor, in which either its hydroxyl or amino group was oriented closest to the acyl–enzyme intermediate. Only for iCALB, phenylglycinol binds with the amino group pointing toward the acylintermediate, whereas in all other enzymes, it binds with the hydroxyl closest to the acyl intermediate.

5.2 The reactions media In water, reactions that form biosurfactants (e.g., esterification, amidation, glycosylation) compete with hydrolysis, which is favored by thermodynamics. Furthermore, the precursors of hydrophobic moieties in surfactants are usually water-insoluble molecules, whose reactivity is limited by mass transfer, as the enzyme usually prefers the aqueous phase. To mitigate these obstacles, water has been substituted by other solvents. Thus free-solvent systems have increased the efficiency of enzymecatalyzed amide or ester formation over hydrolysis. The environmentally friendly DESs are gaining special interest as they support enzymatic catalyzed reactions. DES are composed by mixtures of compounds already in nature, one acting as a hydrogen bond donor and a second acting as hydrogen bond acceptor. Such molecules include choline, urea, glucose, and glycerol, thus arising themselves as greener solvents. Therefore these new solvents have been substituting ionic liquids (ILs) as they arise as greener options. These low fusion temperature compounds have been assayed as solvents in the enzymatic activity of lipases and glycosyl hydrolases [106,107]. Nevertheless, no general solvent exists to favor esterification, transesterification, or amidation reactions over hydrolysis. Hereby, each transformation should be evaluated to establish the conditions for optimal yields of the desired product. In the case of lipases, evolutionary pressure might have adapted these biocatalysts for an efficient function in organic media, as their substrates (TAGs) form a low aw phase. Conversely, other hydrolases such as glycoside hydrolases and proteases, which are used in biosurfactants synthesis, are adapted for working in aqueous media [108]. Although reducing the water content in the reaction media disfavors hydrolysis, enzymes still require the presence of water for activity. This requirement is different for each enzyme. For example, while some glycoside hydrolases require a aw above 0.4 [28]; for lipases, enzyme activity has been observed at aw as low as 0.1 [109]. Not only the dependence of aw is different for each enzyme, but it could follow a different trend. As an example, the esterification of butyric acid and ethanol with CALB performs highest when the aw becomes lowest. This behavior has been explained using molecular dynamics (MD) simulation, which revealed that enzyme mobility is possible thanks to water binding to hotspots that—as the aw increases—occupy sites near the catalytic site favoring hydrolysis and dampening the esterification reaction [109]. These scenarios make it necessary to evaluate enzyme activity at different aw to determine the best solvent composition for each enzyme. When the enzyme tolerates water free systems, residual water (absorbed by the solvent during production and storage) affects the reaction yield. CALB was evaluated for the reaction of vinyl palmitate with glucose; solvents were either tested as received from the suppliers (Condition A) or dried before use (Condition B) [110]. For acetone, tert-butanol, THF, dioxane, and acetonitrile (MeCN), the reaction occurred with conversions between 12% and 52% under Condition A. After drying the solvents (Condition B), the conversion increased above 80%, evidencing the prejudicial effect of residual water in the solvents. Furthermore, when molecular sieves were added to adsorb water during the reaction the yields increased around 10%. The transesterification and hydrolysis rates were also determined. Under Condition B, hydrolysis was observed for all solvents, but for acetonitrile the FA produced was converted into 6-O-Glucosyl-palmitate. Upon addition of the molecular sieves, hydrolysis was reduced and

5 Factors affecting the enzymatic production of biosurfactants

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the complete conversion of the acid into ester was observed for acetonitrile, THF, and acetone. Molecular sieves should be used with caution, though, since in excess, they could stripped enzymes from water, diminishing their activity [81]. An interesting phenomenon was observed in MeCN: 6-Oglucose-palmitate insolubility in this solvent produced its phase separation, displacing the equilibrium to the formation of the surfactant in accordance with the Le Chatelier-Braum principle. The solvent polarity modifies the yield of the surfactant synthesis not only by affecting the solubility of substrates and products, but also by changing the rate of reaction. As a rule of thumb, increasing the solvent hydrophobicity increments the rate for the synthetic reaction. However, even for lipases, this reaction rate declines after reaching an optimum hydrophobicity (measure most of the time as log P). For the esterification with fatty alcohols this optimum is reached at high log P values (Fig. 12). When the acyl acceptor is a polar molecule, such as sugars, AAs, and peptides, the log P optimum falls at lower values of log P (Fig. 13) [114]. The yield of reaction has an optimum in polarity for cases in which it is decreased by the addition of an hydrophobic solvent: subtilisin BPN´-catalyzed acylation of sucrose produced the best results with 20% hexane in a mixture with pyridine; addition of more hexane not just lower acylation yields but also was detrimental for the 10 /6-OH acylation selectivity, which drops to half for 40% hexane [115]. Predictions of the rate of acylation based on the solvent polarity should also consider the substrate solubility as its effect may override that of solvent hydrophobicity. This has been observed with the tripeptide-KHA acylation by iCALB [82], where the best yields were obtained for the solvents in which the tripeptide was the most soluble despite their differences in polarity—e.g., acetonitrile (log P ¼  0.34) yielded 56.9% conversion and 2-methyl-2-butanol (log P ¼ 0.89) yielded 63.30%. In this same line, solvent-free systems have profit from the slightly amphiphilic nature of fatty alcohols and FAs, which also have low polarity; thus for the amidation of

FIG. 12 Correlation of transesterification rates of lipases with solvent log P values: (A) Candida cylindracea lipase and (B) Mucor sp. lipase (replotting from literature data: reaction rates of lipase-catalyzed transesterification of tributyrin and heptanol at 20°C from [111]; log P values from experimental data as presented in [112] except hexadecane and dioxane from calculated data from [113]. Reproduced with permission from Klibanov AM. Why are enzymes less active in organic solvents than in water? Trends Biotechnol 1997;15(3):97–101.

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Chapter 12 The use of biocatalysis for biosurfactant production

100

Conversion (%)

80

60

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) .9 (4 e an 9) ) ct 3. 4 O e ( (3. an e ) n .9 ex a H ex (2 h lo ) de yc 3.1 lori C ( h ne ac le etr Xy n t ) bo 2.6 ( ar C e ) en 2.1 lu ( ) To ne 2.0 e ( 9) nz m 1. r r( Be ofo he t ) or ) l e .2 hl (1 .79 C py 0 ro ne l( a op h t ho Is e or alco hl l ic ty ) D bu 65 r t- 0. ( Te e in rid 6) 9) Py 0.4 0.2 ( ( F ne TH no 23) ta 0. Bu (– 2e on et ) Ac 50 0. (–

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FIG. 13 Effect of solvents on conversion of the transesterification reaction. Experimental conditions: 0.1 mmol D-glucose, 0.4 mmol divinyl butanedioate, 50 mg/mL alkaline protease from B. subtilis, 1 mL solvent. The reaction was carried out at 50°C for 48 h. Pyridine and DMF are outlayers because pyridine is a catalyst in transesterification reactions. Reproduced with permission from Wu Q, Wang N, Xiao Y-M, Lu D-S, Lin X-F. Regiospecific alkaline protease-catalyzed divinyl acyl transesterifications of primary hydroxyl groups of mono- and di-saccharides in pyridine. Carbohydr Res 2004;339(12):2059–67.

phenylglycinol with capric acid, the best result was obtained for the solvent free reaction and the lowest for benzene [116]. The exclusion of water from the enzyme surface can lead to the enzyme loss of flexibility, which not only reduces enzymes’ activity [117,118] but also augments their stability [119] by kinetically preventing its unfolding process [120]. This rigidification has been observed through EPR experiments as in molecular dynamic simulations for the protease subtilisin BPN’, which reveal a greater loss of flexibility with decreasing solvent dielectric constant [121]. Very surprisingly, this rigidification correlates with an increment of the transesterification reaction using subtilisin BPN’. This behavior might be explained by the fixation of water molecules on the enzyme surface in very hydrophobic solvents [122]. The stability of each enzyme varies with the solvent used, but due to the rigidity nonpolar solvents imposed on enzymes, they tend to stabilize these biocatalysts. MD has been used to explain the increment of stability for Bacillus thermocatenulatus lipase (BTL2), in non-polar solvents (toluene and cyclohexane), in terms of its rigidification. In contrast, for water and the alcohols methanol and ethanol, the enzyme’s mobility increases with the temperatures resulting in its denaturalization [123]. However, there have been reports of enzymes that are stable in alcohols, especially as their side chain elongates. For example, tert-butanol has worked as a solvent for transesterification and aminolysis reactions [71]. Some solvents that seem to offer protection against denaturation are DES even when they are made by denaturing agents themselves. The choline:urea DES offers protection against denaturation with

5 Factors affecting the enzymatic production of biosurfactants

287

methanol for the ɑ-amylase AmyA of Thermotoga maritima during the production of methyl glycoside, as demonstrated by an increased in their thermal denaturation temperature (Tm) when DES was present [107]. As in the case of traditional organic solvents, enzymes in DES require water for functioning. In the choline-based DESs, the immobilized lipase from C. antarctica improve both the esterification rate and yield for the synthesis of butyl acetate from acetic anhydride and n-butanol [124]. Although the rate of esterification kept increasing when water was increased, its yield reached an optimum— probably due to the increment of hydrolysis with water content. At this optimum, the yield in DES (70%–80%) surpassed that of the n-heptane (50%) and the imidazolium-based IL [C5mim] [Tf2N] (40%). However, upon addition of water, the half-life time of the enzyme was reduced to 2 days, compared with the anhydrous solvents n-heptane and [C5mim] [Tf2N], in which the residual activity remained unaltered after 15 days. Due to their polar nature, DES become immiscible with FAs and alcohols as their chain elongates. Choline in combination with urea or glucose was also reported for the reaction between glucose and vinyl decanoate catalyzed by iCALB [87]. In the system conformed by choline:urea, the yield was 0.57 μmol/g DES after 24 h of reaction at 50°C. Due to the insolubility of the vinyl decanoate, the reaction yield was not improved after increasing agitation from 60 to 90 rpm in both eutectic solvents. Sonication increased the yield fourfold as it made the vinyl ester more available to react by forming microdroplets. To tackle the solubility problem, more hydrophobic DESs have been designed. For example, Hollenbach et al. [88] reported a ()-menthol:decanoic acid DES for the reaction of glucose with decanoic (or vinyl decanoate) catalyzed by iCALB, which additionally has a viscosity 10-fold lower than the reported for the choline:urea system at 50°C. The decrease in polarity and viscosity, as well as one of the reagents being part of the solvent, substantially incremented the yield to 164.27 μmol/g DES after 24 h of reaction at 50°C.

5.3 Substrate ratio The reactants ratio usually favors the acceptor moiety as it will drive the displacement of the reaction toward the surfactant’s formation. The increase in concentration of the hydrophobic part should be done with caution as its excess might lead to phase-transfer problems and insolubility of the polar moiety of the surfactant; thus the ratio is usually kept below 5:1 [22,82,85,87,116]. In some instances, increments of the acylating agent concentration have inhibited the lipase-catalyzed transesterification carried out in organic solvent [125]; this effect is associated with the formation of nonproductive complexes between the enzyme and this organic molecule [126].

5.4 Time of reaction The velocity of chemical reactions depends on the concentration of reactants, but even in anhydrous starting conditions, the undesired hydrolysis reaction may occur as time advances. That way, the optimum time to obtain the best yield of the product of interest depends on the type of reaction. In general, when the reaction proceeds through reverse hydrolysis, long times are needed to reach the maximum yield established by the thermodynamic equilibrium. On the other hand, when the reaction proceeds through a transfer reaction, to minimize the competing hydrolysis, the reaction time must be limited to obtain the maximum yield. Time might also control the product regioselectivity of the

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reaction, as kinetics and thermodynamics might compete, especially when the reaction is reversible. As an example for the synthesis of phenylglycolamide, different ratios of O–, N–, and diacetyled products were obtained with the diacyl derivative being the representative product for long time of reaction, reaching maximum values at 48 h, while the maximum yield of phenylglycolamide was near to 19 h [116].

5.5 The immobilization of enzymes Enzyme immobilization has been employed to increase enzyme’s productivity. A support compatible with the enzyme can improve its activity and increase its stability to environmental factors such as temperature, pH, salts concentration, and degradation. Furthermore, by allowing its reusability, the overall productivity of the enzyme is augmented. Additionally, by increasing the temperature at which the reaction occurs, the solubility of all substrates increases, as well as the rates of reactions. For example, CALB immobilization in different supports increased the half-life between 5- and 48-fold while maintaining at least 33% of activity [60]. The effect of solid support on reaction yield is directly associated with its chemical nature, for example, the silica matrix is more hydrophobic than epoxy-based supports and its water adsorption is less than for the epoxy derivatives [60]. However, the result of a support with an enzyme depends not only on the nature of the support but also on the nature of the enzyme; the solvent employed for the reaction, and even on the immobilization conditions. For example, for lipases, hydrophobic surfaces on the support matrix favor the exposure of their hydrophobic catalytic site, usually covered by a lid, increasing their activity. In contrast, for these same enzymes, an excess of enzyme could promote the dimerization of enzyme molecules via their hydrophobic catalytic site, rendering inactive dimers, thus decreasing the overall specific activity of the immobilized enzyme [127]. As for the free enzymes, the activity of enzymes on solid supports depends not only on the chosen solvent, but on the aw. This dependency varies from one enzyme to another, as demonstrated by Adlercreutz and Wehtje [128]. In their study, decanoic acid and dodecanol were esterified by three different lipases on a polypropylene EP-100 solid support in diisopropyl ether. The aw ranged from 0.064 to 1. While for Pseudomonas sp. lipase activity diminished with decreased aw, for Rhizopus arrhizus lipase an augmentation in activity was observed after the aw was reduced, with maximum at 0.35. In contrast, Candida rugosa lipase had an optimum of activity for a aw around 0.5. Some reactions not observed for the free enzyme become possible on a support. When immobilized, papain accelerates the esterification rate of alcohols with Boc-alanine, and produced yields up to 85%, with all products present with at least 20% yield, even for substrates not reacting with the free enzyme (Fig. 14). In the same sense, the amidation of Boc-Ala with ethanolamine did not occur with the free papain, in contrast with the 78% yield obtained with the immobilized enzyme [94]. The authors suggested that this effect may be explained by the minimization of water content within the immobilization matrix. In this way, competition of water and alcohol was reduced, favoring the synthesis reaction over hydrolysis. The importance of choosing the right support is highlighted in the esterification of Boc-Asp. It proceeds with diols in larger yields when papain is attached to Sepharose than when it is immobilized in aliphatic adsorbent resin structure, the same support that makes possible the esterification with ethylene glycol with a 40% yield [129] (Fig. 15). This effect on the production of a surfactant is also reflected in changes in the selectivity ratio when the enzyme is immobilized [83] (Table 6).

FIG. 14 Yields of acylation of alcohols with Boc-Ala with free papain (black bars) and immobilized on aliphatic adsorbent resin structure (gray bars). Data was obtained from Cantacuze`ne D, Guerreiro C. Optimization of the papain catalyzed esterification of animo acids by alcohols and diols. Tetrahedron 1989;45(3):741–8.

FIG. 15 Yields of acylation of alcohols with Boc-Asp with papain immobilized on sepharose (black bars) and immobilized on aliphatic adsorbent resin structure (gray bars). Data was obtained from Cantacuze`ne D, Guerreiro C. Optimization of the papain catalyzed esterification of animo acids by alcohols and diols. Tetrahedron 1989;45(3):741–8.

Table 6 Effect of the immobilization in transesterification reaction catalyzed by hydrolytic enzymes. Temperature (yield) Enzyme (immobilization support)

Supported enzyme

Cycles (final yield)

Solvent

Reference

NR

CH2Cl2

[94]

NR

CH2Cl2

[94]

Acetone, propylene carbonate, acetonitrile DMF

[83]

[130]

Product

Free enzyme

Papain (aliphatic adsorbent resin structure) Papain (aliphatic adsorbent resin structure) Protease from Bacillus subtilis (glyoxyl silica)

Boc-Ala ester of mono(di) alcohols (C8-C18) Boc-Ala-CH2CH2OH

37°C (0%–75%) 0°C

37°C (23%–85%) 78°C

Lauroyl glycine

(18.3–23.3%)

(34.2%–42.6%)

NR

Protease from B. subtilis, (S1000SAPTE and S1000STESPM-pHEMA) Subtilisin Carlsberg (pentynyl dextran)

Sucrose 2-O-vinyl adipate

3.24a

6.26 and 31.3a

6 (55% activity)

N-acetyl-L-phenylalanine ethyl ester with different aliphatic and aromatic alcohols N-acetyl-L-tyrosine propyl ester Lauroyl glycine

RT (0%)

RT (42%–91%)

6 (70% activity)

THF

[131]

25°C (92%)

25°C (92%)

2(92%)

[132]

45 or 55°C (4.2%–12.6%)

45 or 55°C (18.4%–49.4%)

NR

NR

NR

12 (50%)

50°C (9.7%)

50°C (1.1%)

NR

1-O-β-D-galactopyranosyl glycerol

40°C (0%)

40°C (27%–34%)

NR

1-Propanol/5% water Acetone, propylene carbonate, acetonitrile 85% Butan2-ol, 15% aqueous phase Aqueous/ alcohol Free solvent

Butyl-β-galactoside

25°C (0.58 mol/mol)

25°C (0.76 mol/mol)

10 (35%)

Trypsin (commercial silica gel) Lipase from Pseudomonas stutzeri

β-glucosidase from Fusarium solani (chitosan) β-Xylosidase from S. sclerotiorum (Celite 545) β-Galactosidase from A. oryzae (Duolite S-761) β-Galactosidase from A. oryzae (glyoxyl-agarose)

sec-Butyl-β-D-glucoside sec-Butyl- β-Dcellobioside Alkyl-β-glycoside

70% (v/v) of 1-butanol

[83]

[133]

[43]

[135]

Lipase from Rhizomucor miehei (ion exchange resin, sn1,3) Lipase from Thermomyces lanuginosa (acrylic resin) iCALB

Mixtures of avocado oil caprilates

NR

30°C (7.5% mol)

NR

Solvent free

[136]

Mixtures of avocado oil caprilates

NR

30°C (30% mol)

NR

Solvent free

[136]

n-Butyl lactate

NR

NR

n-Butyl lactate

NR

[137]

Candida rugosa lipase (porous cross-linked polystyrene resin beads (NKA)) C. rugosa lipase (porous cross-linked polystyrene resin beads (NKA)) C. rugosa lipase (porous cross-linked polystyrene resin beads (NKA)) C. rugosa lipase (porous cross-linked polystyrene resin beads (NKA)) C. rugosa lipase (sol-gel matrix) Surfactant coated C. rugosa lipase (sol-gel matrix) iCALB

Phytosterol and diglyceride esters

50°C (68.6%)

NR

Supercritical fluoromethane Supercritical CO2 n-Hexane

[137]

iCALB

55°C (88.2%) 55°C (75%) 50°C (92.4%)

Phytosterol and diglyceride esters

50°C (68.6%)

50°C (89.7%)

NR

Iso-octane

[138]

Phytosterol and diglyceride esters

50°C (68.6%)

50°C (75.4%)

NR

Cyclooctane

[138]

Phytosterol and diglyceride esters

50°C (68.6%)

50°C (80.6%)

NR

tert-Amyl alcohol

[138]

Sucrose-6-acetate

45°C (5%) 45°C

45°C (60%) 45°C

DMSO

[139]

DMSO

[139]

NR

86°C (90%) 60°C (52.5%) 45°C (67%)

6 (57.81%) 6 (