Formation and Control of Biofilm in Various Environments 9811522391, 9789811522390

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Hideyuki Kanematsu Dana M. Barry

Formation and Control of Biofilm in Various Environments

Formation and Control of Biofilm in Various Environments

Hideyuki Kanematsu Dana M. Barry •

Formation and Control of Biofilm in Various Environments

123

Hideyuki Kanematsu Department of Materials Science and Engineering National Institute of Technology (KOSEN) Shiroko-cho, Suzuka, Mie, Japan

Dana M. Barry Department of Electrical and Computer Engineering Clarkson University Potsdam, NY, USA

ISBN 978-981-15-2239-0 ISBN 978-981-15-2240-6 https://doi.org/10.1007/978-981-15-2240-6

(eBook)

© Springer Nature Singapore Pte Ltd. 2020 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Singapore Pte Ltd. The registered company address is: 152 Beach Road, #21-01/04 Gateway East, Singapore 189721, Singapore

This book is dedicated to Hideyuki Kanematsu’s father, Shoji Kanematsu; mother, Michiko Kanematsu; wife, Reiko Kanematsu; daughter, Hitomi Kanematsu; and son, Hiroyuki Kanematsu. This book is also dedicated to Dana M. Barry’s husband, James F. Barry; her four sons: James D. Barry, Brian P. Barry, Daniel T. Barry, and Eric M. Barry; and her parents, Daniel and Celia Malloy.

Preface: A Message from the Authors

Five years have already passed since we first wrote a biofilm book entitled Biofilms and Materials Science. Time flies. You can confirm the drastic advancement of biofilm research and investigations during that time period. Therefore, major changes exist between our two books. We needed to write this new book to introduce new and exciting information for you. The outline of our book’s contents is described in Chap. 1. Here in the preface, we would just like to write about the basic concept of this new book. The essence of our book is to provide broader applications coupled with deeper basic theories for biofilms. We describe in detail the phenomena in industrial fields, as well as those in our human bodies and in nature. This allows the readers to have a better and fuller understanding of biofilms, and to get information for their commonalities and differences. Since biofilm research has developed tremendously in these past 5 years, we tried to include many new, exciting, and important developments in this book. We hope that readers will get useful and innovative ideas from our book that may result in great achievements for the current and future scientists and engineers. Also we want to inspire and deepen the understanding and learning for students in the areas of biofilm-related studies. We have collaborated together on various research and educational topics for 20 years. We feel that such a long-lasting collaboration became possible by keeping the belief as follows. Take risks with visions and you will find good opportunities! We sincerely wish you great visions and opportunities. Well, let’s use this guide book to explore the world of biofilms together.

Suzuka, Japan Potsdam, USA

Very Best Wishes for the Present and the Future Hideyuki Kanematsu Dana M. Barry

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Acknowledgements

We would like to thank the National Institute of Technology (KOSEN), NIT (KOSEN) Suzuka College in Japan and its Late President Prof. Junichi Yoshida; Former Chair of Clarkson University’s Department of Electrical and Computer Engineering Prof. David Crouse; the current Chair of that Department Professor Paul McGrath; Clarkson University and its Department of Electrical and Computer Engineering; the State University of New York at Canton (SUNY Canton); Executive Vice President of Osaka University Prof. Toshihiro Tanaka; Prof. Eiji Arai of Osaka University; Osaka University in Japan; the American Chemical Society; Prof. Dr. Roger Haw; and Ansted University, for their greatly appreciated support. We are grateful to the Iketani Science and Technology Foundation. They made it possible for us to discuss the book in person. We thank ISIJ (Iron and Steel Institute of Japan), the Iron and Steel Society of Japan, ASM International, and SIAA (the Society of International Sustaining Growth for Antimicrobial Articles) and that Society’s Biofilm Evaluation Committee. We thank the great surface finishing societies in the United States (National Association for Surface Finishing, NASF), the United Kingdom (Institute of Materials Finishing, IMF) and Japan (Surface Finishing Society of Japan, SFSJ). Some information contained in this book was obtained from the work carried out for national funding projects in Japan. We also express our sincerest appreciation to Springer Nature Singapore and to the Editor Dr. Mei Hann Lee. In addition, we would like to thank our families for their continued interest and support: Dr. Kanematsu’s parents (Shoji and Michiko Kanematsu), his wife (Reiko), and children (Hitomi and Hiroyuki); Dr. Barry’s parents (Daniel and Celia Malloy), her husband (James), and children (James, Brian, Daniel, and Eric). Hideyuki Kanematsu, Ph.D. Dana M. Barry, Ph.D.

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Contents

1 5

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2

Fundamentals for Biofilms . . . . . . . . . . . . . . . . . . . . . 2.1 Bacteria and Biofilms . . . . . . . . . . . . . . . . . . . . . . 2.2 General Sketch for Biofilm Formation and Growth . 2.3 Biofilm Constituents . . . . . . . . . . . . . . . . . . . . . . . 2.4 Exopolymeric Substances . . . . . . . . . . . . . . . . . . . 2.5 Quorum Sensing . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Biofilm Collapse and Removal . . . . . . . . . . . . . . . 2.7 Biofilm and Infection . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Animate Substrata and Biofilms . . . . . . . . . . . . . . . . . . 3.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.1 Biofilm on Leaves . . . . . . . . . . . . . . . . . . . 3.3 Biofilms on Animal (and Human) Tissues . . . . . . . . 3.3.1 What Happens Inside of Our Bodies, When Pathogenic Bacteria Enter Them? . . . . . . . . 3.3.2 The Characteristics of Bacteria in Biofilms . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Biofilms in Nature and Artificial Materials . 4.1 Natural Substrates . . . . . . . . . . . . . . . . . 4.2 Artificial Substrates . . . . . . . . . . . . . . . . 4.3 Metals . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Ceramics . . . . . . . . . . . . . . . . . . . . . . . 4.5 Polymers . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

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Laboratory Biofilm Reactors . . . . . . . . . . . . . 5.1 In Vitro Systems . . . . . . . . . . . . . . . . . . . 5.2 Static Systems . . . . . . . . . . . . . . . . . . . . 5.3 Flow Systems . . . . . . . . . . . . . . . . . . . . . 5.4 Quasi-natural Systems (Ex Vivo Systems) 5.5 In Vivo Systems . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Detection and Evaluation of Biofilms . . . . . . . . . . . . . . . . . 6.1 Biological Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1.1 Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1.2 Gene Analysis . . . . . . . . . . . . . . . . . . . . . . . . . 6.1.3 Proteomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Instrumental Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.1 Optical Microscopes . . . . . . . . . . . . . . . . . . . . . 6.2.2 Fluorescence Microscopes . . . . . . . . . . . . . . . . . 6.2.3 Confocal Laser Scanning Microscope . . . . . . . . 6.2.4 Scanning Electron Microscope (SEM)— Energy-Dispersive X-Ray (EDX) Spectroscopy . 6.2.5 Transmission Electron Microscope (TEM) . . . . . 6.2.6 Atomic Force Microscope (AFM) . . . . . . . . . . . 6.2.7 Ultraviolet-Visible (UV-VIS) Spectroscopy . . . . 6.2.8 White Light Interferometer . . . . . . . . . . . . . . . . 6.2.9 Fourier-Transform Infrared Spectroscopy (FTIR) 6.2.10 Raman Spectroscopy . . . . . . . . . . . . . . . . . . . . 6.2.11 Nuclear Magnetic Resonance (NMR) . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Standardization—Current and Future . . . . . . . . . . . . . . . . . . 7.1 The United States . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.1 ASTM E2196 . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.2 ASTM E2562 . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.3 ASTM E2647 . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.4 ASTM E2871 . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.5 ASTM E2799 . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.6 ASTM E3151 . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.7 ASTM E3161 . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1.8 Characteristics of the ASTM Standard in the USA . 7.2 European Union . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.1 European Standards (EN) . . . . . . . . . . . . . . . . . . . 7.2.2 Phase 1—Basic Suspension Test . . . . . . . . . . . . . . 7.2.3 Phase 2 Part 1—Quantitative Suspension Test . . . . 7.2.4 Phase 3—Field Test . . . . . . . . . . . . . . . . . . . . . . . 7.2.5 International Standards (ISOs) . . . . . . . . . . . . . . . .

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7.3 Japan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 168 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 8

Biofilm Problems and Environments . 8.1 Marine Environments . . . . . . . . . 8.2 Soil Environments . . . . . . . . . . . 8.3 Household Environments . . . . . . . 8.4 Food Processing Industries . . . . . 8.5 Pipes and Heat Exchangers . . . . . 8.6 Hospital and Medical Fronts . . . . References . . . . . . . . . . . . . . . . . . . . .

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Biofilm Usefulness . . . . . . . . . . . . 9.1 Energy Applications . . . . . . . 9.2 Environmental Applications . . 9.3 Water Treatment Applications References . . . . . . . . . . . . . . . . . .

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10 Biofilm Control and Thoughts for the Future . . . . . . . . . . . . . . . . . 223 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228

About the Authors

Dr. Hideyuki Kanematsu, Ph.D. FIMF (a Fellow of IMF) is a Full Professor in the Department of Materials Science and Engineering, at the National Institute of Technology (NIT) (KOSEN), Suzuka College in Japan. After holding many administrative positions in the college (Dean of the Department of MS and E, 2010–2014, Deputy President, 2014–2018, etc.), he is still a researcher in Materials Surface Science and Engineering and the research advisor for the college. Dr. Kanematsu is particularly interested in biofilms and the interfacial phenomena between metallic surfaces and organisms from an environmental science point of view. He holds a B.Eng. (1981), a M.Eng. (1983) and a Ph.D. in Materials Science and Engineering (1989) all from Nagoya University. He is a member of the National Association for Surface Finishing (NASF, the USA), as well as the Institute of Metal Finishing (IMF, the UK), the Minerals, Metals and Materials Society (TMS, the USA), ASM International (the USA), Electrochemical Society (ECS, the USA), American Chemical Society (ACS, the USA), the Japan Institute of Metals (JIM) and the Iron and Steel Institute of Japan (ISIJ), Surface Finishing Society of Japan (SFSJ, Japan), the Electrochemical Society of Japan (ECSJ), the Japan Society for Heat Treatment (JSHT), Japan Thermal Spray Society (JTSS), etc. For all of these organizations, he is still very active as a member and also as a board member. He also has many honors and publications to his name. He was awarded a Marquis Who’s Who Lifetime Achievement award for his Outstanding Career and is an NASF Scientific Achievement Award Winner. Dr. Dana M. Barry, Ph.D. is a Research Professor in Clarkson University’s Department of Electrical and Computer Engineering. She also serves as a Professor and Scientific Board President for Ansted University, an Instructional Support Assistant for the State University of New York at Canton (SUNY Canton) and as an officer and Chemistry Ambassador for the American Chemical Society. She previously worked as a Chemical Engineer, a Chemical Consultant, Senior Technical Writer and Editor at Clarkson University’s Center for Advanced Materials

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About the Authors

Processing and as a Host and Co-Producer for the television series Sensational Science (that aired in Northern New York State for several years). She has served as a Visiting Professor overseas numerous times. She also presented invited Keynote Talks in the following countries: Malaysia (2001), Japan (2008), China (2011), Slovenia (2014), and Hungary (2019). She has five graduate degrees including a Ph.D. in Engineering from Osaka University in Japan. In addition, Dr. Barry has many honors and close to 300 professional publications. Her special awards include a Citation of Excellence for her Keynote Speech in Budapest, Hungary (2019); a Marquis Who’s Who Lifetime Achievement Award for her Outstanding Professional Dedication and Accomplishments (2017); an Outstanding Volunteer Award from the American Chemical Society (2017) and 21 Consecutive APEX Awards for Publication and Communication Excellence sponsored by Communication Concepts in VA (1996–2016).

Chapter 1

Introduction

Abstract Biofilms form at the interface between different phases as a result of bacterial activities. They cause many problems for industry and our daily lives. Biofilms have negative problems to solve, but also provide us with benefits if we use them properly and effectively. In this chapter, we outline a world filled with biofilms and their impact on industries and our daily lives. Also we mention why we were compelled to write this important book.

Biofilms form at the interface between different phases by bacterial activities. They affect many industries as well as our daily lives. Figure 1.1 shows problems caused by biofilms and the various fields affected by them [1]. For example, biofilms cause corrosion, scale buildup, etc. for pipes, plumbing, and the shipping industry. They are also responsible for contamination in medical settings, food industries, and in a variety of other environments. Corrosion is a form of materials’ degradation due to the anodic dissolution of metals [2]. For example, steels corrode as a result of the dissolution of iron into the divalent iron ion, as shown in Fig. 1.2. Why would corrosion be related to biofilms? Many mechanisms have been proposed. However, they continue to be controversial [3–14]. The problem has not been solved yet. We would like to say that biofilms assist and accelerate corrosion phenomena of metals. In aqueous solution environments, biofilms form on metallic materials’ surfaces and provide active reaction sites for corrosion. They might produce oxygen Fig. 1.1 Biofilms and their affected fields

© Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_1

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1 Introduction

Fig. 1.2 Corrosion of metals: the case of iron and steel for example

Fig. 1.3 Corrosion of metallic materials in aqueous solutions and biofilms

Fig. 1.4 Water films and corrosion of metals in atmospheric environments [15, 16]

concentration cells to induce corrosion. Also biofilms could accelerate Redox reactions, where metallic ions would interact with proteins and other polymers like EPSs (Extracellular polymeric substances) excreted from bacteria in biofilms (Fig. 1.3). In the case of atmospheric corrosion, we already know that the existence of thin water films would be needed to start the corrosion process. When the water film thickness is smaller than 1 µm, corrosion would be hard to start. However, it would accelerate the corrosion phenomenon at a thickness of around a couple of micrometers. When the water film thickness is larger than this order, the delay of oxygen dissolution would lead to a decrease in the corrosion rate. However, corrosion takes place at a thickness of a couple of micrometers because this size is close to a bacterial size which reminds us of biofilms. Biofilms contain water and could serve as the thin water films themselves (Fig. 1.4).

1 Introduction

3

Keep in mind that pipes and plumbing, with liquids flowing through them, may contain biofilms on their inner walls. At the early stage of investigation (before the 1970s or even in the1980s), people did not realize the scale buildup problems inside of their pipes and plumbing. However, the beginning of such a problem is the attachment of bacteria that is followed by biofilm formation and growth [17–30]. In the food industry, factors such as sanitation, antisepsis, and sterilization are very important. However, the existence of biofilms sometimes blocks these processes and brings about troublesome problems such as food poisoning and others [31–45]. As for medical areas, biofilms are responsible for chronic diseases [46–53] and for nosocomial diseases [54–65]. Since these problems relate to our daily lives, they should be solved as soon as possible. Ships in marine environments often have organisms like oysters and barnacles attached to them (a form of biofouling called macrofouling). This problem could cause ships to use more fuel and to have a longer travelling time to arrive at their destinations. This phenomenon of biofouling is followed by the formation of biofilms, which are considered to be a form of microfouling. These problems need to be solved from the viewpoint of biofilms [66–74]. Biofilms form in many places of the living environment. They can be found in dining and kitchen areas [75–79], toilets/bathrooms [80–90], buildings [91–94], air conditioners [95, 96], laundry machines [97–100] and in some other household items. So far, only negative aspects and problems have been mentioned for biofilms. On the other hand, biofilms have a positive side if used properly and effectively. Biofilms are being used to produce energy in microbial fuel cells and more [101–109]. Also our understanding of the electrochemical characteristics of biofilms continues to grow [110]. The authors have carried out energy projects with Japanese companies in the past. Some projects were funded by NADO (the National Association of Development Organizations) in Japan. The first opportunity involved a project for the generation of solar power. At that time, there was a need to improve the efficiency of electric power generation, since it was hampered and degraded by the fog phenomenon of mirrors reflecting sunlight. We presumed that the degradation was caused by biofilms that formed on the mirrors, so we tackled with some related problems [111]. The second project for NEDO was the generation of geothermal energy. For this project, the scale problem inside the pipes where high temperature groundwater was pumped up (from below the ground) was an issue. This problem also degraded the power generation efficiency. We tried to solve the problem by inventing a new system with the application of alternate electromagnetic waves [112–118]. Environmental reclamation by biofilms is another promising positive application. Biofilms absorb heavy metals and lots of organic and inorganic matter [119]. By using biofilms, we might get a chance to improve many environments. Bioremediation is another application for biofilms [120–129]. Ultimately, the control of biofilms might lead to the control of carbon dioxide and improve the global warming situation [130].

4

1 Introduction

As outlined above, biofilms form in various environments. They are actually very versatile. It seems like the biofilm formation and growth process changes from one environment to another. However, biofilms have a common and essential point for their formation and growth process. Therefore, there should be a certain common way to control them. In this book, we introduce the readers to many examples of biofilm formation and growth processes in various environments. At the same time, we discuss the advantages and disadvantages of biofilms and suggest ways to control them. The authors are mainly chemists and material scientists (especially for the interfaces among heterogeneous phases). Therefore, the book is summarized and integrated in such a way. The book chapters are set up as follows. In this first Chapter, the topic of biofilms is introduced. Also some problems and benefits of biofilms are presented and discussed along with suggested approaches for controlling them. Chapter 2 provides information about the fundamentals for biofilms such as their constituents and a description of their formation and growth process. Chapter 3 includes a discussion about biofilms in natural environments. The main focus is about the growth and effects of biofilms on plants and on tissues of the human body. Chapter 4 describes biofilms from the viewpoint of substrates. As we explained in our former book [1], people sometimes forget that biofilms are formed on substrata and that the combination of bacteria and substrata are very important. Therefore, in regards to solving problems that arise, both environmental and materials’ (substrata) viewpoints should be considered. Chapter 5 presents and describes a variety of laboratory biofilm reactors (LBR). Researchers and engineers design these reactors in order to produce biofilms artificially, rapidly, and with good reproducibility. These studies are needed in addition to natural exposure tests. Chapter 6 discusses various techniques for detecting and evaluating biofilms. They include biological methods such as staining and instrumental analysis. Chapter 7 introduces and describes current standards and tests (relating to biofilms) for the United States, the European Union, and Japan. Suggestions are also provided for the standards of the future. Chapter 8 focuses on the relationship between biofilms and environmental problems for marine, soil, and household environments, etc. Chapter 9 discusses useful applications of biofilms in regards to energy, the environment, and water treatment methods. Chapter 10 emphasizes the importance of science and engineering knowledge and tools for solving current problems to produce a better future. It also mentions challenging problems and topics that need to be solved in the near future. The book Formation and Control of Biofilm in Various Environments is a special publication and guide for professionals and students involved with science and engineering related fields. It includes current information and discussions about leading edge research for biofilms, as well as numerous references. Biofilms are everywhere, so we invite you to use this book as an informative, tourist guide for a trip with us to the world of biofilms. Bon Voyage!

References

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Chapter 2

Fundamentals for Biofilms

Abstract This chapter includes the fundamentals for biofilms. It starts by introducing the topics of bacteria and biofilms. Then it discusses the formation, growth, collapse, and removal of biofilms, which are the result of bacterial activity. Biofilm constituents (which are predominantly water) and exopolymeric substances (EPS) are also presented. EPS, a sticky slime, is the main component of a biofilm’s threedimensional structure and includes proteins, polysaccharides, lipids, and more. Quorum sensing, which is cell to cell chemical communication that allows bacteria to coordinate an activity, is described too. In addition, details are provided about biofilm involvement in a wide variety of infections.

2.1 Bacteria and Biofilms Bacteria and biofilms have an interesting relationship. Biofilms are actually the result of bacterial activity. BACTERIA: Bacteria are considered to be prokaryotic microorganisms. These organisms lack a cell nucleus or any membrane-encased organelles. Bacteria were among the first life forms found on Earth and are present in most of its habitats. They exist in the mouth and on the skin of humans. Generally speaking the bacterial cell is surrounded by a cell membrane, which is primarily made of phospholipids [1]. This membrane encloses the contents of the cell. It acts as a barrier to hold the nutrients, proteins, etc. of the cytoplasm within the cell. Most bacteria have genetic material that is a single bacterial chromosome of DNA located in the cytoplasm. Like other living organisms, bacteria contain ribosomes for the production of proteins. They also have a protective cell wall around their plasma membrane, and some have a capsule (an additional wrapping around that cell wall). The capsule helps disease-causing bacteria evade a human’s protective immune system. Many bacteria have surfaces covered in short hair-like projections called pili (which help them link together or to attach to a surface). Some have one or more whip like structures (flagella) which help them move through fluids. See Fig. 2.1 [2]. It shows a bacterial cell with pili and a flagellum.

© Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_2

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Fig. 2.1 A bacterial cell displays pili and a whip-like tail called flagellum

Bacteria have a variety of shapes and sizes, with a typical length of 0.5–5.0 μm. Most bacteria are either spherical (cocci) or rod-shaped (bacilli). Others are vibrio (like slightly curved rods), spirilla (spiral-shaped) or spirochaetes (tightly coiled). These shapes can influence bacteria’s ability to acquire nutrients, attach to surfaces, etc. Many bacteria exist as single cells, while others tend to group together. For example, streptococcus forms chains and staphylococcus (which causes skin infections) joins together to look like a bunch of grapes. Bacteria can be classified into two large groups (gram-positive and gram-negative) by using crystal violet. The procedure used is called the Gram stain test, which was developed in 1884 by Hans Christian Gram, a Danish Biologist. Gram-positive bacteria give a positive result in the Gram stain test. They take up the crystal violet stain (used in the test) and appear purple when looked at under a microscope. See Fig. 2.2 [3]. Gram-positive bacteria possess a thick peptidoglycan layer. Gram-negative bacteria do not retain the crystal violet used in the test. They contain a thin cell wall

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Fig. 2.2 The purple rods seen in this Gram stain test represent Gram-positive bacteria for anthrax bacilli

consisting of a few layers of peptidoglycan surrounded by a second lipid membrane containing lipopolysaccharides and lipoproteins. These bacteria are a medical challenge because their outer membrane protects them from many antibiotics. BIOFILM: Bacterial activity produces biofilms. What are biofilms? There is no exact definition of biofilm. It varies from researcher to researcher. However, it can be described as a tower-like inhomogeneous film on materials’ surfaces. In nature, microorganisms such as bacteria may exist as single cells that float or independently swim in a liquid. Also they may attach to each other and/or to some surface, especially to one that is moist and has nutrients. This aggregation of microbes is called a biofilm. It is a slimy, sticky layer that encases the bacteria that produced it [4–6]. The biofilm is mostly water and contains polysaccharides, proteins, etc. Figure 2.3 shows a Staphylococcus aureus biofilm on an indwelling catheter [7]. Several examples of biofilms include slime on a rock in a river and dental plaque on our teeth (which over time hardens to form tartar). The first founder of biofilms might be Leeuwenhoek, who invented microscopes in the 17 the century, because he clearly observed biofilms from dental plaque [8]. Biofilms, which mainly consist of water, are more than just slimy layers of bacteria. They are actually biological systems because the bacteria (as a result of cell to cell chemical communication) are organized into a functional community. Bacteria, encased in biofilms, are protected from antibiotics and display resistance to them. Biofilms are negatively associated with human bacterial infections, the clogging of pipes, dental problems, etc. On the other hand, the bacteria within them are useful for animal digestive systems, sewage treatment plants, the production of beverages through fermentation, and more.

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Fig. 2.3 Biofilm can be seen on an indwelling catheter

2.2 General Sketch for Biofilm Formation and Growth A biofilm generally starts to form when a free-swimming (planktonic) bacterium attaches to a surface, typically one that is moist and contains nutrients like organic matter. In addition to nutrients, other factors such as electricity, magnetism and light may influence the movement of free floating bacteria. It is not so easy for bacteria to attach to materials’ surfaces because they must overcome obstacles such as flow, surface roughness, hydrophobic/hydrophilic conditions, etc. Sometimes bacteria attach to material and then leave the surface. This is because the initial adhesion may be through weak reversible forces like van der Waals forces. For a while, the bacteria go through an attachment/detachment process until some eventually stay on the surface. When the bacteria population reaches a certain value, they simultaneously secrete an extracellular polymeric substance (EPS) (EPS is a sticky, slimy matter that binds the bacteria to each other and anchors them to a material’s surface.). This bacterial activity is turned on as a result of cell to cell communication. The aggregation of these microbes is called a biofilm. Biofilms encase and protect bacteria from antibiotics, detergents, and other harmful factors in the environment. Once colonization has begun, the biofilm grows by cell division and recruitment. There are five main stages of biofilm development [9]. See Fig. 2.4 [10]. The first stage is the initial attachment, which is followed by irreversible attachment. The third step is initial maturation, with the fourth step being advanced maturation. The final step is dispersion, which enables biofilms to spread and colonize new surfaces. This is an essential phase of the biofilm life cycle.

2.3 Biofilm Constituents

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Fig. 2.4 Five stages of biofilm formation: (1) initial attachment; (2) irreversible attachment; (3) initial maturation; (4) advanced maturation and (5) dispersion (each stage is paired with a photomicrograph of developing Pseudomonas aeruginosa biofilm)

2.3 Biofilm Constituents Biofilms are not easy to define because they differ in structure and composition depending on their environment. However, their main component is water, which can make up to 90% of their wet weight [11]. Biofilms also have water channels to allow for the delivery of nutrients and the removal of waste products. They are heterogeneous environments that include an organized aggregate of microbial cells. The aggregate of microorganisms (such as bacteria) lives within a self-produced extracellular polymer matrix that contains polysaccharides, proteins, minerals, etc. This matrix of extracellular polymeric substances (EPS) forms a sticky matter that binds the bacteria to each other and anchors them to a surface (substrate). Figure 2.5 shows a sketch of a biofilm [12].

2.4 Exopolymeric Substances Exopolymeric substances (EPS) are a main component of a biofilm’s threedimensional structure. They form in the attachment stage of a biofilm to a surface. Usually the thickness of the EPS matrix is 0.2–1.0 μm [13]. Generally a large portion of the EPS is hydrated and may contain nutrients and elements from the surrounding environment. Hydrophobic EPS such as cellulose can occur too. The extracellular

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Fig. 2.5 This sketch describes a biofilm

polymeric substances establish the functional and structural integrity of biofilms and help determine their physiochemical properties. Some possible components of the EPS matrix include proteins, polysaccharides, nucleic acids, and lipids. These components vary depending on the type of bacteria, the stage of biofilm development, the environment, etc. Using various bacteria, studies have been carried out to determine the components of a biofilm’s extracellular matrix. One study involved the grampositive spore-forming bacterium Bacillus subtilis, which is found in soil. Figure 2.6 displays a TEM micrograph of a B. subtilis cell [14]. Its EPS matrix includes polyDL-glutamic acid (PGA) and the proteins TapA, TasA, BslA [15]. This matrix also contains an exopolysaccharide which is partially composed of poly-N-acetyl glucosamine (PNAG), a widely conserved bacterial polysaccharide [16]. In another study, Pseudomonas aeruginosa was used for biofilm formation. An extracellular matrix was produced that included polysaccharides, extracellular DNA (eDNA), lipids (mainly rhamnolipids), and proteins [17]. The polysaccharides found were Psl, Pel, levan, and alginate. Pel has been studied extensively and found to lend a protective role against aminoglycoside antibiotics [18]. The DNA in this matrix contributes to structural stability and induces antibiotic resistance [19, 20]. A general description of possible EPS matrix components is provided. PROTEINS: Like other living organisms, bacteria contain ribosomes for the production of proteins. Proteins are large, versatile biomolecules. Many proteins are classified as enzymes, which act as catalysts for biological reactions. They also perform

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Fig. 2.6 This is a display of a TEM micrograph of a B. subtilis cell in cross-section with a scale bar equal to 200 nm

many functions within organisms including catalyzing metabolic reactions, responding to stimuli, providing structure to cells and organisms, transporting molecules from one location to another, and DNA replication. They are needed in our diets too, especially for building tissues. Proteins are the main characters within the cell and carry out duties specified by information encoded in genes. They can bind to other proteins as well as to small molecule substrates. They are also involved in the process of cell signaling. In regards to bacteria, it has been estimated that the average-sized bacteria contain about two million proteins per cell (examples are E. coli and Staphylococcus aureus) [21]. TasA is the major protein found in the matrix of B. subtilis biofilms. It polymerizes into amyloid-like fibers that are resistant to degradation [22]. These fibers are used by B. subtilis to build a network that connects cells and may organize the rest of the components of the extracellular matrix. The accessory protein TapA (also found in the matrix) is a minor component of the fibers and is required to anchor the TasA fibers to the cell surfaces. Figure 2.7 shows the protein myoglobin, an iron containing protein in muscle [23]. It receives oxygen from red blood cells and transports it to the mitochondria of muscle cells, where oxygen is used in cellular respiration to produce energy. Proteins are mostly linear polymers that are built from a series of up to twenty different amino acids. They can be hundreds of amino acids long. Proteins differ from one another mainly in their sequence of amino acids, which is genetically controlled. Generally speaking an amino acid contains carbon with the following attached groups: an amine group (R-NH2 ), a carboxylic group (R-COOH), and a side chain. Individual amino acids are bonded together by

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Fig. 2.7 This figure represents the 3D structure of the protein myoglobin, an iron containing protein in muscle. Turquoise α-helices are displayed along with a heme group (in gray) with a bound oxygen molecule (in red)

peptide bonds. The peptide bond contains the following elements: carbon, hydrogen, oxygen, and nitrogen. Figure 2.8 provides a detailed diagram of a peptide bond [24]. POLYSACCHARIDES: The EPS matrix includes polysaccharides. Polysaccharides are constructed from smaller molecules known as monosaccharides or simple sugars. They are all carbohydrates because they contain carbon, hydrogen, and oxygen. An example of a monosaccharide is the simple sugar glucose (C6 H12 O6 ). Polysaccharides usually provide structure or storage-related functions in living organisms. For example, starch (a polymer of glucose) is used as a storage polysaccharide in plants that is available as a source of energy. Cellulose and chitin represent structural polysaccharides. Cellulose is used in the cell walls of plants and other organisms. Chitin has a similar structure, but has nitrogen-containing side branches. Figure 2.9 shows a three dimensional structure of cellulose [25].

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Fig. 2.8 This figure shows the chemical structure of the peptide bond (within the bottom rectangle) and its three-dimensional structure (top insert) between an alanine (small amino acid for protein construction) and an adjacent amino acid

Fig. 2.9 This is a 3D (ball and stick) structure of the polysaccharide cellulose

Pathogenic bacteria often produce a thick, mucous-like layer of polysaccharide over the bacterial surface to protect it from destruction. These polysaccharides are water-soluble and commonly acidic. They are linear and contain regularly repeating subunits of one to six monosaccharides. Structural diversity exists, so nearly two hundred different polysaccharides are produced by E. coli alone. Bacteria and other microbes also secrete polysaccharides to help them adhere to surfaces and to prevent them from drying out.

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Fig. 2.10 The structure of N-acetyl glucosamine is shown above

Cell-surface polysaccharides play various roles for bacterial physiology and ecology. They serve as a barrier between the cell wall and the environment. They also form structural components of biofilms and mediate interactions between a host and pathogen. Lipopolysaccharide (LPS) is an important cell-surface polysaccharide. It is a complex molecule that contains lipid and polysaccharide parts. It is found in the outer membrane of Gram-negative bacteria. LPS contributes to the structure of the bacteria and protects the membrane from attack by certain kinds of chemicals. Poly-N-acetyl Glucosamine (PNAG) has been identified as a major polysaccharide component of the Bacillus subtilis biofilm matrix [26]. PNAG production is essential for the formation of Bacillus subtilis biofilm. The structure of N-acetyl glucosamine is shown in Fig. 2.10 [27]. Biofilm formed by Pseudomonas aeruginosa contains the polysaccharides Psl, Pel, levan, and alginate. Psl is the predominant polysaccharide found in biofilm infections in clinical and environmental settings. It acts as an adhesion and plays an important role in the formation and maintenance of the biofilm matrix of P. aeruginosa [28]. In nature, levan is synthesized from sucrose by fructose (two monosaccharides). Bacterial levans are commonly branched. The shortest levan (6-kestose) is basically a chain of two fructose molecules and a terminal glucose molecule. NOTE: The chemical formulas for fructose and glucose are the same (C6 H12 O6 ). Alginate (which is also called algin or alginic acid) is a polysaccharide present in the cell walls of brown algae. It is also a significant component of the biofilms produced by the bacterium P. aeruginosa, which is a major pathogen in cystic fibrosis. NUCLEIC ACIDS: A nucleic acid is a large molecular chain that is typically associated with the storage and transfer of genetic information [29]. A pair of intertwined nucleic acids forms the double-helical deoxyribonucleic acid (DNA) that stores genetic information and ribonucleic acid (RNA) that is involved in protein synthesis. Nucleic acids are constructed from nucleotides which contain an inorganic phosphate (PO4 ) group, a cyclic monosaccharide (or sugar), and a nitrogenous base. If the sugar of the nucleic acid is ribose, the polymer is RNA (ribonucleic acid). If the sugar is derived from ribose as deoxyribose, the polymer is DNA (deoxyribonucleic acid). The alternating sugar and phosphate groups are very important to the structure of a nucleic acid. Uracil (U), guanine, (G), adenine (A), and cytosine (C) are the nitrogenous bases that appear in RNA. The bases that make up DNA

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are guanine, adenine, cytosine, and thymine carried. Most bacteria have genetic material that is a single bacterial chromosome of DNA located in the cytoplasm. Nucleic acids are important components of NTHi biofilm matrices (Non-typeable Haemophilus influenzae). NTHi is a Gram-negative bacterium that often colonizes the human nasopharynx. It can form biofilms in vitro, producing DNA, etc. The extracellular DNA produced seems to be essential in biofilm maintenance, while the extracellular RNA seems to be required only in the first steps of biofilm formation [30]. LIPIDS: Lipids are a major class of biomolecules. Most of them are nonpolar and mainly consist of carbon and hydrogen, with very little oxygen or nitrogen content. They are relatively insoluble in water and tend to be soluble in nonpolar organic solvents, such as ether. Lipids can be divided into sub classes that include fats, steroids, phospholipids, and waxes. Fats found in animals are generally solids at room temperature and those in plants are usually liquids at room temperature. These liquids are considered to be oils like corn oil, olive oil, etc. Fats and oils store energy. Fats also take in fat-soluble vitamins such as vitamins A (good for vision), D (helps the body absorb calcium for strong bones and teeth), E (an antioxidant for our body), and K (helps our blood clot). They insulate the body against heat loss and temporarily lock foreign substances in new fat tissue to protect organisms. In addition, they help regulate blood pressure and blood lipid levels and are important for inflammatory responses to injury. A fat and oil are often called triglyceride because they result from reactions of three fatty acids and an alcohol group from glycerol. See Fig. 2.11 [31]. It is an example of an unsaturated fat triglyceride (C55 H98 O6 ). Glycerol is on the left. On the right from top to bottom: palmitic acid, oleic acid, and alpha-linolenic acid. Steroids have many biological functions. A well-known steroid is cholesterol, which is responsible for maintaining the permeability of mammals’ cell membranes. Other steroids include the male and female sex hormones (testosterone and estrone respectively), aldosterone which helps regulate blood pressure and cortisone used to treat inflammatory conditions. Steroids have a ring structure with three six-member rings and a five member ring all fused together.

Fig. 2.11 This is an example of an unsaturated fat triglyceride (C55 H98 O6 )

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Fig. 2.12 This figure shows the self-organization of phospholipids: a spherical liposome, a micelle, and a lipid bilayer

Phospholipids contain a glycerol backbone attached to two fatty acids by ester links. Therefore, they are considered diglycerides. Phospholipids store energy and are important for the formation of cell membranes. See Fig. 2.12 [32]. Generally speaking the bacterial cell is surrounded by a cell membrane, which is primarily made of phospholipids. This membrane encloses the contents of the cell. It acts as a barrier to hold the nutrients, proteins, etc. of the cytoplasm within the cell. Phospholipids are able to form cell membranes because the two ends of the phospholipid have different properties (one end is hydrophilic and the other end is hydrophobic). The hydrophobic tails associate into a lipid bilayer to escape the aqueous environment. It is essential for a cell membrane to have a hydrophobic interior and a hydrophilic exterior because they restrict the passage of most molecules from one side of the membrane to the other. Waxes are secretions from plants and animals. They are solid at room temperature, but can melt at relatively low temperatures. Waxes are generally soft and hydrophobic. In nature, wax is typically used as a protective coating. For example, it is found on the leaves of certain trees to prevent the loss of water. They generally contain esters and alkanes. Pseudomonas aeruginosa produces an extracellular matrix that contains lipids like rhamnolipids. Rhamnolipids are considered to be bacterial surfactants that lower surface tension to allow bacteria to move across a surface [33–35]. This is important for swarming motility, which is a rapid (2–10 μm/s) and coordinated translocation of a bacterial population across solid or semi-solid surfaces [36]. Rhamnolipids provide Pseudomonas aeruginosa with virulence (an ability to infect or damage a host). They contribute to establishing and maintaining infection in cystic fibrosis patients by disrupting the cell membranes, etc. Rhamnolipids have a role in causing

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Fig. 2.13 A di-rhamnolipid structure is displayed. The rhamnolipid has a glycosyl head group (a rhamnose moiety) and a fatty acid tail such as 3-hydroxydecanoic acid. A di-rhamnolipid contains two rhamnose groups. Note Rhamnose is a naturally occurring deoxy sugar

the Pseudomonas aeruginosa’s cell surface to become hydrophobic. It should be mentioned that rhamnolipids are involved in main phases of biofilm development. They promote motility and prevent cells from tightly adhering to the substratum. Also it has been reported that rhamnolipids create and maintain fluid channels for water and oxygen flow around the base of the biofilm [37]. They are important for forming structure in biofilms too. A rhamnolipid structure is displayed in Fig. 2.13 [38].

2.5 Quorum Sensing In nature, microorganisms such as bacteria generally live in communities called biofilms, which are thin, slimy layers that encase bacteria and anchor them to a surface. Quorum sensing (cell to cell chemical communication) allows bacteria to coordinate their activity and to group together into communities that provide benefits. Therefore, biofilms are biological systems in which the bacteria are organized into a coordinated functional community. The bacteria in such a community share nutrients and are protected from harmful factors in the environment like antibiotics and disinfectants. Through the process of quorum sensing (QS), the gene expression in bacterial cells is altered when a certain threshold population (cell density) is obtained. Figure 2.14 displays cell density along with an autoinducer for Gram positive bacteria [39]. Bacteria that use quorum sensing produce and secrete a signaling chemical called an autoinducer. These signaling molecules are usually secreted at a low level by individual bacteria. However, at high cell density, the local concentration of signaling molecules may exceed the threshold level and cause changes in gene expression. Gram positive bacteria use autoinducing peptide (AIP) as their autoinducers, while Gram negative bacteria produce N-acyl homoserine lactones (AHL) as their signaling

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Fig. 2.14 This diagram displays cell density along with an autoinducer for Gram positive bacterial quorum sensing

molecule [40–43]. When the AHL molecules reach a certain threshold concentration, they are recognized by specific receptors that belong to a large class of DNA-binding transcription factors called “R-proteins,” such as LuxR in V. fischeri [44]. When the R-proteins bind to the specific AHL molecules, they directly regulate the transcription of the target genes. The general chemical structure of N-acyl homoserine lactone can be seen in Fig. 2.15 [45]. Autoinduction was first noticed in 1970 in the bioluminescent gram-negative bacterium Vibrio fischeri [46]. V. fischeri live in the light organ of the Hawaiian bobtail squid. Here they obtain rich nutrients that allow the bacteria to proliferate. When the bacteria density is high enough, the genes involved in bioluminescence are expressed and light is produced. In addition to bioluminescence, quorum sensing Fig. 2.15 This figure provides a general chemical structure of N-acyl homoserine lactone (AHL)

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plays important roles in regulating a variety of other cellular functions in bacteria such as virulence gene expression, biofilm formation, and antibiotic resistance.

2.6 Biofilm Collapse and Removal BIOFILM COLLAPSE: It should be mentioned that surface-attached communities of bacteria (known as biofilms) must release and disperse cells into the environment so that they can colonize new sites [47]. Biofilm dispersal is a promising area of research that may result in novel chemical agents to prevent and treat biofilms in a variety of settings (industrial, medical, etc.). A basic mechanism of biofilm dispersal is the production of extracellular enzymes that degrade adhesive components in the biofilm matrix. Since the biofilm matrix encases the bacterial cells within the biofilm colony, degradation of the matrix would allow the cells to detach and disperse into the environment. Examples of these degrading enzymes are glycosidases, proteases, and deoxyribonucleases [48]. Biofilms are communities of bacteria that attach to a surface and are difficult to eradicate when they are pathogenic or disease-causing. Researchers are studying various enzymes to collapse biofilms. Several discovered that the enzymes PslG and PelA are used by bacteria for wall building. These enzymes help create sugar molecules which serve as building blocks, but also prevent the buildup of chains of sugars inside the cells of bacteria. The researchers (involved with this project) thought that because the biofilm walls had a chained sugar component, the same enzymes might cause the wall to degrade. They tested this idea. First, the investigators applied Pseudomonas aeruginosa bacteria to culture dishes in their laboratory and waited for them to build biofilms. Then they added synthetic forms of the enzymes, which actually degraded the biofilm walls [49]. Other scientists used a novel approach to disrupt bacterial biofilms. They made a monoclonal antibody (protein) in their lab and directed it against DNABll proteins. The extracellular DNA (eDNA) and associated DNABll proteins are common components in biofilms that help maintain their structure. The monoclonal antibody directed against DNABll proteins resulted in the collapse of bacterial biofilms [50]. Another investigation was carried out to determine the effect of commercial proteases and amylases on biofilms formed by Pseudomonas fluorescens. Biofilm was grown in a diluted medium containing glass wool, which was used as an attachment surface. Extracellular polymeric substances (EPS) were extracted and the EPS composition was determined to be mostly proteins. Protease (savinase, everlase and polarzyme) and amylase (amyloglucosidase and bacterial amylase novo) activities were tested on the biofilms and the extracted EPS. Everlase and savinase were found to be the most effective enzymatic treatments for removing biofilms and degrading the EPS [51].

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Fig. 2.16 A pig is displayed in a cutout section of pipe

In addition to enzymes and chemicals such as biocides, biofilms collapse due to other reasons. Some include the limited amount of space and nutrients available within a biofilm, erosion, abrasion, environmental conditions, and cellular detachment induced through cell to cell communication. REMOVAL: Biofilms can be removed by various methods: chemical, natural, physical (by handling the object that needs to be cleaned) and mechanical. A couple of examples for mechanical removal are provided. Biofilms contribute to slime and scale buildup that result in clogged pipes. These clogged pipes are often cleaned by the pigging method, which uses a bullet-shaped cylinder called a pig to clean the pipelines [52, 53]. Pigs are blown through a pipe and require launching and retrieval. They fit tightly against the interior wall of the pipe and scrape the edges to remove biofilm and debris. To improve their cleaning effectiveness, brushes and blades can be added to the pigs. Figure 2.16 shows a pig in a cutout section of pipe [54]. Another example of mechanical removal of biofilm is the use of ultrasonic dental scalers [55]. A hygienist uses ultrasonic scalers to remove biofilm (bacterial plaque) from teeth. The scalers’ mechanical action of high vibrational energy actually loosens, crushes, and removes tartar and biofilm.

2.7 Biofilm and Infection Biofilms are an important factor in human health. Experts at the Centers for Disease Control and Prevention (CDC) estimate that 70% of human bacterial infections involve biofilms [56]. One approach to prevent biofilm formation is to incorporate

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antimicrobials into surfaces where biofilms might form. Chemical signals that allow quorum sensing are essential for biofilm formation. Therefore, researchers are working to determine the makeup of these chemical signals and hopefully block them. Many pathogens require iron to survive, so controlling the amount of available iron would be useful too. Biofilms are involved in a wide variety of infections. These include both device and non-device associated infections. For device related infections, biofilms usually occur on or within indwelling medical devices. Indwelling devices are made of biomaterials which need to be functional and have useful characteristics such as mechanical properties and biocompatibility. They also need to have high resistance to infection. A material’s resistance to infection is closely related to its resistance to biofilm formation. A biomaterial is a material intended to interface with biological systems in order to evaluate, treat, augment, or replace any tissue, organ, or function in the body [57]. Biomaterials should have an artificial immune system to suppress and control biofilm growth and development [58]. This is generally done in two main ways. One is a physical method in which special topography such as polymer brushes or patterned surfaces are included on the material. The other technique directly relates to a material’s chemical components. For example, bulk materials consisting of silver, copper, and their alloys show antibacterial effects. Also surface coatings (containing antibacterial effects) could be applied to neutral materials that have no effect on biofilm formation. DEVICE ASSOCIATED INFECTIONS: Some examples of these devices include contact lenses, catheters, and mechanical heart valves. Contact lenses can be considered soft or hard. However, microorganisms adhere to both types. The degree of adherence to the lenses depends on the water content, the nature of the substrate, electrolyte concentration, the composition of the polymer, and the type of bacterial strain involved. Several microorganisms that attach to contact lenses are E. coli, P. aeruginosa, and Staphylococcus aureus. Biofilm has been observed on the contact lenses of patients with keratitis (infection of the cornea) by using scanning electron microscopy [59]. This biofilm was produced by P. aeruginosa. Biofilm can form on contact lenses kept in storage cases too. Biofilms form on central venous catheters. However, the extent and location of the biofilm depends on the amount of time used for catheterization. A central venous catheter is a catheter placed into a large vein in the neck, chest, groin, or arms. It is often used to administer medication or fluids. See Fig. 2.17 [60]. Generally speaking, short-term catheters have biofilms form on their surfaces, while long-term catheters (about one month) have more internal biofim formation. The nature of the fluid passed through the central venous catheter affects the growth of the microbes. Gram-positive bacteria like S. epidermidis and S. aureus do not grow well in intravenous fluids, while Gram-negative bacteria such as P. aeruginosa do [61]. Urinary catheters may also contain biofilms. These catheters are usually made of silicon or latex and often used during surgery for urine excretion. These catheters pass through the uretha and go up to the urinary bladder. For an open catheter system, urine is drained into a toilet or other item for collection. However for a closed catheter system, urine accumulates in a plastic bag. Chances for infections like urinary tract

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Fig. 2.17 This diagram shows a non-tunneled central line inserted into the right subclavian vein

infections (UTI) are higher for open catheter systems. Microbes that form biofilms on these devices include E.coli, P. aeruginosa, and other gram negative bacteria [62]. Biofilms form on mechanical heart valves and cause a condition known as prosthetic valve endocarditis [63]. This disease may significantly impact cardiac functions. Endocarditis refers to the layer of the heart that is infected (endocardium). The endocardium is the inner layer of the heart and consists of endothelial tissue and lines the inside of the heart and valves. If tissue damage occurs during the time that the prosthetic heart valves are implanted, then the microbial cells have a better opportunity to colonize these areas. This condition is caused by bacteria like Streptococcus, S. aureus, etc. NON-DEVICE RELATED BIOFILM INFECTIONS: Infection can be defined as bacterial growth exceeding the resistant capability of hosts (human beings in this case) against entering bacteria. The entering bacteria attach to tissues of organs at the beginning stage, since they need to make biofilms for survival. Immediately, antigenantibody complex reactions are initiated by our immune systems. Humans have both an innate immune system and an adaptive immune system [64]. When pathogenic bacteria attach to human tissue for interactions, some fast automatic responses occur against the invading bacteria. Several examples include the physical topography of the tissue itself, the acidic environment of the stomach, antimicrobial agents in saliva, etc. Also some specific proteins might deteriorate the bacteria’s membranes. When these inherent systems can’t stop the bacterial growth and attachments, then phagocytic cells such as macrophages, neutrophils, and dendritic cells try to kill the entering bacteria. Also antiviral and inflammatory processes begin. Antiviral responses, which inhibit the development of the pathogen, lead to the shutdown of protein synthesis by signal deduction processes. On the other hand, the inflammatory

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process defends the body by bringing plasma proteins and phagocytes (white blood cells that engulf and consume foreign material) to the affected area. Human beings also have the adaptive immune response. It includes humoral immunity which is mediated by antibodies to neutralize microbes and cell-mediated immunity by T-lymphocytes that kill virally infected cells. See Fig. 2.18 [65]. The adaptive immune system could compensate the weak points of the original immune response. If the immune system works well, then biofilm could be suppressed and controlled. However, if it does not work well, then bacterial growth could continue and form biofilms through the quorum sensing process. Some non-device related biofilm infections include impetigo, chronic lung infections in cystic fibrosis (CF) patients, community-acquired pneumonia, meningitis, kidney stones, tuberculosis (TB), osteomyelitis, and periodontitis. Impetigo is a superficial infection of the skin that is caused by Staphylococci, Spreptococci, or multiple bacteria [66]. Bullous impetigo is a deep-seated infection of the skin caused by S. aureus. It displays large fluid-filled blisters (bullae) from original vesicles. When these large blisters burst, they leave raw red areas. Treatment involves cleaning the infected areas with soap and water followed by applications of antibiotics. Chronic lung infections in cystic fibrosis (CF) patients are caused by P. aeruginosa [67]. The infection remains present despite aggressive antibody therapy. It is a common cause of death in CF patients because of constant inflammatory damage to the lungs. One treatment during early biofilm development is to use the enzyme DNase to structurally weaken the biofilm. Community-acquired pneumonia (CAP) is mainly caused by S. pneumoniae, a gram positive organism that naturally resides in the upper respiratory tract and can cause pneumonia and other infections [68]. CAP usually occurs in a community setting or within the first 48 h after hospitalization. Meningitis is inflammation of the meanings, protective coverings of the brain and spinal cord [69]. Septic meningitis is caused by bacteria, especially Streptococcus pneumonaie and Neisseria meningitides. Symptoms of severe sepsis include the presence of bacteria in the blood, multiple organ dysfunctions, etc. Outbreaks of N. meningitidis infection are most likely to occur in dense groups such as college campuses and military bases. Factors that increase the risk of getting bacterial meningitis include the use of tobacco, the presence of viral upper respiratory infection, a weak immune system, etc. The kidneys are paired organs that regulate internal water content and balance solute concentrations. They also remove the body’s metabolic wastes, which are normally eliminated in the urine. However, urine flow can be obstructed by kidney stones, which often cause inflammation and infection. Stones are formed when urinary concentrations of substances like calcium oxalate, calcium phosphate, and uric acid increase. Stone formation is not clearly understood, so there are a number of theories about their causes. For example, struvite stones form in alkaline, ammonia-rich urine caused by the presence of urease-splitting bacteria such as Proteus, Pseudomonas, and Staphylococcus [70]. Therefore, infectious kidney stones are formed as a result of the interaction between bacteria and mineral substances

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Fig. 2.18 This diagram shows the T lymphocyte activation pathway. Some T cells direct and regulate immune responses and others directly attack infected cells

2.7 Biofilm and Infection

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present in the urine. Actually a complex biofilm is formed that is composed of infective bacteria and their exoproducts and mineralized stone material. Infectious stones obstruct urine flow and cause severe inflammation and infection that can lead to kidney failure. Mycobacterium tuberculosis is the bacterium that causes tuberculosis (TB). Each year there are about 450,000 new cases of TB that are resistant to various drugs. Close to one-third of TB-related deaths are due to antimicrobial resistance. One TB target that scientists have been studying is the enzyme phosphopantetheinyl transferase, which plays a role in the biosynthesis of mycolic acids that make up the bacterium’s cell wall and virulence lipids that the bacterium needs to suppress its hosts’ immune system [71]. Researchers found a compound (known as 8918) that fights TB by inhibiting the previously described enzyme [72]. The 8918 compound prevents the bacterium from reproducing itself in the lungs of mice infected with TB. (See Science 2019, DOI: https://doi.org/10.1126/science.aau8959) Progress has been made in the fight against TB. However, more work needs to be done because 8918 quickly metabolizes. Osteomyelitis is an infection of the bones that results in inflammation, pain, etc. [73]. It may be caused by bacterial cells. Bacteria enter the bones through the bloodstream, or from previous infections, or as a result of traumatic experiences. More than half of bone infections are caused by Staphylococcus aureus. Other pathogens include the Gram-positive organisms Streptococci and Enterocci, as well as Gram-negative bacteria like Pseudomonas [74]. Osteomyelitis may spread to adjacent soft tissues and joints. If the infectious process is not properly treated, then a bone abscess forms. This abscess cavity contains dead bone tissue that does not easily liquefy and drain. Therefore, it can’t properly heal. New bone growth forms and surrounds that area, which appears to be healed. However, recurring abscesses are produced throughout the patient’s life. This is considered to be chronic osteomyelitis. Periodontitis is an infection of the gums that damages soft tissues and the bones supporting the teeth [75]. Biofilms are present in the human body on the teeth as dental plaque, where they cause gum disease and tooth decay. Dental plaque is a biofilm that adheres to the teeth and contains many species of bacteria (such as Streptococcus mutans) embedded in salivary polymers and microbial extracellular products. Periodontitis is normally the result of poor oral hygiene, but may be caused by agents such as P. aerobicus and Fusobacterium nucleatum. These agents have the ability to form biofilms on a variety of surfaces including those in the oral cavity.

References 1. Slonczewski, J. L., & Foster, J. W. (2013). Microbiology: An evolving science (Third Ed.) (p. 82). New York: W W Norton. ISBN 9780393123678. 2. Zifran, A. File: Prokaryote cell.svg. Date: October 12, 2015. License: Creative Commons Attribution-Share Alike, 4.0 International. https://commons.wikimedia.org/wiki/File: Prokaryote_cell.svg.

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3. Yuval. File: Gram stain Anthrax.jpg. Date: November 25, 2005. This work is in the public domain. (It is a work of the Centers for Disease Control and Prevention, part of the United States Department of Health and Human Services, for the U.S. Federal government.) https:// commons.wikimedia.org/wiki/File:Gram_Stain_Anthrax.jpg. 4. Lappin-Scott, H. M., Jass., J. & Costerton, J. W. (1993). Microbial biofilm formation and characterization. In Society for Applied Bacteriology technical series, Society for Applied Bacteriology Symposium (p. 30). 5. Costerton, J. W. (1999). Introduction to biofilm. International Journal of Antimicrobial Agents, 11, 217–221; discussion, pp. 237-239. 6. Lappin-Scott, H. M. & Costerton, J. W. (1995). Microbial biofilms. Cambridge, New York: Cambridge University Press. 7. Dolan, R., & Carr, J. File: Staphylococcus aureus biofilm 01.jpg. Date: April 19, 2006. This work is in the public domain. (It is a work of the Centers for Disease Control and Prevention, part of the United States Department of Health and Human Services, for the U.S. Federal government.) https://commons.wikimedia.org/wiki/File:Staphylococcus_aureus_biofilm_ 01.jpg. 8. Chandki, R., Banthia, P., & Banthia, R. (2011). Biofilms: A microbial home. Journal of Indian Society of Periodontology, Apr–June, 15(2), 111–114. https://doi.org/10.4103/0972124x.84377. 9. O’Toole, G., Kaplan, H. B., & Kolter, R. (2000). Biofilm formation as microbial development. Annual Review of Microbiology, 54, 49–79. https://doi.org/10.1146/annurev.micro.54.1. 49.ISSN0066-4227.PMID11018124. 10. Davis, D. File: Biofilm.jpg. Date: November 13, 2007. License: Creative Commons Attribution 2.5 Generic. https://commons.wikimedia.org/wiki/File:Biofilm.jpg. 11. Garrett, T. R., Bhakoo, M., & Zhang, Z. (2008). Bacterial adhesion and biofilms on surfaces. Progress in Natural Science, 18(9), 1049–1056. https://doi.org/10.1016/j.pnsc.2008.04.001. 12. Sketch by Hideyuki Kanematsu. 13. Flemming, H.-C., Neu, T. R., & Wozniak, D. J. (2007). The EPS matrix: The house of biofilm cells. Journal of Bacteriology, 189(22), 7945–7947. https://doi.org/10.1128/JB.00858-07. 14. Allonweiner. File: Bacillus subtilis.jpg. Date: January 18, 2007. This work is in the public domain. (It was taken by a Tecnai T-12 TEM.). https://commons.wikimedia.org/wiki/File: Bacillus_subtilis.jpg. 15. Mitchell, K., Zarnowski, R., & Andes, D. (2016). The extracellular matrix of fungal biofilms. In I. Christine (Ed.), Fungal biofilms and related infections (Vol. 3, pp. 21–24). Springer. 16. Roux, D., Cywes-Bentley, C., Zhang, Y. F., Pons, S., Konkol, M., Kearns, D. B., et al. (2015). Identification of Poly-N-acetylglucosamine as a major polysaccharide component of the Bacillus subtilis biofilm matrix. Journal of Biological Chemistry, 290(31), 19261–19272. https:// doi.org/10.1074/jbc.M115.648709. 17. Mann, E. E., Wozniak, D. J. (2012). Pseudomonas biofilm matrix composition and niche biology. FEMS Microbiology Review, 36(4), 893–916. https://doi.org/10.1111/j.1574-6976. 2011.00322.x 18. Colvin, K. M., Gordon, V. D., Murakami, K., Borlee, B. R., Wozniak, D. J., Wong, G. C., et al. (2011). The pel polysaccharide can serve a structural and protective role in the biofilm matrix of Pseudomonas aeruginosa. PLoS Pathogens, 7(1), e1001264. https://doi.org/10.1371/journal. ppat.1001264. 19. Mulcahy, H., Charron-Mazenod, L., & Lewenza, S. (2008). Extracellular DNA chelates cations and induces anti-biotic resistance in Pseudomonas aeruginosa biofilms. PLoS Pathogens, 4(11), e1000213. https://doi.org/10.1371/journal.ppat.1000213. 20. Yang, L., Hu, Y., Liu, Y., Zhang, J., Ulstrup, J., & Molin, S. (2011). Distinct roles of extracellular polymeric substances in Pseudomonas aeruginosa biofilm development. Environmental Microbiology, 13(7), 1705–1717. https://doi.org/10.1111/j.1462-2920.2011.02503.x. 21. Milo, R. (2013). What is the total number of protein molecules per cell volume? A call to rethink some published values. BioEssays, 35(12), 1050–1055. https://doi.org/10.1002/bies. 201300066.PMC3910158.PMID24114984.

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22. Romero, D., Vlamakis, H., Losick, R., & Kolter, R. (2014). Functional analysis of the accessory protein TapA in Bacillus subtilis amyloid fiber assembly. Journal of Bacteriology, 196(8), 1505–1513. https://doi.org/10.1128/JB.01363-13. 23. Toth, A. File: Myoglobin.png. Date: 2008. This work is in the public domain. https://commons. wikimedia.org/wiki/File:Myoglobin.png. 24. Chem. Grad. Student. File: Peptide-Figure-Revised.png. Date: September 5, 2011. License: Creative Commons Attribution—Share Alike 3.0 https://commons.wikimedia.org/wiki/File: Peptide-Figure-Revised.png. 25. Benjah-bmm27. File: Cellulose-Ibeta-from-xtal-2002-3D-balls.png. Date: April 24, 2009. This work is in the public domain. https://commons.wikimedia.org/wiki/File:Cellulose-Ibeta-fromxtal-2002-3D-balls.png. 26. Roux, D., Cywes-Bentley, C., Zhang, Y.-F., Pons, S., Konkol, M., Kearms, D., et al. (2015). Identification of Poly-N-acetyl Glucosamine as a major polysaccharide component of the Bacillus subtilis biofilm matrix. Journal of Biological Chemistry. https://doi.org/10.1074/jbc.m115. 648709. 27. Yikrazuul. File: N-Acetylglucosamine.svg. Date: November 20, 2008. This work is in the public domain. https://commons.wikimedia.org/wiki/File:N-Acetylglucosamine.svg. 28. Ma, L., Lu, H., Sprinkle, A., Parsek, M., & Wozniak, D. (2007). Pseudomonas aeruginosa Psl is a galactose- and mannose-rich exopolysaccharide. Journal of Bacteriology, 189(22), 8353–8356. https://doi.org/10.1128/JB.00620-07. 29. Dahm, R. (2008). Discovering DNA: Friedrich Miescher and the early years of nucleic acid research. Human Genetics, 122(6), 565–581. https://doi.org/10.1007/s00439-007-04330.PMID17901982. 30. Jurcisek, J., Brockman, K., Novotny, L., Goodman, S., & Bakaletz, L. (2017). Non typeable Haemophilus influenzae releases DNA and DNABll proteins via T4SS-like complex and ComE of the type IV pilus machinery. PNAS, 114(32), E6632–E6641. https://doi.org/10.1073/pnas. 1705508114. 31. Schaefer, W. File: Fat triglyceride shorthand formula.png. Date: April 21, 2005. This work is in the public domain. https://commons.wikimedia.org/wiki/File:Fat_triglyceride_shorthand_ formula.PNG. 32. Villarreal, M. R. File: Phospholipids aqueous solution structures. svg. Date: November 6, 2007. This work is in the public domain. https://commons.wikimedia.org/wiki/File:Phospholipids_ aqueous_solution_structures.svg. 33. Desai, J. D., & Banat, I. M. (1997). Microbial production of surfactants and their commercial potential. Microbiology and Molecular Biology Reviews, 61(1), 47–64. 34. Lang, S., & Wullbrandt, D. (1999). Rhamnose lipids–biosynthesis, microbial production and application potential. Applied Microbiology and Biotechnology, 51(1), 22–32. https://doi.org/ 10.1007/s002530051358.PMID10077819. 35. Soberón-Chávez, G., Aguirre-Ramírez, M., & Sánchez, R. (2005). The Pseudomonas aeruginosa RhlA enzyme is involved in rhamnolipid and polyhydroxyalkanoate production. Journal of Industrial Microbiology and Biotechnology, 32(11–12), 675–677. https://doi.org/10.1007/ s10295-005-0243-0.PMID15937697. 36. Glick, R., Gilmour, C., Tremblay, J., Satanower, S., Avidan, O., Dézie, E., et al. (2010). Increase in rhamnolipid synthesis under iron-limiting conditions influences surface motility and biofilm formation in Pseudomonas aeruginosa. Journal of Bacteriology, 192(12), 2973–80. https:// doi.org/10.1128/JB.01601-09.PMC2901684.PMID20154129. 37. Davey, M. E., Caiazza, N. C., & O’Toole, G. A. (2003). Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PAO1. Journal of Bacteriology, 185(3), 1027–36. https://doi.org/10.1128/jb.185.3.1027-1036.2003.PMC142794.PMID12533479. 38. Boghog. File: Rhamnolipid.tif. Date: November 20, 2011. This work is in the public domain. https://commons.wikimedia.org/wiki/File:Rhamnolipid.tif. 39. Gira, J. File: Gram Positive Bacteria Quorum Sensing.pdf. Date: December 19, 2016. License: Creative Commons Attribution-Share Alike 4.0 International https://en.wikipedia.org/wiki/ File:Gram_Positive_Bacteria_Quorum_Sensing.pdf.

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40. Miller, M. B., & Bassler, B. L. (2001). Quorum sensing in bacteria. Annual Review of Microbiology, 55, 165–199. 41. Solano, C., Echeverz, M., & Lasa, I. (2014). Biofilm dispersion and quorum sensing. Current Opinion in Microbiology, 18, 96–104. 42. Ikegai, H. (2015). Genomics approach. In: Kanematsu, H., Barry, D. M. (Eds.), Biofilm and Materials Science. New York, The USA: Springer. 43. Whiteley, M., Diggle, S. P., & Greenberg, E. P. (2017). Progress in and promise of bacterial quorum sensing research. Nature, 551, 313–320. 44. Li, Zhi, & Nair, Satish. (2012). Quorum sensing: How bacteria can coordinate activity and synchronize their response to external signals. Protein Science, 21(10), 1403–1417. https:// doi.org/10.1002/pro.2132. 45. Hbf878. File: N-Acyl Homoserine Lactone.svg. Date: December 24, 2017. License: Creative Commons CCO 1.0 Universal Public Domain https://commons.wikimedia.org/wiki/File:NAcyl_Homoserine_Lactone.svg. 46. Nealson, K. H., Platt, T., & Hastings, J. W. (1970). Cellular control of the synthesis and activity of the bacterial luminescent system. Journal of Bacteriology, 104, 313–322. 47. Chapman, J. (2015). Detachment of bacteria. In H. Kanematsu & D. M. Barry (Eds.), Biofilm and Materials Science. New York, The USA: Springer. 48. Kaplan, J. B. (2010). Biofilm dispersal: Mechanisms, clinical implications, and potential therapeutic uses. Journal of Dental Research, 89(3), 205–218. https://doi.org/10.1177/ 0022034509359403. 49. Yirka, B. (2016). Enzymes found that can tear down bacterial biofilm walls Phys.org (report). https://phys.org/news/2016-05-enzymes-bacterial-biofilm-walls.html. 50. Novotny, Laura, et al. (2016). Monoclonal antibodies against DNA-binding tips of DNABll proteins disrupt biofilms in vitro and induce bacterial clearance in vivo. EBioMedicine. https:// doi.org/10.1016/j.ebiom.2016.06.022. 51. Molobela, P., Cloete, T. E., & Beukes, M. (2010). Protease and amylase enzymes for biofilm removal and degradation of extracellular polymeric substances (EPS) produced by Pseudomonas fluorescens bacteria. African Journal of Microbiology Research, 4(14), 1515–1524. 52. Sadaf, M. (2013). Calculation of pigging effectiveness for petroleum (product) pipelines. International Journal of Scientific and Research Publications, 3(9), 2123. ISSN 2250-3153. 53. Cloyde, C. (2011). Pig trap design and assessment consideration. Pipeline & Gas Journal, 36–42. 54. Barrison, H. File: Pipeline PIG.jpg. Date: February 24, 2009. License: Creative Commons Attribution-Share Alike 2.0 Generic. https://commons.wikimedia.org/wiki/File:PipelinePIG. jpg. 55. Ryan, D. L., Darby, M., Bauman, D., Tolle, S., & Naik, D. (2005). Effect of ultrasonic scaling and hand-activated scaling on tactile sensitivity in dental hygiene students. Journal of Dental Hygiene, 79(1), 1–13. 56. Tortora, G., Funke, B., & Case, C. (2016). Microbiology: An introduction (12th ed.) (pp. 156– 157). U.S. Pearson. 57. Black, J. (Ed.). (2005). Biological performance of materials—Fundamental of biocompatibility. Boca Raton, FL: CRC Press-Taylor & Francis. 58. Kanematsu, H., Barry, D. M., Ikegai, H., Yoshitake, M., & Mizunoe, Y. (2017). Biofilm evaluation methods outside body to inside—Problem presentations for the future. Medical Research Archives, 5, 1–17. 59. Szczotka-Flynn, L., Imamura, Y., Chandra, J., Yu, C., Muherjee, P., Pearlman, E., et al. (2009). Increased resistance of contact lens related biofilms to antimicrobial activity of soft contact lens care solutions. Cornea, 28(8), 918–926. https://doi.org/10.1097/ICO.0b013e3181a81835. 60. Blaus, B. File: Blausen 0181. Date: November 6, 2013. License: Creative Commons Attribution 3.0 https://commons.wikimedia.org/wiki/File:Blausen_0181_Catheter_ CentralVenousAccessDevice _NonTunneled.png.

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61. Garcia-Caballero, J., Heruzo-Cabrera, H., Vera-Cortes, M. L., Garcia de Lorenzo, A., VazquezEncinar, A., Garcia-Caballero, F., del Rey-Calero, J. (1985). The growth of micro-organisms in intravenous fluids. Journal of Hospital Infection, 6(2), 154–157. https://doi.org/10.1016/ S0195-6701(85)80092-X. 62. Nicolle, Lindsay. (2014). Catheter associated urinary tract infections. Antimicrobial Resistance and Infection Control, 3, 23. https://doi.org/10.1186/2047-2994-3-23. 63. Ivanovic, B., Trifunovic, D., Matic, S., Petrovic, J., Sacic, D., & Tadic, M. (2019). Prosthetic valve endocarditis—A trouble or a challenge? Journal of Cardiology, 73(2), 126–133. 64. Nicholson, L. (2016). The immune system. Essays in Biochemistry, 60(3), 275–301. 65. Hazmat2. File: T cell activation.svg. Date: January 28, 2012. License: This work is in the public domain. https://commons.wikimedia.org/wiki/File:T_cell_activation.svg. 66. Pereira, L. B. (2014). Impetigo-review. Anais Brasileiros de Dermatologia, 89(2), 293–299. 67. Doring, G., Flume, P., Heijerman, H., & Elborn, S. (2012). Treatment of lung infection in patients with cystic fibrosis: Current and future strategies. Journal of Cystic Fibrosis, 11(6), 461–479. 68. Wunderink, R., & Waterer, G. (2014). Community-acquired pneumonia. The New England Journal of Medicine. https://doi.org/10.1056/NEJMcp1214869. 69. Bamberger, D. (2010). Diagnosis, initial management and prevention of meningitis. American Family Physician, 15; 82(12), 1491–1498. 70. Borghi, L., Nouvernne, A., & Meschi, T. (2012). Nephrolithiasis and urinary tract infections: ‘The chicken or the egg’ dilemma? Nephrology, Dialysis, Transplantation, 27(11), 3982–3984. 71. Ballinger, E., Mosior, J., Hartman, T., et al. (2019). Opposing reactions in coenzyme A metabolism sensitive Mycobacterium tuberculosis to enzyme inhibition. Science, 363(6426), eaau8959. https://doi.org/10.1126/science.aau8959. 72. Rawal, T., & Butani, S. (2016). Combating tuberculosis infection: A forbidding challenge. Indian Journal of Pharmaceutical Sciences, 78(1), 8–16. 73. Calhoun, J., Manring, M. M., & Shirtliff, M. (2009). Osteomyelitis of the long bones. Seminars in Plastic Surgery, 23(2), 59–72. https://doi.org/10.1055/S-0029-1214158. 74. Hatzenbuehler, J., & Pulling, T. J. (2011). Diagnosis and management of osteomyelitis. American Family Physician, 84(9), 1027–1033. 75. Hajishengallis, G. (2015). Periodontitis: From microbial immune subversion to systemic inflammation. Nature Reviews Immunology, 15(1), 30–44.

Chapter 3

Animate Substrata and Biofilms

Abstract Biofilms form on natural animate substrata. For substrata discussed in this chapter, we selected plants’ surfaces and the tissues inside of human bodies. Biofilms are a result of bacterial activity. First bacteria attach to a surface (at an interface), aggregate, and then increase their number to a certain threshold value. At this point they excrete extracellular polymeric substances to form biofilms. The phenomenon is brought about by quorum sensing, a sort of signal transmission process. This process is similar to that of biofilms in other environments. With an understanding of the proposed mechanisms and phenomenon, one has a chance to utilize the benefits of biofilms and to control their negative effects. In this chapter, biofilms are described for an animate natural environment.

3.1 Introduction Biofilms generally form at heterogeneous interfaces. Therefore, they also form on biological tissues. In nature there are many kinds of interfaces and substrates. This chapter describes two cases. One of them relates to plants and biofilms and the other one relates to humans and biofilms. These are representative cases for animate organisms. Plants are very important for our global environmental system. They absorb inorganic matter and light to produce chemicals, as shown in Fig. 3.1. Approximately 40% of the total carbon absorbed by plants is transformed to carbon compounds excreted from the root areas of plants [1, 2]. As shown in Fig. 3.1, many chemicals are produced in the root area (called “Rhizosphere”, as described below). These chemicals attract bacteria, so it is no wonder that a certain bacterial flora form in this area. It was already mentioned that the aggregation of bacteria form biofilms through the process of quorum sensing. In such a way, biofilms form in the root area (rhizosphere). However, biofilm formation should not be restricted to root areas. They can form on other parts of plants too.

© Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_3

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Fig. 3.1 Carbon circulation and the role of plants

The space around plants is classified as the phyllosphere and the rhizosphere. These topics are described in Sect. 4.1. The rhizosphere corresponds to the underground space, and the phyllosphere is the space above ground, as shown in Fig. 3.2. The underground space for plants includes roots and rhizomes. (A rhizome is a rootstock that contains nodes from which roots and shoots originate.) Therefore, a plant’s space underground is classified into two groups. On the other hand, the space for plants above ground is called phyllosphere. It is classified into four categories. Figure 3.3 shows the classification schematically. Caulosphere is part of the caulome and refers to stems. Phylloplane is the surface of leaves. Anthosphere is the flower part. Carposphere is the fruit part. For all of these spaces, bacteria exist and inevitably form biofilms. Fig. 3.2 The classification of space around plants

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Fig. 3.3 This is a detailed classification of the phyllosphere. Caulosphere refers to stems

The rhizosphere is defined as the space affected by plants’ roots [3, 4]. Usually the rhizosphere corresponds to the area that is about several meters from the root. It is classified into three regions as shown in Fig. 3.4. Endorhizosphere is the inner area under the epidermis. The rhizoplane corresponds to the surface of the roots. The exorhizosphere refers to the soil parts around the plants’ roots. On the other hand, the non-rhizosphere area is called the “bulk soil”. As previously mentioned, there is an abundance of nutrition (chemicals) in the rhizosphere. Therefore, it contains lots of bacteria and biofilms. Biofilms in the rhizosphere have some special roles and affect the rhizosphere and the soil. On the other hand, biofilms also form on human tissues. They can form inside of our bodies. In this environment, bacteria and their activities are controlled by the immune systems. Therefore, the competitive formation of biofilms against the immune systems could lead to chronic diseases and hospital acquired infections. It is sometimes difficult to distinguish between non-biofilm infections and biofilm caused infections. However, there are already many findings that make correlations between diseases and biofilms. It has been well-known (since Antonie van Leeuwenhoek’s discovery) that biofilms form easily on teeth. Therefore, diseases related to our teeth might be caused by biofilms. Caries [5–7] are obviously caused by biofilms and their development. Fig. 3.4 This figure displays the three regions of the rhizosphere

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Dental plaque has been confirmed by the use of light microscopy, electron microscopy and/or both [8–11]. It was also confirmed by using Fluorescent In situ Hybridization Probes (FISH) [10]. Periodontitis is also caused by biofilms [12–21]. The investigation (about it) started at an early stage of biofilm research. Theilade observed the presence of biofilms by using electron microscopy [22]. Zijnge et al. investigated the biofilm problem by using FISH [23]. The biofilms on teeth are very often caused by Streptococcus mutans at the early stage [22–28]. In the western world, cystic fibrosis (a lung disease) has appeared often, since quite a few people have had the genetic factor for the disease [29–37]. The disease is caused by Pseudomonas aeruginosa which form biofilms easily. Therefore, the disease has been investigated from the viewpoint of biofilms. Chronic wounds are also related to biofilms [38–43]. In addition, it was found in the early part of this twentyfirst century that Otitis media is closely related to biofilms [44–48]. The relation of chronic diseases to biofilms should be attributed to the resistance of bacteria to antibiotics and to our natural immune systems.

3.2 Introduction Biofilms form at interfaces as a result of bacterial activities. They also form on plants’ tissues in nature. As described later (Chap. 4), the interfaces where biofilms form are classified into two categories: above ground and underground. The former space is generally called phyllosphere. In this category, biofilms are present on leaves and stalks of plants. The second category is the rhizosphere. The role of biofilms is very important for plants and for soils. The nutrition needed for plants to survive is mostly absorbed in the rhizosphere. Here biofilms play an important role between the soil and the plants.

3.2.1 Biofilm on Leaves The surface of leaves is described in Fig. 3.5 schematically. The outermost surface of leaves is called cuticula, a transparent hydrophobic layer. Cuticula is composed of cutin, a polymer of unsaturated fatty acid, and non-aqueous fatty ester (wax). Since cuticula is a hydrophobic layer, water does not enter the layer. Due to the existence of the cuticula layer, the surface of a leaf repels water. Also it is hard for water to evaporate from the inside of leaves. On such a surface, bacteria must exist and survive in the phyllosphere. Therefore, such an environment must be very tough and severe for bacteria. Usually, the number of bacteria on leaves is 106 –107 /cm2 . This number differs from plant to plant. The space on a leaf is generally a severe oligotrophic (low in nutrients) environment for

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Fig. 3.5 The surface structure of leaves

bacteria with a scant amount of nutrition and water. Therefore, the bacteria form biofilms in order to survive on leaves. When we come to think about the effect on bacteria (that are on leaves) we can classify the flora into two main groups. One of them is called “epiphyte” and they live on the surface of leaves, etc. In the past, all of the bacteria in the phyllosphere would have been epiphytes. We now know that some bacteria live inside of the leaves or plant tissues. They are called endophytes. However, most bacteria in the phyllosphere live on the surface of leaves (epiphyte). The number of epiphyte bacteria could be estimated as several million to tens of millions per 1 cm2 . However the correct numbers depend on the kinds of plants. From numerous test results, the total number of bacteria in the phyllosphere could be estimated as 1026 on the globe. The biocide is pretty large and it could affect the carbon and nitrogen circulation cycles. The factors affecting bacterial biota on leaves are summarized in Fig. 3.6. The effect of host plants would generally lead to differences of surface (topographical) properties and chemical properties. Table 3.1 shows the microbes observed well in the phyllosphere. The weather obviously affects the survival condition for bacteria. The tendency depends on whether they are epiphytes or endophytes. The total number of bacteria Fig. 3.6 General factors that affect biofilms on leaves

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Table 3.1 Some typical examples of Microbes observed in the phylloshere Bacteria

Habitats

References

Alternaria sp. Microsphaeropsis sp. Cladosporium sp. Pestalotiopsis sp.

Epiphyte of ever green broad leaves’ trees

[49–57]

Phomopsis sp. Phyllosticta sp. Colletotrichum spp.

General endophyte

[58–61]

Epicoccum sp., Botrytis sp., Phoma sp., Mucor sp., Trichoderma sp.

General epiphyte

[62–69]

in the phyllosphere does not generally change as a whole. The flora components on the sites seem to be affected by the weather [70, 71]. Lambais et al. suggest that a few percent of bacteria would be common for all leaves. Most bacteria differ from plant to plant, since the chemical matter and structures differ from plant to plant. Therefore, the flora in the phyllosphere would be affected [72]. It should be mentioned that the host plants obviously affect the flora in the phyllosphere, since chemical reactions must exist between bacteria and the host plants. In general, ultraviolet light impairs the DNA of organisms. Also biocides on leaves of plants are very tough for bacteria. Therefore, bacteria in the phyllosphere (particularly the epiphytes) have a high tolerance for ultraviolet light as compared to the endophytes. Some bacteria are able to recover the DNA that was impaired by UV. Actually Psuedomonas syringae have demonstrated the recovery function of DNA after impairment by UV [73, 74]. Biocides affect bacterial flora and can change them drastically. When agrichemicals are used, they generally decrease the number of bacteria drastically. However, the effect differs from bacteria to bacteria, since the action mechanism depends on the combination of bacteria and biocides. Usually bacteria tend to aggregate and form biofilms. This process also takes place in the phyllosphere. Since biofilms protect bacteria, they provide a survival strategy for them. Biofilms must have higher resistance and tolerance to harmful factors than bacteria [75]. This means that bacteria in biofilms have higher resistance to ultraviolet light, drying, biocides, etc. [76]. Particularly, they have high resistance to drying. It is very natural when we come to think about biofilms, since biofilms contain lots of water. Actually, a researcher pointed out that the quorum sensing capability indirectly increased the resistance to drying [77]. Some researchers pointed out that the horizontal gene transfer would occur very often in biofilms [78]. Therefore, the structural changes must occur with high probability in biofilms. We presume that the characteristics would lead to the high tolerance and resistance to biocides, UV tolerance, etc. The gene characteristics also produce bacteria that can survive the drastic change of conditions in the phyllosphere. Biofilms in the rhizosphere are very important to bacteria in the region and to the soil environments. As shown in Fig. 3.1 in the previous section, plants produce many chemicals particularly in the rhizosphere. Most of the chemicals produced by plants attract bacteria through chemotaxis (a movement toward nutrients). Therefore, the number of bacteria at the surface of roots increases to produce biofilms. They get

3.2 Introduction

45

lots of matter from the roots (exudates, substances secreted by the plant) and also from soil in the rhizosphere. Therefore, biofilms must have an intermediate function in order to exchange chemical matter, as shown in Fig. 3.7. Therefore, by controlling the biofilm’s function, one could change the properties of the soil. Generally, biofilms in the rhizosphere have some characteristic functions, as shown in Fig. 3.8. Some bacteria in the phyllosphere are well-known to work for each function. However, most of them belong to bacteria living in the rhizosphere. Biofilms that form in the rhizosphere could mainly increase chemicals as compounds and water. Therefore, the number of bacteria would increase as a result. This phenomenon (the increase of bacteria and microbe numbers) is called the rhizosphere

Fig. 3.7 Roots, soils, and biofilms: a schematic of their mutual relationships

Fig. 3.8 Biofilms in the rhizosphere have some characteristic functions

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3 Animate Substrata and Biofilms

effect [79–81]. As a result, biofilm formation would accelerate and microbiomes would increase. Such a rhizosphere effect would repeat cyclically. In such a case, bacterial flora might bring plants beneficial results and harmful effects. A beneficial effect would be an increase in the beneficial bacteria [82–85] to enrich the bulk soil as well as the rhizosphere, since they would increase the nutrition and symbiosis in the rhizosphere. On the other hand, there are some pathogenic bacteria in the rhizosphere such as Agrobacterium tumefaciens, Burkholderian cepacia, etc., [86, 87]. In some cases, beneficial bacteria in the rhizosphere could increase the host plants’ resistance to pathogenic bacteria [88–90]. This is carried out by excreting lipopolysaccharides, flagellin, salicylic acid and siderophores, cyclic lipopeptides, etc. [91–94]. The phenomenon is called Induced Systemic Resistance (ISR) [95, 96]. As described above, biofilms in the rhizosphere activate and accelerate bacterial activities and numbers, as well as plants’ activities. As a result, a soil’s environment could change. From the viewpoint of biofilm utilization, positive investigations and ideas are expected.

3.3 Biofilms on Animal (and Human) Tissues Bacteria and fungi on animal and human tissues have been studied to a great extent because they lead to diseases. This section focuses on such a topic. As Fig. 3.9 shows, biofilms not only form on rocks and artificial solid materials, but also on human tissues. Outside of the human body, bacteria grow and produce biofilms that result in sticky and slimy surfaces. Biofilms cause many industrial problems like corrosion, scales, hygiene deterioration, etc. On the other hand, problems might occur inside of our bodies. The human body has some natural countermeasures to control the growth of bacteria. However, bacteria can sometimes overcome a person’s natural

Fig. 3.9 Schematic illustration of biofilms in the extracorporeal and intracorporeal areas

3.3 Biofilms on Animal (and Human) Tissues

47

resistance and begin to grow. This causes an infection. Since bacteria in biofilms have high tolerance to antibiotics, biofilms could lead to chronic diseases and hospital acquired infection. The close relationship between diseases and biofilms has already been mentioned. Therefore, some medical researchers may insist that biofilms are equal to infection itself. The case outside of our bodies is described in other chapters. This chapter discusses the case inside of our bodies.

3.3.1 What Happens Inside of Our Bodies, When Pathogenic Bacteria Enter Them? When pathogenic bacteria enter our bodies, we have three automatic responses to remove exogenous matters, as shown schematically in Fig. 3.10 [97]. The first response is the innate immunity reaction. When bacteria try to enter our bodies, the surface area of skin (composed of keratin) provides a strong physical barrier to the entering bacteria. Mucous membranes also secrete many kinds of enzymes (for example: lysozyme, a component of salvia, sweat, tears, nasal and vaginal secretions) to kill bacteria. Also hydrochloric acid (which is secreted from the stomach) can kill bacteria. There is another innate reaction for us too. It includes defensive cells called phagocytes and macrophages. Figure 3.11 shows a schematic illustration of how phagocytes swallow bacteria and kill them. All of these protective reactions are innately provided to humans. These reactions start very early when the body encounters bacteria. In addition to macrophages, we can mention dendritic cells, neutrophills, natural killer cells (NK cells), etc. which are included in the innate immune systems.

Fig. 3.10 Various immunity reactions inside of the human body

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Fig. 3.11 Schematic illustration of phagocytosis (ingestion of bacteria, etc. by phagocytes)

On the other hand, human beings have an adaptive immune system. This system uses antibodies to protect us from toxins. (The presence of antigens induces the immune system to produce antibodies.) This system begins to work, when the innate system is not able to kill the bacteria and microbes entering our human bodies. Generally, two cells are mentioned as adaptive immune cells: B cells and T cells. B cells are also called B lymphocytes. They are a type of lymphocyte produced in bone marrow. On the other hand, T cells are produced in the thymus, and are a sort of lymphocyte. The acquisition of immunity can be carried out through a signal deduction process in the following way. As for the innate immune response, macrophages or dendritic cells quickly reach the infection site where bacteria or microbes enter our body. They try to combat the microbes and collect information about them. Then they transmit the information to T cells (helper T cells). The helper T cells transmit the signals (proteins such as cytokines and hormones such as interleukin-2) to killer T cells in order to increase the growth of killer T cells. Killer T cells could destroy bacterial cells and viruses. Helper T-cells, that get information from macrophages, also transmit the information to B cells. B cells get the information from the B cell receptors and finally produce antibodies. The relationship among the immune cells is displayed in Fig. 3.12. If bacteria overcome the multi-step immune systems, then biofilms continue to grow and cause infections. What kind of interactions might occur between biofilms and the immune systems? Such questions might be answered after lots of further investigations take place. At this point, there are some research reports about the interaction between biofilm formation/growth and macrophages’ activities [98–106]. They suggest that biofilms could control phagocytosis. This means that biofilms could accelerate the spread

3.3 Biofilms on Animal (and Human) Tissues

49

Fig. 3.12 Diagram of our adaptive immune system

of infections. Other ongoing studies include the relationship between T1 cells and biofilms [107] and the signal transmission processes of cytokines [108–114]. The connection of biofilms to infections will be clarified on the molecular level.

3.3.2 The Characteristics of Bacteria in Biofilms Bacteria form and continue to grow, when infections occur. Bacteria in biofilms could have tolerance and resistance to antibiotics. This situation could lead to chronic diseases, due to some changes of the bacteria in biofilms. What are the changes? The answers have not been fully established yet. However, some hypotheses have been proposed and might be true to some extent. One of them is the efflux pump system [115–121]. The efflux pump is actually a pumping out system by multimolecular base membranes. The function was found in gram-negative bacteria. Here, the antibiotic drugs would be moved (pumped) out of the cells (outside of the bacteria in biofilms). The second example is a classic one. Many antibiotics have difficulty diffusing into biofilms because of the existence of EPSs (Extracellular polymeric substances) [122–126]. Therefore, the antibiotics would not be able to reach the targeted bacteria. We presume that these phenomena could make a great contribution to the explanation of the mechanism involved. However, some antibiotics could penetrate and diffuse into biofilms, even though they might not work as antibiotics. A third example is the physiological changes due to dormant conditions [127– 130]. Physiological changes of bacteria in biofilms might give them tolerance and resistance to antibiotics. The fourth one is phenotypic change of bacterial cells in biofilms [131–135]. If the phenotype changes for some bacteria, then the antibiotics could miss the original target of bacteria.

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The fifth one is persister cells, a sub-population of non-growing bacterial cells. They have a high resistance to antibiotics [136–140]. Some bacteria neither grow nor die in the presence of bactericidal agents. These bacteria (existing in biofilms) would be dormant and have a slow rate of bacterial growth. In such a situation, persister cells are produced and exhibit multidrug tolerance. All of the above mechanisms suggest that bacteria in biofilms are very different from the planktonic state. When we come to think about biofilms, they serve as a survival strategy for bacteria and protect them. As a result, bacteria have high resistance and tolerance to antibiotics inside our bodies as well as to biocides outside of our bodies.

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76. Beattie, G. A., & Lindow, S. E. (1995). The secret life of foliar bacterial pathogens on leaves. Annual Review of Phytopathology, 33(1), 145–172. 77. Quiñones, B., Dulla, G., & Lindow, S. E. (2005). Quorum sensing regulates exopolysaccharide production, motility, and virulence in Pseudomonas syringae. Molecular Plant-Microbe Interactions, 18(7), 682–693. 78. Bailey, M. J., Lilley, A. K., & Diaper, J. P. (1996). Gene transfer between micro-organisms in the phyllosphere. In Aerial plant surface microbiology (pp. 103–123). Boston, MA: Springer. 79. Pearce, D., Bazin, M. J., Lynch, J. M., Lappin-Scott, H. M., & Costerton, J. W. (1995). The rhizosphere as a biofilm. Microbial Biofilms, 207–220. 80. Dennis, P. G., Miller, A. J., & Hirsch, P. R. (2010). Are root exudates more important than other sources of rhizodeposits in structuring rhizosphere bacterial communities? FEMS Microbiology Ecology, 72(3), 313–327. 81. Timmusk, S., Paalme, V., Pavlicek, T., Bergquist, J., Vangala, A., Danilas, T., et al. (2011). Bacterial distribution in the rhizosphere of wild barley under contrasting microclimates. PLoS ONE, 6(3), e17968. 82. Van Nieuwenhove, C., Van Holm, L., Kulasooriya, S. A., & Vlassak, K. (2000). Establishment of Azorhizobium caulinodans in the rhizosphere of wetland rice (Oryza sativa L.). Biology and Fertility of Soils, 31(2), 143–149. 83. Yanni, Y. G., Rizk, R. Y., Corich, V., Squartini, A., Ninke, K., & Philip-Hollingsworth et al. (1997). Natural endophytic association between Rhizobium leguminosarum bv. trifolii and rice roots and assessment of its potential to promote rice growth. In Opportunities for biological nitrogen fixation in rice and other non-legumes (pp. 99–114). Dordrecht: Springer. 84. Kim, C., Kecskés, M. L., Deaker, R. J., Gilchrist, K., New, P. B., Kennedy, I. R., et al. (2005). Wheat root colonization and nitrogenase activity by Azospirillum isolates from crop plants in Korea. Canadian Journal of Microbiology, 51(11), 948–956. 85. Iniguez, A. L., Dong, Y., & Triplett, E. W. (2004). Nitrogen fixation in wheat provided by Klebsiella pneumoniae 342. Molecular Plant-Microbe Interactions, 17(10), 1078–1085. 86. Hawes, M. C., & Smith, L. Y. (1989). Requirement for chemotaxis in pathogenicity of Agrobacterium tumefaciens on roots of soil-grown pea plants. Journal of Bacteriology, 171(10), 5668–5671. 87. Balandreau, J., Viallard, V., Cournoyer, B., Coenye, T., Laevens, S., & Vandamme, P. (2001). Burkholderia cepacia genomovar III is a common plant-associated bacterium. Applied and Environment Microbiology, 67(2), 982–985. 88. Bittel, P., & Robatzek, S. (2007). Microbe-associated molecular patterns (MAMPs) probe plant immunity. Current Opinion in Plant Biology, 10(4), 335–341. 89. Van Peer, R., Niemann, G. J., & Schippers, B. (1991). Induced resistance and phytoalexin accumulation in biological control of Fusarium wilt of carnation by Pseudomonas sp. strain WCS 417 r. Phytopathology, 81(7), 728–734. 90. Wei, G., Kloepper, J. W., & Tuzun, S. (1991). Induction of systemic resistance of cucumber to Colletotrichum orbiculare by select strains of plant growth-promoting rhizobacteria. Phytopathology, 81(11), 1508–1512. 91. Van Loon, L. C. (2007). Plant responses to plant growth-promoting rhizobacteria. In New perspectives and approaches in plant growth-promoting Rhizobacteria research (pp. 243–254). Dordrecht: Springer. 92. Ongena, M., Jourdan, E., Adam, A., Paquot, M., Brans, A., Joris, B., et al. (2007). Surfactin and fengycin lipopeptides of Bacillus subtilis as elicitors of induced systemic resistance in plants. Environmental Microbiology, 9(4), 1084–1090. 93. Iavicoli, A., Boutet, E., Buchala, A., & Métraux, J. P. (2003). Induced systemic resistance in Arabidopsis thaliana in response to root inoculation with Pseudomonas fluorescens CHA0. Molecular Plant-Microbe Interactions, 16(10), 851–858. 94. Schuhegger, R., Ihring, A., Gantner, S., Bahnweg, G., Knappe, C., Vogg, G., et al. (2006). Induction of systemic resistance in tomato by NacylLhomoserine lactoneproducing rhizosphere bacteria. Plant, Cell and Environment, 29(5), 909–918.

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112. Spiliopoulou, A. I., Kolonitsiou, F., Krevvata, M. I., Leontsinidis, M., Wilkinson, T. S., Mack, D., et al. (2012). Bacterial adhesion, intracellular survival and cytokine induction upon stimulation of mononuclear cells with planktonic or biofilm phase Staphylococcus epidermidis. FEMS Microbiology Letters, 330(1), 56–65. 113. Zhou, Y., Guan, X., Zhu, W., Liu, Z., Wang, X., Yu, H., et al. (2014). Capsaicin inhibits Porphyromonas gingivalis growth, biofilm formation, gingivomucosal inflammatory cytokine secretion, and in vitro osteoclastogenesis. European Journal of Clinical Microbiology and Infectious Diseases, 33(2), 211–219. 114. Takayama, S., Saitoh, E., Kimizuka, R., Yamada, S., & Kato, T. (2009). Effect of eel galectin AJL-1 on periodontopathic bacterial biofilm formation and their lipopolysaccharide-mediated inflammatory cytokine induction. International Journal of Antimicrobial Agents, 34(4), 355– 359. 115. Zhang, L., & Mah, T. F. (2008). Involvement of a novel efflux system in biofilm-specific resistance to antibiotics. Journal of Bacteriology, 190(13), 4447–4452. 116. De Kievit, T. R., Parkins, M. D., Gillis, R. J., Srikumar, R., Ceri, H., Poole, K., et al. (2001). Multidrug efflux pumps: expression patterns and contribution to antibiotic resistance in Pseudomonas aeruginosa biofilms. Antimicrobial Agents and Chemotherapy, 45(6), 1761–1770. 117. Soto, S. M. (2013). Role of efflux pumps in the antibiotic resistance of bacteria embedded in a biofilm. Virulence, 4(3), 223–229. 118. Kvist, M., Hancock, V., & Klemm, P. (2008). Inactivation of efflux pumps abolishes bacterial biofilm formation. Applied and Environment Microbiology, 74(23), 7376–7382. 119. Yoon, E. J., Chabane, Y. N., Goussard, S., Snesrud, E., Courvalin, P., Dé, E., et al. (2015). Contribution of resistance-nodulation-cell division efflux systems to antibiotic resistance and biofilm formation in Acinetobacter baumannii. MBio, 6(2), e00309–e00315. 120. Maira-Litran, T., Allison, D. G., & Gilbert, P. (2000). An evaluation of the potential of the multiple antibiotic resistance operon (mar) and the multidrug efflux pump acrAB to moderate resistance towards ciprofloxacin in Escherichia coli biofilms. Journal of Antimicrobial Chemotherapy, 45(6), 789–795. 121. Liao, J., Schurr, M. J., & Sauer, K. (2013). The MerR-like regulator BrlR confers biofilm tolerance by activating multidrug efflux pumps in Pseudomonas aeruginosa biofilms. Journal of Bacteriology, 195(15), 3352–3363. 122. Anderl, J. N., Franklin, M. J., & Stewart, P. S. (2000). Role of antibiotic penetration limitation in Klebsiella pneumoniae biofilm resistance to ampicillin and ciprofloxacin. Antimicrobial Agents and Chemotherapy, 44(7), 1818–1824. 123. Walters, M. C., Roe, F., Bugnicourt, A., Franklin, M. J., & Stewart, P. S. (2003). Contributions of antibiotic penetration, oxygen limitation, and low metabolic activity to tolerance of Pseudomonas aeruginosa biofilms to ciprofloxacin and tobramycin. Antimicrobial Agents and Chemotherapy, 47(1), 317–323. 124. Werner, E., Roe, F., Bugnicourt, A., Franklin, M. J., Heydorn, A., Molin, S., et al. (2004). Stratified growth in Pseudomonas aeruginosa biofilms. Applied and Environment Microbiology, 70(10), 6188–6196. 125. Anderl, J. N., Zahller, J., Roe, F., & Stewart, P. S. (2003). Role of nutrient limitation and stationary-phase existence in Klebsiella pneumoniae biofilm resistance to ampicillin and ciprofloxacin. Antimicrobial Agents and Chemotherapy, 47(4), 1251–1256. 126. Stewart, P. S. (2002). Mechanisms of antibiotic resistance in bacterial biofilms. International Journal of Medical Microbiology, 292(2), 107–113. 127. Mah, T. F. C., & O’Toole, G. A. (2001). Mechanisms of biofilm resistance to antimicrobial agents. Trends in Microbiology, 9(1), 34–39. 128. Fletcher, M., & Savage, D. C. (Eds.) (2013). Bacterial adhesion: Mechanisms and physiological significance. Berlin: Springer Science & Business Media. 129. Blenkinsopp, S. A., & Costerton, J. W. (1991). Understanding bacterial biofilms. Trends in Biotechnology, 9(1), 138–143.

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130. Davies, D. (2003). Understanding biofilm resistance to antibacterial agents. Nature Reviews Drug Discovery, 2(2), 114. 131. Drenkard, E., & Ausubel, F. M. (2002). Pseudomonas biofilm formation and antibiotic resistance are linked to phenotypic variation. Nature, 416(6882), 740. 132. Gilbert, P., Das, J., & Foley, I. (1997). Biofilm susceptibility to antimicrobials. Advances in Dental Research, 11(1), 160–167. 133. Drenkard, E. (2003). Antimicrobial resistance of Pseudomonas aeruginosa biofilms. Microbes and Infection, 5(13), 1213–1219. 134. Mah, T. F. (2012). Biofilm-specific antibiotic resistance. Future Microbiology, 7(9), 1061– 1072. 135. Gilbert, P. E. T. E. R., Collier, P. J., & Brown, M. R. (1990). Influence of growth rate on susceptibility to antimicrobial agents: Biofilms, cell cycle, dormancy, and stringent response. Antimicrobial Agents and Chemotherapy, 34(10), 1865. 136. Lewis, K. (2008). Multidrug tolerance of biofilms and persister cells. In Bacterial biofilms (pp. 107–131). Berlin, Heidelberg: Springer. 137. Lewis, K. (2005). Persister cells and the riddle of biofilm survival. Biochemistry (Moscow), 70(2), 267–274. 138. LaFleur, M. D., Kumamoto, C. A., & Lewis, K. (2006). Candida albicans biofilms produce antifungal-tolerant persister cells. Antimicrobial Agents and Chemotherapy, 50(11), 3839– 3846. 139. Keren, I., Shah, D., Spoering, A., Kaldalu, N., & Lewis, K. (2004). Specialized persister cells and the mechanism of multidrug tolerance in Escherichia coli. Journal of Bacteriology, 186(24), 8172–8180. 140. Lewis, K. (2007). Persister cells, dormancy and infectious disease. Nature Reviews Microbiology, 5(1), 48.

Chapter 4

Biofilms in Nature and Artificial Materials

Abstract This chapter presents various substrates and their capabilities for biofilm formation, taking into account influential factors like van der Waals forces, hydrophobicity, hydrophilicity, the presence of polar side chains on polymers, and more. It includes information about natural substrates such as roots of plants and rocks (which are slimy in rivers when covered with biofilm). Also the artificial substrates of metals, ceramics, and polymers are described in terms of their interaction with bacteria and the formation/control of biofilms. Studies have been carried out with ceramic materials used in dentistry. The results showed that the greater the surface roughness in crowns, etc. the greater the accumulation of biofilm (called plaque in its hardened form). As for metals, silver has an antibacterial action that depends on the silver ion. It interrupts the ability of a bacterial cell to form chemical bonds that are necessary for survival.

4.1 Natural Substrates Firstly, when the concept of biofilms was better understood, people got to know why rocks and sand in the rivers and seas are slimy and sticky. As shown in these examples, natural solid matter such as rocks, sand, etc. form biofilms when they are located in humid environments. In seas, lakes, and rivers, every rock has its own biofilm. As described many times, biofilms are formed by bacterial activities. Since biofilms contain polymeric substances, other larger organisms are attracted to them because they are abundant sources of nutrition. Biofilms can be seen everywhere in nature. Even though the concept of biofilm was proposed in the late 1970s or 1980s, they have existed for a long time. They are very important for bacteria because they protect them from antibiotics, etc. According to some biological research results [1, 2], the mass generation of cyanobacteria occurred around 37 billion years ago [3]. Before the biological event, we presumed that our primary earth did not have much oxygen. Therefore, anaerobic bacteria had to exist. Since then, our atmosphere started to have oxygen and everything changed. Figure 4.1 shows the schematic illustration for the metabolism of cyanobacteria.

© Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_4

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Fig. 4.1 The outline of metabolism for cyanobacteria

The traits of biofilms can be seen in fossils of stromatolites [4–8] even today. Stromatolites are a sort of rock-like matter or mound and are found in shallow rivers, seas, etc. They contain layers of lime-secreting cyanobacteria and trapped sediment. They are found in Precambrian rocks as the earliest fossils. Therefore, these fossils demonstrate the existence of bacteria and biofilms around that time period. Figure 4.2 shows the schematic illustration about the mechanism of the formation process for stromatolites. We already know that bacteria exist in biofilms and that more than 98% of bacteria on earth do not exist as planktonic bacteria (individuals), but exist cooperatively in biofilms. In addition, the analytical results about various stromatolites help us predict not only the archaeological data of microorganisms, but also the existence of biofilms

Fig. 4.2 The formation of stromatolites

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themselves. According to such an investigation, we could predict that more than 50% of the biomass on earth would belong to biofilms. As described above, bacteria form biofilms and mostly exist in them. Also biofilms (formed by various bacteria) on rocks accumulate in the soil layer by layer. As a result, microbial mats are formed. The microbial mat is defined as the association of bacteria formed at the interfaces between a water phase and a solid one or between a water phase and air. The environment generally changes with time, and affects the microbiota. The slightly changing factors with time are temperatures, oxygen concentrations, nutrient salt concentrations, light qualities (how to be irradiated by light), redox components, etc. Therefore, the changing microbiota produces the corresponding unique biomat layer by layer [9–11]. Such a biomat could show us colorful and magnificent views of nature [12–14]. Figure 4.3 provides some examples. All photos in Fig. 4.3 were taken by the authors in Yellow Stone National Park in the US. As you can see from the photos, the color changes from location to location, which corresponds to differences of the microbial biota present. Biofilms (on rocks, on the ground, in the oceans, lakes, ponds, and rivers, etc.) form microbial biomats. Volcanic action, crustal movements, etc. also add to the biological phenomena to produce magnificent views in nature. Actually people don’t really know very much about the contributions of biofilms. The other important aspect about biofilms in nature is that they form on plants’ leaves, stems, roots, etc. In particular, roots have lots of nutrients and affect areas around them in various ways. The area affected by roots is called the rhizosphere. Note that each area around various parts of plants has its own name, as shown in Fig. 4.4. The area above ground for plants is called phyllosphere. The rhizosphere is classified into three categories: endorhizosphere, rhizoplane and the exorhizosphere. The endorhizosphere is the inside of the roots. The rhizoplane is the surface of a root. The exorhizosphere corresponds to the area around the roots that is affected by the roots. The area other than the rhizosphere in soils is called the “bulk soil”. The size of the rhizosphere is generally several millimeters, even though it depends on the size of the roots and species of the plants. Due to the abundance of nutrients, the rhizosphere forms biofilms on the roots’ surfaces and changes the qualities and properties of areas where they exist. For example, abundant nutrients of biofilms in the rhizosphere attract bacteria which form a certain unique bacterial biota. Since biofilms contain lots of water components, rhizospheres with abundant biofilms might somewhat change arid regions to humid sites. Such a change could lead to an ecological system (ecosystem) that drastically affects our food supply (Fig. 4.5).

4.2 Artificial Substrates Artificial substrates (materials) are classified into three main types based on the viewpoint of binding forces. They include metallic materials, ceramics, and polymers (Fig. 4.6) [15]. Metallic materials are constituted through metallic bonding. As shown in Fig. 4.7,

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Fig. 4.3 Concrete examples of microbial mats observed in Yellow Stone National Park

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Fig. 4.3 (continued)

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Fig. 4.4 Rhizosphere and phyllosphere

Fig. 4.5 Biofilms in the rhizosphere and the ecological (ecosystem) system

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Fig. 4.6 A general classification of industrial materials and their bonding

Fig. 4.7 Metallic surfaces and metallic ions

the equilibrium between metals and metallic ions is usually established. Biofilm formation is affected by the interaction between metallic ions and bacteria or excreted EPS. Figure 4.8 shows the schematic illustration for the concept. In ceramics, atoms are bound to each other by ionic bonds and covalent bonds. For some ceramics, the surface atoms could react with organic and inorganic materials derived from organisms. For others, the surface might be inert. According to such a standard, the reactive ceramics are glass, carbon materials and calcium phosphate such as hydroxyl apatite, etc. On the other hand, the inert ceramics are oxides such as alumina, zirconia, nitrides such as silicon nitrides, etc. (Figure 4.9). The above diagram displays the viewpoint of interactions between ceramics and bacteria/biofilms. As already described, we have bioactive ceramics and bio-inert ceramics from the viewpoint of biofilm formation. Ceramics usually become bioactive, when biopolymers such as proteins bind to them for various reasons. Hydroxyapatite (Ca10 PO4 (OH)2 ) can easily exchange surface ions with those in the environment. Therefore, the electrostatic force between ions would work fairly well, as shown in Fig. 4.10. Organic materials are constituted mainly through covalent bonding and van der Waals forces. When we come to think about the correlation between organic polymeric materials, polymers should be classified into two types: polar polymers and

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Fig. 4.8 Metallic materials and correlated forces Fig. 4.9 The classification of ceramics from the viewpoint of reactions with biofilms

Fig. 4.10 Ceramics and involved forces at the vicinity of surfaces

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non-polar ones. The demarcation is sometimes difficult and delicate, since macromolecule polymers are polarized locally to some extent. However, the big difference between the two kinds of polymers could be contributed to the dominant and peculiar side chains of their structures that are composed of hydroxyl, amino, and carboxyl groups (Fig. 4.11). Biofilms form at the interface between solid and gas phases or between solid and liquid phases. When we use the technical term, “interphase,” we anticipate the area that is around the interface. It has the potential profiles, as shown in Fig. 4.12, due to the formation of electric double layers at the interface. The figure shows the interphase between solid and liquid phases. However, the ambient atmosphere usually contains humidity. Therefore, the interphase in the case between solid and gas phases can be correlated like that in Fig. 4.12. When two different phases have contact with each other, the potential profile is produced due to the difference of Fermi energy. The rearrangement of electric charges leads to the formation of electric double layers [16–18]. According to the Bockris–Devanathan–Müller model, the electric double layer is composed of an inner Helmholz layer and an outer Helmholz one (where non-hydrated anions specifically absorb on solid surfaces and hydrated cations are arranged, respectively). The diffusion layer exists on the outside of the electric double layer. The potential continues to gradually decrease with an increasing distance from the surface in the diffusion layer. The decrease also continues outside of the diffusion layer. The characteristic potential at the boundary between the Helmholz double layer and the diffusion layer is called the Stern potential. The border between the diffusion layer and the bulk fluid is called a slipping plane and the corresponding potential is called ζ (zeta) potential. The potential drops with increasing distances from that, corresponding to the Fermi energy of the solid phase through the two layers. Then the potential also continues to decrease in the diffusion layer and bulk layer outside of the slipping plane with changing rates, respectively. On the other hand, bacteria approaching the solid surface also have the electrical double layer at the interphase between their outer membrane and liquid phase. When bacteria approach a solid surface, their double

Fig. 4.11 The classification of polymers from the viewpoint of biofilm formation

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Fig. 4.12 Schematic image of electric double layer of materials’ surfaces and zeta potential

layers overlap with each other. It means that they repel each other because of osmotic pressure. Simultaneously, other factors might be added. For example, van der Waals forces work between two matters (solids and bacteria) for covalent bonding surfaces. When the force of the latter exceeds the former, then bacteria attach to solid surfaces [19–21]. On the contrary, if the former exceeds the latter, then bacteria can’t attach to solid surfaces, as shown in Fig. 4.13.

Fig. 4.13 Polymers and involved forces at the vicinity of surfaces

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The general force relationship between bacteria and a material’s surface (solid) is just like what is described above. In addition to the general tendency, other factors (depending on the kinds of materials involved) would be influential too. Therefore, the materials involved would profoundly affect the characteristics of biofilm formation.

4.3 Metals Biofilms can grow on different surfaces including floors, countertops, slimy rocks, medical devices, and more. They are produced by bacteria to provide a protective outer casing against potential threats such as detergents and antibiotics. Biofilms start to form when free floating microorganisms (like bacteria) become attached to a surface. Studies show that some metals (especially their metal ions) are useful for controlling biofilms because they are toxic to bacteria. The use of Silver for antimicrobial purposes dates back thousands of years and it is still being used today [22]. Silver is a chemical element with the symbol Ag (from the Latin argentum). It is a shiny white transition metal with the atomic number 47. It is located in group 11 and period 5 of the periodic table. Silver has very high electrical and thermal conductivity and is used in jewelry, currency, etc. SILVER’S medical applications are diverse. Dilute solutions of silver nitrate and other silver compounds are used as disinfectants and microbiocides. For example, they are often added to bandages. See the drawing in Fig. 4.14 [23]. Silver is used in wound dressings, creams, and as an antibiotic coating on medical devices. A limited amount of evidence shows that silver coatings on endotracheal breathing tubes may reduce the occurrence of pneumonia associated with ventilators [24]. Also the use of silver-alloy with indwelling catheters for short term catheterizing will reduce the risk of catheter-acquired urinary tract infections [25]. Silver and most silver compounds have an oligodynamic effect and are toxic for bacteria. The oligodynamic effect is generally defined as the ability of small amounts of heavy metals to kill bacterial cells. The antibacterial action of silver depends on the silver ion, Ag+ . Silver interrupts the ability of a bacterial cell to form the chemical bonds that are necessary for survival [26]. These bonds are needed to produce the physical structure of the cell. Also applying an electric current across silver electrodes enhances the antibiotic action [27]. One study involving silver nanoparticles suggests that the antimicrobial activity could be controlled by regulating the silver ion release, possibly by particle size, shape, etc. [28]. Investigators for another project, to better understand the antimicrobial action of silver, carried out a chemical genetic screen of a mutant library of Escherichia coli (the Keio collection) to identify silver sensitive or resistant deletion strains [29]. Their data shows that the activity of silver within the bacterial cell is extensive. It encompasses genes involved in cell wall maintenance, quinone metabolism and sulfur assimilation.

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Fig. 4.14 Above is a display of a bandage containing silver compounds

COPPER is an element with the symbol Cu and the atomic number 29. It is a soft metal with high thermal and electrical conductivity. It is also a trace dietary mineral and is in group 11 and period 4 of the Periodic Table. Medical use of copper dates back many, many years. An Egyptian medical text, that was written about 2000 B.C. describes the use of copper to sterilize drinking water and wounds [30]. A 1983 report documented the benefits of using brass (copper and zinc) and bronze (copper and tin) on doorknobs in hospitals to prevent the spread of microbes [31]. Currently there is interest in using metallic copper as an antimicrobial surface. Bacteria, yeasts, and viruses are quickly killed on metallic copper surfaces. This process is referred to as contact killing. In many studies for contact killing, a wet inoculation technique is used by applying a small volume of cell suspensions to coupons. However, this might not be similar to the dry copper surfaces in health care environments. For the alternative dry method, a small volume of liquid is added to the coupon with a cotton swab. The thin film’s liquid evaporates quickly and allows the cells to have direct contact with the metal surface. For a study using these conditions, E. coli and other bacteria were inactivated within a few minutes of exposure to the copper surface [32]. The work showed that large amounts of copper ions were taken up by E. coli during the killing phase, which suggests that the cells became overwhelmed by the copper. Cell survival on metallic copper is also influenced by oxidative stress (from the generation of reactive oxygen species). Research results show that the absence of oxygen did not prevent contact killing of

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E. coli, but increased the amount of time for it to take place [33]. This process seems to target the cell membrane, leading to a loss of cytoplasmic content. Touch surfaces (like door handles, toilet seats, bed rails, etc.) in hospitals contain microbes. Therefore, copper surfaces (with their antimicrobial properties) could provide protection from infectious microbes by reducing contamination. Successful hospital trials support the above statement. For example, a trial was carried out in a consulting room of a primary health care clinic in South Africa. In this case, items (like desks, window sills, cupboards) frequently contacted by patients and staff, were covered with copper sheets. For six months the surfaces were sampled every six weeks for a four to five day period with multiple samples per day. There was an overall 71% reduction in the bacterial load of the copper surfaces as compared to the control surfaces, non copper surfaces such as aluminum, etc. [34]. A different type of study investigated the antimicrobial effect of copper surfaces on bacteria isolated from poultry meat. The aim of this study was to evaluate the antibacterial effect of copper surfaces on the transmission of two food borne pathogens (Salmonella enteric and Listeria monocytogenes) and a poultry bacterial species (Entero bacter cloacae). The results showed that copper surfaces quickly reduced (in less than four minutes) the bacterial load when the microorganisms were exposed to polished copper surfaces. This suggests that copper surfaces have a potential role in controlling the microbiological hazards of the poultry industry [35]. ZINC is an element with the symbol Zn and atomic number 30. It has a bluishwhite appearance, is a component of brass, and an essential mineral. Zinc has antimicrobial properties too. Its ions (Zn+2 ) show antimicrobial activity against a variety of fungal and bacterial strains [36]. A study was carried out to investigate the antibacterial effect of zinc oxide (ZnO) nanoparticles (NP) on Campylobacter jejuni, a leading cause of microbial food-borne illness worldwide. The results showed that C. jejuni was very sensitive to treatment with ZnO nanoparticles. Examination by scanning electron microscopy showed that many of the bacterial cells changed from spiral shapes to coccoid forms after being exposed to 0.5 mg/ml of ZnO nanoparticles for 16 h, which is consistent for the changes of C. jejuni when it is under stress conditions. The overall results of the investigation suggest that the antibacterial mechanism of ZnO nanoparticles is most likely due to oxidative stress and disruption of the cell membrane [37]. Since ZnO is generally recognized as safe by the Food and Drug Administration (21 CFR 182.8991), it has been used in the linings of cans containing food items such as meat, corn, peas, etc. to prevent spoilage. A project, supporting this use, compared the antibacterial potential of zinc oxide nanoparticles with conventional ZnO powder against food-borne bacteria. The researchers found the nanoparticles to be more efficient and that Gram positive bacteria were generally more sensitive to ZnO than the Gram negative bacteria. For example, exposure of Salmonella typhimurium (Gram negative bacteria) and Staphylococcus aureus (Gram positive bacteria) to their relevant minimal inhibitory concentrations from ZnO nanoparticles, reduced the cell number to zero within 8 and 4 h respectively [38]. Coatings (that contain silver, copper, etc.) are also used to provide substrates with antibacterial properties. Some metal alloys like brass have these properties too.

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Since metallic materials are used for biomaterials, they need to have good mechanical characteristics, and resistance to corrosion and biofilm formation. STAINLESS STEEL is a good candidate for biomaterials, especially because of a new strategy where antibacterial surfaces are created by using nanotechnology on diverse materials. Stainless steel is an alloy that has about 10% chromium by mass and about 1.2% carbon by mass. It is noted for corrosion resistance, which increases as the chromium content increases. Also the addition of molybdenum increases corrosion resistance. A variety of stainless steels exist. Bacterial adhesion to stainless steel 316L (SS316L), which is generally used in medical devices (such as surgical tools and orthopedic implants), can cause serious infections. Researchers have demonstrated that nanotextured SS316L surfaces (prepared by electrochemical etching) inhibit bacterial adhesion of both Gram-negative Escherichia coli and Gram-positive Staphylococcus aureus [39]. The nanotextured SS316L (which contains nanopores and nanoprotrusions) offers great potential for medical applications. It provides a reduction of bacterial adhesion without using antibiotics or chemical modification. At the same time, it has corrosion resistance and exhibits cytocompatibility and no toxicity toward mammalian cells in vitro.

4.4 Ceramics Ceramics are generally inorganic, non-metallic materials made from compounds of a metal and non metal, formed by the process of heat and subsequent cooling. Ceramic substrates include a wide variety of materials, so it is almost impossible to come up with an exact definition for them. However, they can be divided into two main classes. One is called traditional ceramics, which includes clay products like pottery, silicate glass, and cement. See Fig. 4.15. [40]. The other class is referred to as advanced ceramics, which includes silicon carbide (SiC), oxides like aluminum oxide (Al2 O3 ), nitrides like silicon nitride (Si3 N4 ), non-silicate glass, etc. A general description of some examples of advanced ceramics is provided. Aluminum nitride (AlN) has high thermal conductivity, excellent thermal shock resistance, and corrosion resistance. It is used in power electronics and aeronautical systems. Figure 4.16 shows aluminum nitride powder [41]. Zirconium dioxide (ZrO2 ), also known as zirconia, is a white crystalline oxide of zirconium. It is used as a refractory material and in the production of hard ceramics (like those used in dentistry). Also because of its high ionic conductivity and low electronic conductivity, it is very useful for electroceramics. Silicon nitride (Si3 N4 ) is a white solid with a high melting point. It is very hard and has high thermal stability. It is used for engine parts in automobiles, ball bearings, cutting tools, and insulators. Silicon nitride ceramics have good shock resistance too. Silicon carbide (SiC), which is mostly synthetic, is also known as carborundum. It has high hardness, wear resistance, and excellent chemical resistance. Silicon carbide is a semiconductor that has many uses. Several of its uses include abrasives, cutting tools, the production of graphene, the hard ceramics in car brakes, etc. See Fig. 4.17 [42].

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Fig. 4.15 A traditional ceramic dome is displayed

Fig. 4.16 Aluminum nitride powder

Aluminum oxide (Al2 O3 ) is produced from bauxite and often called alumina. Its most common crystalline form is corundum. Because of its hardness, it is used as an abrasive and as a component in cutting tools. See Fig. 4.18 [43]. Alumina is also an electrical insulator, but has relatively high thermal conductivity for a ceramic material. In addition, it is important in the production of hard ceramics such as crowns and implants used in dentistry. Researchers investigated the surface characteristics and biofilm development on selected dental ceramic materials. Dental ceramics are used in restorative dentistry that includes crowns and implants. These materials are used because they are biocompatible, have low thermal conductivity, color stability, and aesthetics. Dental bioceramics include a wide range of glass-ceramics, reinforced porcelains, oxide

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Fig. 4.17 A silicon carbide sample used for making steel

Fig. 4.18 A sample of corundum from Brazil

ceramics (zirconia, alumina, and spinel), fiber–reinforced ceramic composites, and multilayered ceramic structures. The aims of this study were to characterize the surface ultrastructure and roughness of four ceramic materials (used by dentists and sold commercially) and to assess their promotion of biofilm development following adjustments that simulated clinical intraoral polishing. The four ceramic materials were prepared, adjusted, and polished, following the manufacturers’ instructions. Specimens of each type were analyzed by SEM imaging, confocal microscopy, and crystal violet assay. As a result, the SEM images showed more irregular surface topography in adjusted specimens than their

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respective controls did. It was concluded that simulated intraoral polishing methods resulted in greater surface roughness and increased biofilm accumulation [44]. In addition, Haroon Rashid reviewed the literature to find out the overall effect of surface roughness on ceramics used in dentistry [45]. He found that a rough material surface accumulates more plaque (hardened biofilm). Another investigation was carried out to evaluate the formation of oral biofilm on various dental ceramics in vivo [46].The five different ceramic materials that were tested include a veneering glass-ceramic, a lithium disilicate glass-ceramic, a yttrium-stabilized zirconia (Y-TZP), a hot isostatically pressed (HIP) Y-TZP ceramic, and HIP Y-TZP ceramic with 25% alumina. Test specimens were attached to acrylic appliances that were worn by five volunteers in their maxillary arch for 24 h. Then the samples were removed from the appliances and the biofilms were stained. The two-dimensional surface coating and thickness of the biofilms were determined by using confocal laser scanning microscopy. The results showed that the lowest surface coating and biofilm thickness were on sample HIP Y-TZP ceramic. The highest amounts were found on the lithium disilicate glass-ceramic. It was concluded that zirconia exhibited a low level of plaque accumulation. Other researchers found reduced bacterial adhesion on ceramics used for arthroplasty applications [47]. Bacterial adhesion depends on a material’s surface features (including their chemical and physical properties). This study compared materials used for bearings of total hip ceramics (alumina and zirconia-platelet toughened alumina composites), metals (cobalt-chromium-molybdenum alloy), and polymers (highly cross-linked polyethylene). They were examined in terms of wettability and protein adsorption. Specimens were infected with Staphylococcus aureus and Staphylococcus epidermidis biofilm for 24 or 48 h. Evaluations of bacterial adhesion properties were made in terms of biofilm viability, thickness, the condition of surface roughness, etc. As compared to the metal and polymer surfaces, the bioceramics displayed reduced bacterial adhesion and less biofilm formation. In a new study, an advanced engineering technology was successfully applied to TiAl4 V external fixation pins. It actually converted the surface of the self-drilling pins into a hard ceramic coating. The surface of the TiAl4 Vexternal fixation pins was converted into a TiO2 layer supported by an oxygen hardened case [48]. Characterization of these treated pins was determined by using a scanning electron microscope (SEM), X-ray diffraction, etc. External, fracture fixation is a common orthopedic procedure that involves the placement of pins and/or wires to support a limb. Stainless steel and titanium self-drilling pins are good candidates for the procedure because they do not require pre-drilling. Stainless steels have been used for external fixation pins, due to their corrosion resistance, good mechanical properties, and adequate biocompatibility. However, these pins often lead to pin track infections because the pin entry site is exposed to the environment and permits an interface between bone and internal soft tissues (with the external stainless steel pin). Titanium dioxide has been found to have antibacterial properties [49–51]. Therefore, the researchers decided to carry out a procedure to convert the titanium alloy surface (of the titanium type pins) into a TiO2 ceramic layer. For testing, both treated and untreated pins were inserted into high density bone simulation material. Also

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Staphylococcus aureus was used to assess the antibacterial behavior. The post testing SEM results revealed that the treated pins (with the improved surface hardness) performed better than the untreated ones. They had less wear and lower numbers of bacteria than the untreated pins.

4.5 Polymers Polymers’ surfaces are constituted by covalent bonds. Therefore, van der Waals forces could affect the binding between materials’ surfaces and bacteria, when bacteria try to attach to materials’ surfaces. Thus, the balance between repulsion by the overlapping of the electric double layers and the attraction due to van der Waals forces might determine the possibility of a successful attachment. However, other forces might work in favor of the polymers. Even though other factors might be involved for the dynamic balance, we could mention hydrophobicity and hydrophilicity as the most important factors. Hydrophilicity is the characteristic of being water loving and binding water molecules easily. On the other hand, hydrophobicity is the act of repelling water. For polymers, some side chains have hydroxyl, carboxyl, and amino groups that are locally polarized and generally show hydrophilicity. However, C–H and C–F bonds are not so polarized and show hydrophobicity [52–54]. When polymers are compared with metallic materials, the polymers don’t interact with EPS or bacteria very strongly. Therefore, polymers don’t form biofilms as much as metallic materials. However, when you compare the capability of biofilm formation among polymers, hydrophilicity and hydrophobicity are important key factors. Generally, people say that bacteria tend to attach to hydrophobic materials’ surfaces more easily. As shown in Fig. 4.19, hydrophobic surfaces would remove water molecules. When hydrophobic surfaces of materials and bacteria come closer to each other, both of them tend to remove water molecules and form their “voids”. Then they could easily bind to each other. That is the reason why hydrophobic surfaces are favorable to form biofilms. However, this is only one factor for biofilm formations— in the light of bacterial attachment. As already described, many other factors would be involved in biofilm formation and growth. If other factors would dominate this one factor, then the hydrophobic surface might not work well to control biofilm formation. For this viewpoint, biofilm formation capability could not be explained just by hydrophilic—hydrophobic characteristics. However, the hydrophobic-hydrophilic scale could be sort of standard for us to evaluate biofilm formation capability to some extent, especially when polymer materials have polarization within their structures [55, 56]. On the other hand, there are some polymers that have “water repellency”. Concretely speaking, polytetrafluoroethylene (PTFT) can be mentioned as a typical example (Fig. 4.20). The polymer does not have any polar characters. Therefore, van der Waals forces don’t work between the polymer and other molecules. The unique

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Fig. 4.19 Hydrophilic/hydrophobic polymers and bacterial attachment

Fig. 4.20 Chemical structure of polytetrafluoroethylene (PTFE)

characteristics would lead to a strong repulsion force against water molecules. Actually, the contact angle (Fig. 4.21) of water on PTFT would be more than 70° and might reach 120–150° in some cases. From the viewpoint of biofilm formation, various conditions and factors need to be considered. Figure 4.22 shows the relationship between each process of biofilm formation and the factors involved. When we come to think about the contribution of polymers’ surfaces to biofilm formation, there are three steps for us to take into account. One of them is the conditioning film, the previous step for the attachment of bacteria. Since the attached matter would mostly be carbon compounds at this stage, van der Waals forces are the most important. Non-polar polymers would not form conditioning film easily, due to the lack of van der Waals forces between polymers’ surfaces and the attached carbon compounds. The phenomenon would affect the following attachment of bacteria and biofilm formation.

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Fig. 4.21 Surface tension, Young’s equation and water repellency

Fig. 4.22 A schematic illustration for the correlation among polymeric substrates: hydrophobicity/hydrophilicity, and water repellency, from the viewpoint of biofilm formation

At the second stage of bacterial attachment, the high water repellency of the surface might be favorable for bacterial attachment itself, since such a surface might get a chance to form a “water void” in the vicinity of the surface. However, non-polar surfaces would not attract EPS. Also only the repulsion due to the overlapping of the electric double layers between them would be dominant. As a result, the biofilm formation would be difficult (The third step). Even if it could form biofilms to some extent, they might detach from materials’ surfaces during the growth process.

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Since the biofilm formation process is composed of multi steps, the balance among forces would be very complicated. It is very important for us to know which process would be considered and how dominant the process would be against other steps to control biofilm formation.

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20. Rijnaarts, H. H., Norde, W., Lyklema, J., & Zehnder, A. J. (1999). DLVO and steric contributions to bacterial deposition in media of different ionic strengths. Colloids and Surfaces B: Biointerfaces, 14, 179–195. 21. Birdi, K. (1979). Adherence of bacteria to solid surfaces and the surface forces. Journal of Dentistry, 7, 230–234. 22. Alexander, J. W. (2009). History of the medical use of silver. Surgical Infections, 10(3), 289– 292. 23. Pearson Scott Foresman. File: Bandage (PSF) png. This work is in the public domain. Retrieved September 8, 2008 from https://commons.wikimedia.org/wiki/File:Bandage_(PSF).png. 24. Bouadma, L., Wolff, M., & Lucet, J.-C. (2012). Ventilator- associated pneumonia and its prevention. Current Opinion in Infectious Diseases, 25(4), 395–404. https://doi.org/10.1097/ qco.0b013e328355a835. 25. Lederer, J. W., Jarvis, W. R., Thomas, L., & Ritter, J. (2014). Multicenter cohort study to assess the impact of a silver-alloy and hydrogel- coated urinary catheter on symptomatic catheterassociated urinary tract infections. Journal of Wound, Ostomy and Continence Nursing, 41(5), 473–480. 26. Barry, D. M., & McGrath, P. B. (2016). Rotation disk process to assess the influence of metals and voltage on the growth of biofilm. Materials, 9(7), 568. https://doi.org/10.3390/ma9070568. 27. Akhavan, O., & Ghaderi, E. (2009). Enhancement of antibacterial properties of Ag nanorods by electric field. Science and Technology of Advanced Materials, 10(1). https://doi.org/10.1088/ 1468-6996/10/1/015003. 28. Xiu, Z.-M., Zhang, Q. B., Puppala, H. L., Colvin, V. L., & Alvarez, P. J. (2012). Negligible particle-specific antibacterial activity of silver nanoparticles. Nano Letters, 12(8), 4271–4275. https://doi.org/10.1021/nl301934w. 29. Gugala, N., Lemire, J., Chatfield-Reed, K., Yan, Y., Chua, G., & Turner, R. J. (2018). Using a chemical genetic screen to enhance our understanding of the antibacterial properties of silver. Genes, 9(7), 344. https://doi.org/10.3390/genes9070344. 30. Dollwet, H. H. A., & Sorenson, J. R. J. (1985). Historic uses of copper compounds in medicine. Trace Elements in Medicine, 2, 80–87. 31. Kuhn, P. J. (1983). Doorknobs: A source of nosocomial infection? Copper Development Association, New York, NY. http://www.copperinfo.co.uk/antimicrobial/downloads/kuhn-doorknob. pdf. 32. Espirito Santo, C., et al. (2011). Bacterial killing by dry metallic copper surfaces. Applied and Environment Microbiology, 77, 794–802. 33. Espirito Santo, C., Taudte, N., Nies, D. H., & Grass, G. (2008). Contribution of copper ion resistance to survival of Escherichia coli on metallic copper surfaces. Applied and Environment Microbiology, 74, 977–986. 34. Marais, F., Mehtar, S., & Chalkley, L. (2010). Antimicrobial efficacy of copper touch surfaces in reducing environmental bioburden in a South African community healthcare facility. Journal of Hospital Infection, 74, 80–82. 35. Parra, A., Toro, M., Riocardo, J., Navarrete, P., Troncoso, M., Figueroa, G., et al. (2018). Antimicrobial effect of copper surfaces on bacteria isolated from poultry meat. Brazilian Journal of Microbiology, 49(1), 113–118. 36. Pasquet, J., Chevalier, Y., Pelletier, J., Couval, E., Bouvier, D., & Bolzinger, M.-A. (2014, September, 5) The contribution of zinc ions to the antimicrobial activity of zinc oxide. Colloids and Surfaces A: Physicochemical and Engineering Aspects, 457, 263–274. 37. Xie, Y., He, Y., Irwin, P. L., Jin, T., & Shi, X. (2011). Antibacterial activity and mechanism of action of zinc oxide nanoparticles against Campylobacter jejuni. Applied and Environmental Microbiology, 77(7), 2325–2331. 38. Tayel, A., El-Tras, W. F., Moussa, S., El-Baz, A. F. Mahrous, H., Salem, M. F., & Brimer, L. (2011). Antibacterial action of zinc oxide particles against food-borne pathogens. Journal of Food Safety. https://onlinelibrary.wiley.com/doi/abs/10.1111/j.1745-4565.2010.00287.x. 39. Jang, Y., Choi, W. T., Johnson, C. T., García, A. J., Singh, P. M., Breedveld, V., Hess, D. W., & Champion, J. A. (2018). Inhibition of bacterial adhesion on nanotextured stainless steel 316L

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Chapter 5

Laboratory Biofilm Reactors

Abstract Biofilms form at the interface between different phases in nature. To investigate biofilms and to solve practical problems, we have to reproduce the phenomena in laboratories. To achieve this purpose, we need to produce biofilms in laboratories and to evaluate them properly and accurately. As for the artificial production of biofilms, we need laboratory biofilm reactors. They have to mimic natural phenomena as much as possible, while the conditions should be idealized without some unnecessary factors. However, the simplification and idealization are very often difficult. It depends on the design and production of the laboratory biofilm reactors. In this chapter, we describe some representative laboratory biofilm reactors and discuss the efficacy and also the problems.

Biofilms form in nature. They affect our daily lives and cause many industrial problems. To solve these problems and/or utilize biofilms, we must first investigate various biofilm topics. Figure 5.1 shows the concept for the study of biofilm. As described above, biofilms originally form in nature. Therefore, it is best for us to investigate the natural biofilms which form in natural (or practical) environments. To mimic or to reproduce the biofilm phenomena in nature, we sometimes need to carry out experiments in natural environments. Such an experiment is called an in vivo experiment. However, natural environments generally have many complicated factors that affect biofilm’s formation and growth processes. To fix the mechanism and to solve such problems, scientists and engineers often simplify the complicated factors to reach a solution. In addition, a real phenomenon in the natural environment needs a relatively long time to occur. Actually, it may take anywhere from a couple of weeks to many decades. Therefore, to reduce the complex factors and also accelerate the test term, laboratory experiments are very often carried out in laboratory environments. Such a laboratory test is usually called an in vitro experiment. However, there are always worries about the discrepancy between natural (and practical) results in natural environments, since the factors might be too simplified or the experimental conditions might be too accelerated. To mediate the discrepancy, some intermediate experiments are carried out. These experiments have a slit orientation: half natural and half laboratory oriented. Such an experiment is called an ex vivo experiment. © Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_5

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Fig. 5.1 The concept of biofilm research

In this chapter, we introduce many possibilities for in vivo, ex vivo and in vitro experiments. Their problems are also discussed.

5.1 In Vitro Systems The research and development for biofilms are basically composed of two steps, as shown in Fig. 5.2. These steps include the production of biofilms and the evaluation process. The system, along with its environment, where biofilms would form is called the “Biofilm Reactor”. It may be natural or artificial. In any case, it should be called the “Biofilm Reactor”. When this process is carried out in a laboratory, the system is called a “Laboratory Biofilm Reactor” [1]. Laboratory biofilm reactors include two steps: the artificial production of biofilms and the evaluation of them. The basic principle for biofilm study is to make and measure biofilms. As for the evaluation process, the detailed information will be discussed in Chap. 6. On the other hand, the artificial production process of biofilms is discussed in this chapter. As described in the introduction, the experiments will be carried out in nature and in laboratories. In this section of the chapter, we focus on laboratory scale experiments. Usually, most of them correspond to in vitro systems. In these systems, where biofilm formation/growth would occur, the reactor is called a laboratory biofilm reactor (LBR). Lebeaux et al. wrote a good review paper about the laboratory test processes [2]. They divided the in vitro system into three types and named them as a Closed or Static system, Open or Dynamic system, and as a Microcosms system. The classification was reasonable and clear. However, we would like to classify them into three types that take into account the viewpoint of materials science and engineering. They are shown in Fig. 5.3.

5.1 In Vitro Systems

Fig. 5.2 Biofilm research and the steps involved

Fig. 5.3 Laboratory Biofilm Tests and their classification

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(1) Static systems This set almost overlaps with the static system for Lebeaux’s classification. In this classification, the system does not have any flow factors and the specimens (or objects) would just be immersed into biofilm environments. (2) Flow systems In these systems, the flow factor would be added to the LBRs. (3) Quasi-natural systems. In these systems, many practical and complicated factors are involved to adjust the practical (natural) and laboratory scale experiments.

5.2 Static Systems In these systems, flow factors are not usually considered and specimens or objects are just immersed and exposed in the concrete examples provided for Static Systems. (1) Colony Biofilm [3, 4] Colonies are grown over agar. Basic biofilm characteristics are maintained like the structured environment and chemical ingredients. The advantages are reproducibility and simplicity. This method would be used for antibiotic susceptibility assessment, morphology observations (essentially upon polysaccharide production). (2) Microtiter plate [5, 6] This method has often been used for biofilm evaluations. Bacteria attach to well surfaces. However, from the viewpoint of materials science and engineering, the material specimens should be immersed in wells. The characteristics are simple to run, amenable to high throughput screening, and suited to molecular genetic tests. This method has been used for evaluation of biofilm formation of strains, biofilm antibiotic tolerance and resistance, and efficiency of antibiofilm/antimicrobial products. (3) Biofilm Ring Test [7] This method utilizes mobility measurements of magnetic beads that are mixed with bacterial suspensions in microplates. Biofilm formation could be evaluated rapidly. The adhesion process could be evaluated. This method could avoid washing and staining procedures. Particularly, the early stage of biofilm formation could be evaluated. (4) Calgary Biofilm Device [8] This method is based on a 96-well microtiter plate assay. It includes a lid with 96 pegs on which biofilms develop. Pegs can be removed individually without opening the system, which avoids contamination. Consistent shear force across all pegs is possible. This facility is already available commercially. Using this method, one could evaluate biofilm’s antibiotic resistance and tolerance, efficiency

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Fig. 5.4 A static method by the authors

of antibiofilm/antimicrobioal products, etc. Biofilm development with time could be evaluated too. Usually, we (the authors) use a plastic plate of 12-wells to quickly investigate the anti-biofouling for biofilm formation [9–13]. Figure 5.4 is a schematic illustration for the process. It shows the following: (a) the appearance observed from the perpendicular direction and (b) the cross sectional view from the horizontal direction. Each well is filled with a liquid culture containing bacteria. For example, LB culture with E. coli, or HI culture with S. epidermidis. Conventionally, the wells themselves have been made for such a procedure. Specimens to be investigated are also added to each well. After immersion for a certain period of time, the specimens are removed from the wells and either stained or evaluated using Raman spectroscopy. Sadly, the flow is missing for this method. However, it allows us to effectively investigate the formation and growth processes depending upon the combination of substrate materials and bacteria used. Our method basically belongs to the Microtiter plate method.

5.3 Flow Systems In these systems, an important factor for the biofilm formation/growth process, “Flow”, is added to the test. Actually, researchers have pointed out that this concept would be very important for biofilm formation/growth [14–24]. From this viewpoint, we can say the liquid flow would lead to biofilm formation on materials. Pipes with liquids flowing inside of them are typical examples. In those cases, the factors derived from the liquid flow would be shear stress, nutrient concentration, exposure times, Reynold’s number, etc. Originally, the biofilm affected by the flow form on materials in natural environments. The flow condition differs from case to case. For example,

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the pipe changes for some liquids, depending on what kind of applications the pipe has. Sample applications include tap water in houses, cooling water in factories, and catheters for our bodies. The problems involved to control biofilms would define the conditions required in the laboratories. It means that the flow type laboratory tests should imitate the natural conditions as much as possible. A variety of flow systems have already been proposed. Several examples include the Drip Flow biofilm reactors and the Kadouri System. The Drip Flow biofilm reactors are designed for the study of biofilms grown under low shear conditions. This type of reactor has four parallel test channels that can hold one standard microscope slide each. It is good for general biofilm studies, high biomass production, and medical material evaluations [25]. The Kadouri system is a low-flow system that is a bridge between static assays and continuous-flow systems. We (the authors) developed some methods to investigate materials’ resistance and characteristics for biofilm formation/growth. They are shown in Fig. 5.5, schematically. Figure 5.5a shows the closed loop type of LBR. In our case, a single species for example, E. coli [26] or S. epidermidis [27] were inserted into the system and biofilms were formed on specimens placed in the upper column (Fig. 5.6). Such a flow mimicked the urine system and biofilm formations to produce urinary track stones, etc. The first generation of a LBR produced by us was a circulating LBR. This LBR had a three-necked beaker, which was composed of the culture part for E. coli and the column part where the sample was inserted. When the two parts were connected using a tube, at least 6 joint parts were created. The more binding sites there were, the more likely contamination would occur. Actually, bacterial solutions were weakly leaked at some joint parts. In particular, during the autoclaving, fatigue accumulated in the parts and leakage was likely to occur. Such a difficulty to handle the LBR might be an important and serious problem (for this type of reactor). The loop type LBR allows the experiment to be performed in an environment closer to the real environment, since the liquid culture including bacteria is constantly flowing. As described above, the possibility of leakage at some joint parts might be problematic. However, this problem is also related to the skill of the person who actually performs the experiment. Therefore, we expect that the possibility of

Fig. 5.5 The three flow types of LBRs developed by the authors: a closed loop type, b rotation type 1 (specimens are fixed), c rotation type 2 (specimens are rotated on the disk)

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Fig. 5.6 A concrete example of a flow type of LBR developed by the authors and the application to a series of experiments

problems will decrease with the improvement of the experimenters’ skills. The loop type of LBR may be a little difficult to handle as already described. However, the sample area has less oxygen to make the environment harsh for bacteria. From the viewpoint, this apparatus might simulate the real environment for some bacteria. Actually, the result of crystal violet staining for specimens by this method showed that biofilm formation was most noticeable. To improve the difficulty of handling the loop-type of LBR, we developed two kinds of rotation type LBRs. Figure 5.5b shows the rotation type where the specimens are fixed and the jig is rotated to produce circulation flow. Figure 5.7 shows a schematic example of practical applications. On the other hand, Fig. 5.5c shows another rotation type of LBR where the specimens on a plate are rotated to produce rotation flow by themselves. The specimens are fixed to the plate by a special organic compound at this point (Fig. 5.8). By using

Fig. 5.7 A concrete example of a flow type of LBR (rotation type 1) developed by the authors and the application to a series of experiments

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Fig. 5.8 A concrete example of a flow type LBR (rotation type 2) developed by the authors and the application to a series of experiments

these LBRs, we could avoid the leakage problems completely. In these processes, the part where the bacterial culture is going on is quite the same with that where biofilm formation would occur. Therefore, the biofilm formation would occur easier and faster. And in addition, the possibility of contamination would be lower and the sterilization process would be easier. However, it would be hard for us to measure the flow rate in this case. It must be analyzed theoretically, by using the analysis of fluid dynamics. Apart from those LBRs mentioned above, we have developed a flow chip apparatus independently. Figure 5.9 shows one of them. Using this type of LBR, we confirmed that the biofilm formation/growth process occurred as usual and the acceleration process was available. This allows us to investigate the biofilm formation/growth process more rapidly and more easily. Therefore, we expect further development of this LBR type from practical viewpoints.

5.4 Quasi-natural Systems (Ex Vivo Systems) As can be seen in the previous two sections, various laboratory biofilm reactors have been developed to mimic natural biofilm formation and growth processes in laboratories. Obviously, such laboratory biofilm reactors would sacrifice some factors in the case of natural biofilms. This is because we need to speed up the process and simplify the factors in order to easily and accurately analyze the phenomena. However, we might miss the essential factors in such a procedure. Therefore, we need the natural exposure where specimens are immersed in natural environments. As already described repeatedly, the natural exposure always requires a longer amount of time. The two extremes often need intermittent measuring systems to obtain an accurate solution. For example, we could modify the bacterial biota by single species, or by multiple ones. Maybe, we might need to add some nutrients to the system. Also we

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Fig. 5.9 A flow chip LBR developed by the authors: a process/principle, b appearance

could modify the structure of the LBRs, to make them more like the natural environment. This type of modification is called a Microcosms system by Lebeaux et al. [2]. This system closely mimics in situ conditions and includes more environmental factors. References with medical examples related to this system are provided [28–34]. References relating to medical ex vivo conditions are also provided [35–40]. The authors developed modifications to establish intermediate LBRs between those for natural environments and laboratory environments [41–50]. Figure 5.10 shows a typical one. In this LBR, the bottom tank is filled with tap water, sea water, etc. The pump electrically moves liquid from the tank upward into tubes and rotates it into the system. The specimens are placed in the upper column where the center line should be parallel with the surfaces of the specimens. In the column, liquid moves from the left side to the right side. The liquid flows out from the system once and flows down on the punching metal where the concavo convex profile (of the surface) is made to increase the contact area between the plate and liquid. Then the liquid flows down into the tank. At the punching metal plate environmental biota are able to mix with the liquid. This process is repeated for a certain period of time. For example, it could be for a couple of days, a couple of weeks or a couple of months. Using this method, biofilms form with good reproducibility, since the wild species lead to biofilm formation. They survive by living in oligotrophic environments. Therefore, such bacteria generally need to form biofilms for their survival. On the other hand,

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Fig. 5.10 Flow type LBR to form biofilms by environmental biota. a A display of the setup, b a diagram of the process

the cultured bacteria sometimes lose their capability to form biofilms. This is because they do not always need nutrition from biofilms. Their nutrition is often provided in the laboratories. In this case, we are not able to get information about the primary bacteria that form the biofilms. Also the results might not relate well to general cases, because the LBR would be placed in a certain (specific) type of environment. Usually experiments are carried out for screening and comparing the antibiofilm characteristics among specimens. In most cases, the order among specimens would be the same and not depend on the environments. By comparing the results with exposure tests in natural environments, we could estimate the life cycles of materials’ specimens. These results have already been applied to research and development activities for some companies. ROTATION DISK REACTOR Researchers Barry and McGrath of Clarkson University designed and used a Rotation Disk Reactor for carrying out a preliminary investigation to assess the biofilm forming ability of a variety of items used in hospitals. Biofilms are groups of microorganisms that adhere to each other on a surface, especially as a result of exposure to water and bacteria. They can pose health risks to humans as they grow in hospital settings that include medical supplies and devices. Most bacteria that cause health problems exist in biofilms [51–53]. They are embedded within a self-produced EPS (extracellular polymeric substance) that makes them resistant to antibiotics. Therefore, from a medical point of view, the researchers used their unique reactor to test six medical items: a nebulizer (made of polyvinyl chloride), a syringe (made of polypropylene), a pipette (made of polyethylene), cannula tubing (made of polyvinyl chloride), a suction catheter (made of polyvinyl chloride with added plasticizers), and an external male catheter (made of latex, natural rubber).

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Fig. 5.11 Rotation Disk Reactor with attached samples and dip tank

Their reactor was created to simulate a hospital setting (where items are exposed to moisture and air), and to accelerate biofilm growth without resort to artificial acceleration substrates [54]. This setup included a HDPE dip tank with 30 L of deionized/distilled water and a rotating wheel that could hold multiple samples. A motor was used to turn the wheel at 3 RPM. This process allowed the samples to become wetted and exposed to microorganisms in the ambient air during each revolution of the wheel. Figure 5.11 is a display of the Rotation Disk Reactor used for two preliminary studies [55]. After several weeks of testing, the latex catheter sample was wetted and had a surface coating that looked white and slimy. See Fig. 5.12 [55]. The other samples were still hydrophobic. Based on this work, the best material for use in hospitals is polyethylene (which was represented by a pipette for this study). Upon further testing, SEM images for the latex catheter samples displayed rod shaped bacteria. The rapid growth of bacteria on the latex catheter samples concerned the researchers. As a result, they decided to carry out a second preliminary study to find methods of inhibiting bacteria/biofilm growth on latex, especially since it is used in catheters and other medical devices. They searched the literature, but were not able to find information that directly related to their work. However, they did find published articles about metal ions and the use of electric fields exerting antibacterial effects [56–60]. Therefore, the investigators launched their new project to assess the influence of metals and voltage for the growth of biofilm on latex samples.

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Fig. 5.12 The latex catheter sample displays a white film on its surface after several weeks of testing

Once again the unique Rotation Disk Reactor and setup conditions (exposure to ambient air and wetted with deionized/distilled water) were used for three trials of the experiment. This time all of the samples were latex from an external male catheter. For each trial, the test items included a control sample of latex, latex with silver, latex with stainless steel, latex with brass, latex with both stainless steel and brass, and latex with paired electrodes and applied voltage (the use of a 9.5 V battery). The samples were repeatedly immersed in water followed by an air exposure period. The continuous motion of the wheel ensured that microorganisms in the ambient air had an opportunity to mix with the water in the dip tank. Since medical items in hospitals are often exposed to both moisture and air, this system provided a comparable environment in an accelerated manner. The water level in the dip tank was kept constant and the temperature, conductivity, and pH were continually monitored. Throughout the test, the samples were observed for wettability and biofilm formation. Observations were made using the unaided eye, magnifiers, cameras, a scanning electron microscope, etc. The results (as viewed from SEM images) showed that when compared to the control sample (where more than 90% of the surface was covered with rod shaped bacteria), the presence of metals (brass, stainless steel, and silver) was generally effective in preventing bacterial growth. Figure 5.13 is a SEM of the aged control sample. The use of voltage essentially eliminated the appearance of rod shaped

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Fig. 5.13 The SEM image (at 2,500X) of the aged control latex sample displays many rod shaped bacteria

bacteria in some of the samples. It can be concluded that the presence of metals significantly reduced bacterial growth on latex and the application of voltage was able to essentially eliminate bacteria, providing appropriate electrode combinations were used. This work (using the unique Rotation Disk Reactor) and the encouraging results are important from a medical point of view because the goal was to obtain ways to prohibit the growth of bacteria/biofilm on latex that is used in catheters and other clinical devices. The results showed brass to be the most effective single metal with less than 1% of the latex surface covered with rod shaped bacteria. Also the use of voltage essentially eliminated the appearance of rod shaped bacteria when like metals were used for the electrodes such as brass with brass [61]. It should also be mentioned that since bacteria are often encased in biofilm, a reduction in the amount of bacteria will also reduce the amount of biofilm. EX VIVO systems were originally defined as experimental ones using samples taken out of organisms. Usually experiments are performed inside organisms to demonstrate the biofilm phenomena. For example, mice and other animals are often used to investigate the biofilm formation/growth process and to check its effect on the inside of the animals’ bodies. This type of experiment is called in vivo and will be explained later. On the other hand, ex vivo systems are done outside of an

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organism’s body. Therefore, these concepts (in vivo and ex vivo) should be used for investigations about the relationship of biofilms to infection. From this viewpoint, ex vivo systems for biofilms should be experiments where the animal or human body environments are replaced with various kinds of cells, e.g. mammalian cells. Ex vivo experiments have the following two merits. One of them is to carry out practical experiments and evaluations that would be very dangerous or impossible within living organisms. The other merit is not to violate the ethical code, for example, animal protection rights, etc. Some biofilm research activities have been devoted to clarify the correlation between biofilm formation and mammalian cells [62–69]. In many cases mammalian cells were used for biofilm research in order to clarify the competitive correlation between the cells’ attachment to materials and the bacterial growth on them. It could shed light on a process by which materials inserted into human bodies (as biomaterials or implants) might control the recurrence of infections by bacterial growth with biofilm formation. Such a materials science approach could successfully lead to the further clarification of BAI (Biofilm Associated Infection or Biomaterials Associated Infection).

5.5 In Vivo Systems In vivo systems utilize the natural environments to form biofilms and to clearly fix the effects on materials. We would like to differentiate the biofilm related phenomena into two main types. One of them corresponds to the extracorporeal case. In such a case, materials are exposed to the same or similar natural environments to check the antibiofilm characteristics. Figure 5.14 shows a research project about the corrosion characteristics of materials in a certain type of atmospheric environment. Metallic specimens are fixed to a special jig and exposed to a natural environment. Their environment included some rain and some sunshine. However, the environment was very natural and depended on the weather. The angle of the jig could be changed, due to the conditions required for the investigation of the materials’ performance. Figure 5.15 shows the experimental outline for the immersion of metallic specimens into marine environments. In this experiment, we wanted to investigate biofilm formation and growth on immersed specimens in a certain actual marine environment. We used the floating dock in a marina of Ise Bay near Suzuka, Japan. The specimens were suspended from the docks. Strictly speaking, the technical term, “in vivo” is used for biological and medical applications. Therefore, the two extracorporeal cases may sound a bit strange. However, these are also in vitro cases, since the experiments were carried out in a natural environment. The cases applied to medical problems are clearly in vivo. To clarify the infectious diseases and the correlation among diseases, immune systems, and biofilms, researchers and engineers generally use two kinds of organisms as shown in Fig. 5.16.

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Fig. 5.14 A practical example of corrosion tests for metallic specimens in an atmospheric environment. The authors wanted to investigate the correlation between corrosion phenomena and biofilm formation/growth characteristics in this experiment. Specimens were fixed to jigs and exposed to a certain natural environment

Fig. 5.15 Metallic specimens were suspended from the floating dock in a marine environment to investigate biofilm formation/growth behavior in a natural environment

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Fig. 5.16 In vivo experiments for biofilms and their organisms

Using various kinds of organisms, researchers could analyze the correlation between organisms’ metabolism and biofilms. In most cases, the final target should be human beings’ reactions to biofilms. In the light of that, higher organisms should have reactions closer to those that humans would have with biofilms. Many in vivo experiments have been summarized by Lebeaux et al. [2]. Some references for non-mammalian in vivo studies (relating to biofilms) are provided [70–81]. Many references relating to mammalian in vivo studies are provided too [82–208].

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173. Lucke, M., Schmidmaier, G., Sadoni, S., Wildemann, B., Schiller, R., Haas, N. P., et al. (2003). Gentamicin coating of metallic implants reduces implant-related osteomyelitis in rats. Bone, 32, 521–531. 174. Lucke, M., Wildemann, B., Sadoni, S., Surke, C., Schiller, R., Stemberger, A., et al. (2005). Systemic versus local application of gentamicin in prophylaxis of implant-related osteomyelitis in a rat model. Bone, 36, 770–778. 175. Prabhakara, R., Harro, J. M., Leid, J. G., Keegan, A. D., Prior, M. L., & Shirtliff, M. E. (2011). Suppression of the inflammatory immune response prevents the development of chronic biofilm infection due to methicillin-resistant Staphylococcus aureus. Infection and Immunity, 79, 5010–5018. 176. Li, D., Gromov, K., Soballe, K., Puzas, J. E., O’Keefe, R. J., Awad, H., et al. (2008). Quantitative mouse model of implant-associated osteomyelitis and the kinetics of microbial growth, osteolysis, and humoral immunity. Journal of Orthopaedic Research, 26, 96–105. 177. Fitzgerald, R. H., Jr. (1983). Experimental osteomyelitis: description of a canine model and the role of depot administration of antibiotics in the prevention and treatment of sepsis. Journal of Bone and Joint Surgery. American Volume, 65, 371–380. 178. Petty, W., Spanier, S., Shuster, J. J., & Silverthorne, C. (1985). The influence of skeletal implants on incidence of infection. Experiments in a canine model. Journal of Bone and Joint Surgery. American Volume, 67, 1236–1244. 179. Philipov, J. P., Pascalev, M. D., Aminkov, B. Y., & Grosev, C. D. (1995). Changes in serum carboxyterminal telopeptide of type I collagen in an experimental model of canine osteomyelitis. Calcified Tissue International, 57, 152–154. 180. Williams, D. L., Haymond, B. S., Woodbury, K. L., Beck, J. P., Moore, D. E., Epperson, R. T., et al. (2012). Experimental model of biofilm implant- related osteomyelitis to test combination biomaterials using biofilms as initial inocula. Journal of Biomedical Materials Research Part A, 100, 1888–1900. 181. Schurman, D. J., Trindade, C., Hirshman, H. P., Moser, K., Kajiyama, G., & Stevens, P. (1978). Antibiotic-acrylic bone cement composites. Studies of gentamicin and Palacos. Journal of Bone and Joint Surgery. American Volume, 60, 978–984. 182. Blomgren, G., & Lindgren, U. (1981). Late hematogenous infection in total joint replacement: Studies of gentamicin and bone cement in the rabbit. Clinical Orthopaedics and Related Research, 244–248. 183. Southwood, R. T., Rice, J. L., McDonald, P. J., Hakendorf, P. H., & Rozenbilds, M. A. (1985). Infection in experimental hip arthroplasties. Journal of Bone and Joint Surgery. British Volume, 67, 229–231. 184. Bernthal, N. M., Stavrakis, A. I., Billi, F., Cho, J. S., Kremen, T. J., Simon, S. I., et al. (2010). A mouse model of post-arthroplasty Staphylococcus aureus joint infection to evaluate in vivo the efficacy of antimicrobial implant coatings. PLoS One, 5, e12580. 185. Berra, L., Curto, F., Li Bassi, G., Laquerriere, P., Baccarelli, A., & Kolobow, T. (2006). Antibacterial-coated tracheal tubes cleaned with the Mucus Shaver: A novel method to retain long-term bactericidal activity of coated tracheal tubes. Intensive Care Medicine, 32, 888–893. 186. Fernandez-Barat, L., Li Bassi, G., Ferrer, M., Bosch, A., Calvo, M., Vila, J., et al. (2012). Direct analysis of bacterial viability in endotracheal tube biofilm from a pig model of methicillin-resistant Staphylococcus aureus pneumonia following antimicrobial therapy. FEMS Immunology and Medical Microbiology, 65, 309–317. 187. Olson, M. E., Harmon, B. G., & Kollef, M. H. (2002). Silver-coated endotracheal tubes associated with reduced bacterial burden in the lungs of mechanically ventilated dogs. Chest, 121, 863–870. 188. Tollefson, D. F., Bandyk, D. F., Kaebnick, H. W., Seabrook, G. R., & Towne, J. B. (1987). Surface biofilm disruption. Enhanced recovery of microorganisms from vascular prostheses. Archives of Surgery, 122, 38–43. 189. Bergamini, T. M., Bandyk, D. F., Govostis, D., Kaebnick, H. W., & Towne, J. B. (1988). Infection of vascular prostheses caused by bacterial biofilms. Journal of Vascular Surgery, 7, 21–30.

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207. Rediske, A. M., Roeder, B. L., Nelson, J. L., Robison, R. L., Schaalje, G. B., Robison, R. A., et al. (2000). Pulsed ultrasound enhances the killing of Escherichia coli biofilms by aminoglycoside antibiotics in vivo. Antimicrobial Agents and Chemotherapy, 44, 771–772. 208. Engelsman, A. F., van Dam, G. M., van der Mei, H. C., Busscher, H. J., & Ploeg, R. J. (2010). In vivo evaluation of bacterial infection involving morphologically different surgical meshes. Annals of Surgery, 251, 133–137.

Chapter 6

Detection and Evaluation of Biofilms

Abstract This chapter describes a variety of ways for detecting and evaluating biofilms. The biological methods include staining, gene analysis, and proteomics. An instrumental analysis section is available too. It introduces many microscopes and discusses their uses for observing and analyzing biofilms.

6.1 Biological Methods 6.1.1 Staining Staining is an important method for detecting biofilms, especially from the industrial viewpoint. It is mainly classified into two categories: non-fluorescent and fluorescent types. The former usually utilizes the phenomena where some pigments adsorb to specific components of cells, such as cell membranes, organelles, etc. On the other hand, the latter is based on the color produced when certain proteins bind with pigments. Researchers and engineers usually observe the stained parts by using microscopes. Some staining processes are described below.

6.1.1.1

Crystal Violet

Crystal violet is usually used for Gram stain [1]. This method makes it possible for us to tell gram-negative from gram-positive bacteria. Figure 6.1 shows the general process for Gram staining. The method was invented by Hans Christian Gram in 1884. The first step is the staining process using crystal violet. After about 30 s of staining, the dye is removed (from the sample) by water. Bacteria display a purple color. During the second step, iodine is applied to the stained bacteria. This makes the staining of the cells firmly fixed. Thirdly, alcohol is applied to the purple bacteria for about 30 s and then removed by rinsing with water. At this stage, some bacteria lose

© Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_6

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Fig. 6.1 Usual process for gram staining

the stained purple color, while others keep it. The former is called gram-positive bacteria, and the latter is gram-negative bacteria. As already mentioned, the difference occurs due to the structural change of the cell walls. Then at the final stage, safranin is applied to the bacteria. After the final rinsing with water, the gram-negative bacteria would be invisible by microscopes, while the gram-positive bacteria could be observed clearly due to the deep purple color. The explanation described above corresponds to the original Gram stain test. Gram-positive bacteria have a thick peptidoglycan layer. Therefore, the bacteria are stained very strongly. On the other hand, the peptidoglycan layer for gram-negative bacteria is thin and the pigment can be removed easily. Due to the difference of stain characteristics, gram-positive bacteria show violet color clearly, while gram-negative bacteria show red. Crystal violet for biofilm detection does not have different targets to stain. The target which we aim to stain in this method is EPS (the extracellular polymeric substances). Figure 6.2 shows the chemical structure of crystal violet. As shown in this figure, crystal violet is composed of anionic and cationic parts. In the aqueous solution, it dissociates and the cations specifically bind with the anionic parts. As a result, the absorbed parts show violet color after washing with water. This means that crystal violet is basically absorbed to the negative parts of compounds based on the electrical force. EPS is composed of various organic compounds derived from organisms. It is classified into four main types: polysaccharides, proteins, nucleic acids and lipids. Since polysaccharides generally have strong polarity in some of their parts, crystal violet adsorbs to them very strongly and generally stains them with a violet color.

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Fig. 6.2 Chemical structure of crystal violet

However, crystal violet also adsorbs to the negatively charged parts of other EPS components to show violet color. Therefore, any organic components in EPS would be stained. Usually, researchers and engineers remove adsorbed crystal violet by dissolution into some appropriate solvents and the solvents’ colors are checked. Figure 6.3 shows an example of our work [2]. Biofilms were formed on small coupon-like specimens by an artificial method. Then they were moved into some appropriate wells and washed three times using sterilized water. Each well with a specimen was filled with 0.1% crystal violet solution for one hour or 30 min. The Fig. 6.3 Stained glass due to the existence of biofilms

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duration of time depended on the particular situation. After the immersion, specimens were removed from the wells and rinsed with sterilized water several times (three times in most cases). By using this process, the specifically absorbed crystal violet could be removed to a certain extent. As a result, specimens could show a violet color with the remaining absorbed crystal violet on the EPSs [3]. The absorbed crystal violet was dissolved and extracted with 1 ml ethanol for a certain amount of time (for example, 15 min.). The absorbance of the solution was measured at about 590 nm using spectrophotometers or plate readers. The extent of absorbance was also analyzed. On the other hand, the reflection or transmission of stained solid surfaces could be measured directly. Ogawa et al. carried out experiments that confirmed the stained specimens (by crystal violet) could quantitatively show the amount of biofilm formation [4]. Using a certain type of LBR (Laboratory Biofilm Reactor), Ogawa and her group artificially produced biofilms on some kinds of specimens in an accelerated way. Then the specimens were freeze dried by substituting the water with ethanol and t-butyl alcohol. They were immersed into 0.1% crystal violet solution for 30 min at 25 °C. The samples were washed with water to remove the nonspecifically absorbed crystal violet from the specimens. Scotch tape was attached to the stained surface for a certain period of time. Then the tape was removed. The crystal violet staining biofilms were transferred to the scotch tape. The tape with the stained biofilms was attached to glass slides and the color was measured by a color reader. The degree of staining corresponds to the amount of biofilm formation, which was evaluated by the color difference based on parameters, L*, a* and b*. An example is shown in Fig. 6.4. The degree of staining is displayed in a*-b* plane.

Fig. 6.4 An example for the evaluation of color by crystal violet staining

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Even though many staining methods have been investigated in biology, it seems like the staining for biofilms has been concentrated on crystal violet [5–9]. Actually, SIAA (a Japanese non-profit organization containing many Japanese companies in antibacterial industries) is now going to establish an international standard for biofilm evaluation based on crystal violet [10]. This decision could be attributed to the following reasons. (A) Crystal violet could attach to negative parts of huge molecules ranging from polysaccharides, proteins, nucleic acids and lipids. The pigment seems to be suitable to stain complicated compounds as a whole. On the other hand, organic polymers in biofilms are very complicated and composed of the four kinds mentioned above. Therefore, the complicated EPS would be hard to stain and analyze. For these reasons, crystal violet seems to be the best to evaluate biofilms at this point. (B) Being compared with the fluorescent pigments, the treatment is generally much better. (C) The cost would be lower. (D) The stained specimens could be analyzed intuitively. Also the detailed methods could be diversified and widened in various ways.

6.1.1.2

Other Possibilities

As already described, there are many pigments and staining methods available. Some pigments and methods have been applied to biofilm formation. However, it seems like crystal violet is the best so far. This section presents other possibilities with concrete examples. Bismarck Brown [11–16] is a historical pigment that turns acid mucins (proteins) to a yellow color. Some researchers used it for biofilm evaluation. Carmic acid is a red pigment known as cochineal extract. It is produced as an extract from some insects and can stain glycogen a red color. Therefore, it can stain the polysaccharide parts of EPS derived from Biofilm [17–19]. Coomassie Brilliant Blue (CBB) has been used as a pigment to stain proteins a blue color. Therefore, this pigment might be able to stain the protein components of EPS [20–22]. Alcian Blue [23–26] is a basic pigment belonging to the phthalocyanine group. It binds to carboxyl groups and sulfates, and stains sticky polysaccharides. However, the binding and staining depend on pH values. Silver staining [27–29] uses the characteristics of silver ions, which tend to bind to proteins favorably. After silver ion binds to proteins, it would be reduced by formalin or citric acid, and the stained parts would turn black. Benzidine dye [30–32] could stain amyloids (a kind of protein) a red color. The staining called Congo red can be used to detect proteins [33–36]. Methyl green is a cationic or positively charged stain as well as crystal violet [37–39]. It could basically stain DNA green and RNA red. Therefore, nucleic acids in biofilms might be detected properly. Sudan III [40–42] is basically lysochrome diazo dye (1-(w-(Phenyldiazenyl) phenyl) azonaphthalen-2-ol) and could stain fat components. Therefore, it might be

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Fig. 6.5 Non-fluorescent types of pigments and their possibilities to stain components of biofilm

used to stain lipid components. All of these pigments could stain one or more components of EPS, but not all of them. Therefore, we recommend that they be used in combination with crystal violet, especially when one needs to investigate a certain component for some specific purposes. The concept is shown in Fig. 6.5 in the case of non-fluorescent pigments. On the other hand, there are many fluorescent types of pigments. DAPI (4’6diamidino-2-phenylindole) could bind to adenine-thymine strongly and emit fluorescence. Therefore, this could be used to detect nucleic acid components in biofilms. Ethidium bromide (EthBr) is a fluorescent type of pigment that strongly bonds to DNA. Since this is considered as a carcinogen, it may not be used in the future. Hoechst stains [43–45] were originally developed by the company (Hoechst). Therefore, this term is a trademark. The blue fluorescent dye is called Bis-benzimide and is used to stain DNA. However, it is potentially considered a carcinogen. In any case, we can presume that many fluorescent types of pigments could be used to tell nucleic acid components from EPS. The third group of pigments is composed of chemicals to stain bacteria themselves. Crystal violet was originally used with iodine, alcohol, and safranin to stain bacteria, as described earlier. Such pigments have been investigated a lot. For example, Eosin could stain the cell cytoplasm red [46–48]. Malachite green is a basic organic pigment that is also called Aniline green, Basic green 4, Victoria green B, etc. It could stain bacteria themselves and is sometimes used as a substitute for crystal violet. These pigments could stain bacteria as a whole and the staining shows the existence of bacteria in biofilms. As explained earlier, biofilms are composed of water, bacteria, and EPS. Therefore, the staining indirectly shows the biofilms just like the pigments staining EPS. However, there are some problems for the pigments staining bacteria when used as an indicator for biofilms. A few are mentioned below. (A) Even though bacteria exist as dormant in biofilms, some might change their structures. Both dormant and modified bacteria could be stained just like planktonic bacteria. (B) In biofilms, live and dead bacteria exist simultaneously. Both could be stained by those pigments.

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(C) A lot of bacteria might be broken down under bacteriolytic reactions by bacterial phages. Could the pigments estimate and take the phenomenon into consideration? Therefore, the staining of bacteria themselves should be carried out in parallel with other types of pigments.

6.1.2 Gene Analysis Gene analysis is defined as the process where the genome of a certain organism would be analyzed holistically. Basically, the base arrangement is decided from segments of DNA constituting genomes. Therefore, the analysis can be defined also as a genome analysis. Genome is the technical term corresponding to the entire genetic information contained in DNA. However, the process should include the analysis for RNA and proteins. The general process is shown in Fig. 6.6 as a flow chart [49–58]. First of all, EPS (derived from biofilms) has to be collected from biofilms formed on sessile matter. Then polysaccharides, proteins, and lipids are removed from the collected EPS and genome DNA is extracted (process 1 in Fig. 6.6). Since the extracted DNA is still too large to be analyzed, it is cut into fragments by using various enzymes. The fragments produced through this process bind to vector DNA such as plasmid. (Plasmid is a small DNA molecule within a cell that is physically separated from a chromosomal DNA and can replicate independently. It is often found in bacteria.) Therefore, the number increases in E.coli. The process is called cloning (process 2). Then the sequencer is used to decide the base arrangement and fluorescent materials are attached to the target DNA (process 3). The process is composed of two steps. First of all, DNA polymerase, an enzyme relating to the DNA synthesis, lengthens the DNA chains. Then the base arrangement is determined by DNA electrophoresis, which is used to decide the nucleotide sequence of a piece of DNA. Therefore, the base arrangement used for process 3 is for fragments that are shorter than the original DNA genome. In order to rebuild the original arrangement, overlapped material should be removed and the data needs to be edited and combined (process 4). After the rebuilt base arrangement is confirmed, (process 5), then the estimation of the Open Reading Frame (ORF) is carried out with the aid of computers (process 6). ORF is the part of the base arrangement that is available for transaction and translation. Finally, the base arrangement is translated (with the meaning of functions) based on the past research data and a huge database through comparisons and investigations (process 7). The importance of gene analysis for biofilm research is very clear. Biofilms are basically complicated systems and each biofilm in nature is composed of many kinds of bacteria and other matter including EPS. To simplify and idealize the system for research activities, we often use a single species of bacteria in order to analyze the biofilm phenomena. However, such a research method often makes it impossible for

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Fig. 6.6 General processes for gene analysis

us to grasp the real images of biofilms, since biofilms generally contain many kinds of unknown bacteria. Most bacteria in biofilms are not only diversified, but also VBNC bacteria. VBNC stands for “viable but non-culturable bacteria”. Therefore, the pure culture is impossible in most cases and does not have any serious meanings. This is because one cannot determine the primary bacteria in natural biofilms. (Primary bacteria are defined as bacteria which first produce the biofilms.) As a conclusion, we can say the identification of bacteria within biofilms is impossible or unavailable to solve any problems relating to biofilms. However, the identification of bacteria contained in the target biofilms is still very important. To solve such a contradictory problem, gene analysis can prove to be effective. We introduce an applied case by Kanematsu and his collaborators [59, 60]. During the summer, some specimens were immersed in Ise Bay, Japan for a certain period of time. The sea around Japan’s islands, particularly in Ise Bay (Fig. 6.7), has two seasons in regards to biofilm research. In the first half of the year (from April to

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Fig. 6.7 The map of Japan’s Islands

September), biofilm formation is accelerated, while it is slow from October to March due to the temperature changes. Therefore, our immersion test in a practical marine environment was done during the active season for biofilm formation. The specimens for the immersion were carbon steel (JIS SS400), stainless steel (AISI 304, tin plated steel, and pure copper. The size of these specimens was 50 × 50 mm. They were immersed into the Bay Area for a week. After the immersion, the specimens were taken out of the sea. Even though the immersion time was only one week, the specimens’ surfaces changed due to biofouling. It looked like biofilms formed on the specimens’ surfaces. We scraped all of the surface products (composed mostly of biofilms) together and extracted DNA. From this DNA, 16S-rRNA, a special region of the bacterial gene, was amplified by PCR (Polymerase Chain Reaction). Each 16S-rRNA was incorporated into vectors to form a 16S-rRNA library. Also the base arrangement was determined by a sequencer for DNA. Using such a measurement system, the researchers obtained a variety of interesting results which have never been found before. For example, the bacterial flora differed from substrate to substrate as shown in Fig. 6.8. It suggests that some unknown reactions might have occurred between biofilms (or bacteria) and the substrate. As shown in Fig. 6.8, bacterial flora changed, depending on the kind of substrates. For example, bacteria belonging to Fimicutes occupied the flora on carbon steel. Likewise, those belonging to γ-Proteobacteria were on tin plated steel. On the other hand, diversified bacteria were found on stainless steel. However, the flora changed with time, as shown in Fig. 6.9. In this case, flora changed on the tin plated specimen. However, the same phenomena would occur on most substrates. This could be attributed to the biofilm formation. In the biofilm, growth behaviors changed dramatically from that in the planktonic state [59]. As described in this practical case, gene analysis is a powerful tool to learn a lot about biofilms and the phenomena relating to them such as a material’s degradation [60–62].

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(1) Stainless steel

(2) Tin-plated carbon steel

Fig. 6.8 Bacterial flora on stainless steels and tin-plated carbon steels

Fig. 6.9 The change of bacterial flora on tin-plated carbon steels with time

6.1.3 Proteomics Proteomics (according to a broad definition) was originally a research field about the entire discipline of proteins and proteomes [63–65]. A proteome is a set of proteins produced in a system, organism, or some biological context. Conventional studies focus on the purification of proteins and the use of mass spectrometry for the analysis. Proteins are the essence of organisms. For example, our human bodies are composed of 60% water and 20% proteins. Proteins help support lives of organisms in various ways. EPS, an important biofilm component, is actually composed of four types of carbon compounds at the early stage of biofilm formation. The four types include polysaccharides, proteins, nucleic acids, and lipids [66–69]. Polysaccharides are the essence of biofilms, since they are produced by quorum sensing. However, quorum sensing is a kind of signal deduction process. In this case, a signal deduction is carried out by the moving and cascade reactions of proteins. Proteins are also used for adhesion [70]. They are used for adhesion of bacteria onto sessile matter.

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From all of these results, one can see that it is very important to analyze proteins for biofilm evaluation and analyses. In this chapter, we have focused on some analytical methods for proteins to obtain information about biofilms.

6.1.3.1

Proteins and Biofilms

Biofilms are made by bacterial activities. Bacteria move toward materials’ surfaces driven by chemotaxis. This is because carbon compounds (which serve as nutrition) adsorb to materials’ surfaces [66, 67]. Generally, bacteria scatter lots of proteins for various reasons. One could be to send signals, so bacteria would take a certain action to survive. Another reason could be for adhesion, by which bacteria could attach firmly to materials’ surfaces. On the other hand, bacteria contain lots of proteins in their cells. On the surfaces of cells or within bacterial cells, many protein cascading, chain reactions occur. When bacteriolysis occurs in biofilms (which is quite often.), proteins in bacterial cells end up being added to the biofilm components. Based on the above reasons, proteins are sometimes more important than polysaccharides.

6.1.3.2

Vibrational Spectroscopy

As explained in Sects. 6.2.9 and 6.2.10, vibrational spectroscopy such as Infrared spectroscopy (FTIR) and Raman spectroscopy could be used to determine what exists in biofilms [71]. Infrared spectroscopy uses infrared light as incident light that is absorbed or transmitted through a sample. These values are plotted on the y-axis, while the frequency or wavelength values are plotted on the x- axis. The reflection type of analysis is also available. It uses the surface correlation between the incident infrared light and a sample’s surface molecules. Raman spectroscopy uses the Raman Scattering phenomenon. By comparing the incident light (usually laser light) and scattered light, the energy shift could be observed. The shift corresponds to the energy loss for the sample’s own vibration energy. The sample can be fixed using the energy loss values. Proteins have complicated structures. Two methods (presented below) are available to analyze them. (1) The theoretical calculations and predictions. (2) The fingerprint method by comparing experimental data to an existing database. As for the first approach, one has to precisely predict the structure of a molecule in advance. For the second approach, the data volume in the databases is sometimes small and the compounds corresponding to Raman shifts are missing. Also overlapping of Raman shifts for various compounds often occurs. In such a case, it is hard for us to fix the compounds by only using vibrational, spectroscopic data. Basically,

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Raman spectroscopy and FTIR are related in a complementary way, as shown in Sects. 6.2.9 and 6.2.10. However, other analytical methods should be added to both vibrational methods in order to precisely identify proteins. Such methods will be described.

6.1.3.3

Uv-Vis

UV-VIS stands for Ultraviolet-Visible Spectroscopy. If biofilms would form on transparent or semi-transparent substrates, UV-VIS might be used for biofilm evaluations. UV-VIS usually uses the light of wavelengths from 200 to 1,500 nm. When the light irradiates the specimens’ surfaces, the electron transitions of specimens’ components would occur in various ways. The π-π* transition, n-π* transition, d-d transition, metal-ligands charge transfer, etc. can be mentioned for examples. The energy of incident light would be consumed for the transitions or transfer phenomena. Also the light at a certain wavelength would be absorbed by the specimen. Protein-containing solutions would generally show the absorption of ultraviolet light at 200–215 nm and at 280 nm. The former corresponds to that of peptides and the latter to the side chains of polyaromatic amines. The former peaks might be overlapped with others derived from different matter. The latter would not be overlapped with other peaks so often. Therefore, the latter absorption could be used for biofilm evaluations.

6.1.3.4

Mass Spectrometry

Mass spectrometry is basically the analytical method to determine the ratio of mass to charge for certain samples. These samples are generally composed of atoms, molecules, and ions. Specimens could be ionized by several methods. Then an ionic flow would be generated. The ionic flow would bend through magnetic fields, since the perpendicular force (Lorentz Force) would be applied to the ionic flow. The force would depend on mass numbers. The ionic flow that reaches the detector would have special ions of a certain mass. The results would generally be displayed as a spectrum with the mass to charge ratio (m/z) at the horizontal axis and the ion abundance (%)/ionic strength at the vertical axis. Mass spectrometry is classified according to ionization methods [72, 73]. On the other hand, it could be classified into some categories based on the separation of ionic flows. One method utilizes an electromagnetic field to separate the ionic flows. This method is generally called a double-focusing mass spectrometer, since the first electric field sector accelerates ionic flow and the second magnetic sector bends the ionic flow under the balance of the Lorenz Force and a centrifugal force. As for quadruple mass spectrometry, four electrodes are used, through which ionized flows would move. Since the high frequency voltage would be applied to the electrodes in addition to the direct voltage, a certain ion would be allowed to pass through the electrodes and reach the detector.

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TOF (time-of-flight) Mass Spectrometry utilizes the phenomenon that the flight speed differs from ion to ion, when ions are accelerated by the applied voltage. The primary ion would be irradiated to the specimens’ surfaces and the emitted secondary ion flies at a certain speed, depending on the ratio of mass number to charge. In this method, mass separation would be carried out due to the flight time difference of the secondary ions. This method makes it possible to analyze organic and inorganic materials. It generally has a high degree of precision and the image analyses would be available too. For Ion Trap Mass Spectrometry, ions would be kept in the trap section of the apparatus. However, when the applied high frequency voltage would increase, the ion would be released, depending on the ratio of mass number to charge. Therefore, mass separation would be available for this method. As compared to other methods, this one has a high advantage in the light of qualitative analysis. In addition to these methods, other special methods have been developed. One of them is the FT-ICR method which stands for Fourier-Transform Ion Cyclotron Resonance. In this method, ions are introduced in the cell under the combination of a static electric field and a magnetic one. When the high frequency voltage is applied, the ions begin a circular motion and the cycle for the ion cyclotron motion would be used for the mass separation based on Fourier Transformation. The cost for the introduction of the apparatus is generally very high. The Acceleration Mass Spectrometry (AMS) is another special example. It utilizes accelerators and the mass separation would be carried out based on the energy loss in the process within the accelerators. Inevitably, the apparatus might be extensive. This method has been used for carbon dating quite often. As already described in other chapters repeatedly, EPS and its analysis are very important for biofilm evaluation, because EPS causes industrial problems in various ways. However, EPS is a complicated, complex system, since it is composed of versatile molecules produced by various bacteria. The metabolic substances produced by bacterial life activities are generally called metabolites. An investigation to analyze metabolites is called metabolomics. For this method, one would analyze the metabolites by cataloging them holistically. Since biofilm components form a complicated system, this method would be very useful to analyze them. Mass spectrometry is essential for such an analysis.

6.1.3.5

NMR (Nuclear Magnetic Resonance Method)

From the1940s to the 1950s, the analytical method using nuclear magnetic resonance phenomenon was established independently by two groups. One group was led by Dr. Purcell at Harvard University and the other was led by Dr. Bloch at Stanford University [74, 75]. When a strong magnetic field is applied to samples, their nuclear magnetic moments (nuclear spins) align in the direction opposite to that of the magnetic field. Since the nuclear spin against the magnetic field has higher energy than that aligned in the direction of the magnetic field, the spin with lower energy increases to the

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energy state (with the higher energy). The phenomenon is called nuclear magnetic resonance. When the electromagnetic waves at radio frequencies are applied to matter, the energy of the electromagnetic wave is absorbed by the sample. When the excited state returns to the original one, the energy for the difference between the two states is emitted. The change would be transferred to electric currents. In such a way, the sample could be analyzed and identified from the NMR spectrum. This method is also very useful for analyzing metabolites produced in biofilms. As well as mass spectrometry, NMR will be more important for investigating biofilms in the future [76–78].

6.2 Instrumental Analysis 6.2.1 Optical Microscopes There are many types of microscopes. From the viewpoint of materials science and engineering, optical microscopes are the simplest to use for observing materials’ surfaces. As for optical microscopes, the probe is the visible light and the signal is the reflected or penetrating light. Since the probe is the visible light, the resolution is determined by the following Eqs. (6.1 and 6.2) [79]. d = λ/(2AN )

(6.1)

AN = n x sin(θ)

(6.2)

The symbols used for the equations are as follows. d: distance between the objects, λ: wavelength of the incident light. AN : the numerical aperture of the microscope and defined as Eq. 6.2. n: the refractive index of the medium. Usually, the value in air is the unit. θ: one-half of the largest angle at which the lens can accept light reflected from the object. Using these equations, one can estimate the resolution for an optical microscope with visible light. One can also see how the resolution could be improved by using several options. Optical microscopes are mainly classified into biological microscopes (BM), metallographic microscopy, and stereomicroscopes. For the BM system, the light is placed under the specimen (object) and irradiates the specimen from underneath to penetrate the specimen. Then the penetrated light is observed. Therefore, it is a transmission type of optical microscope. Metallographic microscopy irradiates light onto the specimen from above and the reflected light is observed. Therefore, it is called a reflecting microscope. This

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technique is used to look at the physical structure and components of metals and alloys. The stereomicroscope has its own independent optical systems for both eyes and the difference is used to give 3D appearances. The difference is called the disparity. Therefore, the sample can be observed as it really is. In this system, the lenses move up and down at various relative positions between the object and the lenses. At various heights, the images are taken at the focal points. For each height, the proper relative positions are obtained. These proper points are combined to draw 3D images of the object. Kanematsu et al. used an optical microscope with the system of Depth from Defocus (DFD) to observe biofilms formed on various kinds of materials [80–83]. Figure 6.10 shows some examples. Since biofilms generally have concavo—convex surface profiles, this method allowed the researchers to get three dimensional images of biofilms. Even though this method might be relatively simple and easy to get 3D images of biofilms, the resolution is low when compared to apparatus that uses electron beams, etc. However, no special preparations are needed in order to observe the specimens. As for biofilm observation, the optical microscope is the simplest one to use. If one acquires proficiency in the skill of observing, then the optical microscope would be the best choice. For example, Fig. 6.11 shows an image of biofilms formed on a material. It was obtained when the authors used optical spectroscopy attached to Raman spectroscopy. The white spots correspond to biofilms. In this case, the spots were confirmed as biofilms by Raman spectroscopy. Without the combination of both types of spectroscopy, it might be impossible to confirm the existence of the biofilms. It is suggested that the optical method be used for initial screening, and that other methods be used for further investigations.

Fig. 6.10 Biofilms observed by an optical microscope with the DFD function

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Fig. 6.11 Biofilms on PVC (polyvinyl chloride) sheets observed by an optical microscope

6.2.2 Fluorescence Microscopes Fluorescence microscopy takes advantage of fluorescence, the ability of substances to absorb short wavelengths of light (ultraviolet) and give off light at a longer wavelength (visible). Some organisms are able to fluoresce naturally under ultraviolet light. Specimens that do not fluoresce naturally are stained with a fluorescent dye (a fluorochrome). When stained specimens are examined under a fluorescence microscope with an ultraviolet or near-ultraviolet light source, they look like luminescent, bright objects against a dark background. Fluorochromes have a special attraction for different microorganisms. For example, the fluorchrome fluorescein isothiocyanate causes Bacillus anthracis (the anthrax causing agent) to appear apple green [84]. The main use for fluorescence microscopy is for diagnostics, especially the fluorescent-antibody (FA) technique or immunofluorescence [85]. This method can be used to detect bacteria or other pathogenic microorganisms within cells, tissues, etc. Figure 6.12 shows fluorescence microscopy images [86]. A few ways are possible for staining biofilm components in fluorescence microscopy. One is to use fluorescence in situ hybridization (FISH) probes, where the staining takes place on dead microorganisms [87]. This method allows researchers to detect the presence and location of selected microorganisms in the biofilm. Hybridization probes are single-stranded fragments of DNA or RNA designed to attach to complementary nucleotide sequences in a sample. The probes have fluorescent markers,

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Fig. 6.12 Fluorescence microscopy images of sun flares pathology in a blood cell showing the affected areas in red

so their location can be determined in the sample. In order for in situ hybridization to take place, the probe needs to enter the microbial cell and hybridize with the nucleotide it was designed for. Another way to stain biofilm components in fluorescent microscopy is by using fluorescent proteins (FPs), which were initially extracted from a particular type of jellyfish [88]. Today a variety of FPs is being used. When stimulated with light from a particular wavelength, a FP emits fluorescent light of a specific color (such as green, yellow, or red), which depends on the structure of the protein. The advantages for using this technique are that FPs can be genetically encoded and tracked in living cells. These proteins are used to enhance the visibility of bacterial cells in biofilms.

6.2.3 Confocal Laser Scanning Microscope Confocal microscopy is a technique in light microscopy used to reconstruct threedimensional images. It is good for obtaining high resolution optical images with depth selectivity. See Fig. 6.13, which shows a stack of confocal microscope images [89]. For the confocal microscopy method, specimens are stained with fluorochromes so that they will emit or return light. However, only one plane of a small region of the specimen is illuminated with a short-wavelength (blue) light. The returned light is passed through a pinhole aperture aligned with the illuminated region. Successive regions and planes are illuminated until the complete specimen has been scanned.

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Fig. 6.13 A stack of confocal microscope images showing the distribution of actin filaments throughout a cell

This method provides clear two-dimensional images. Confocal microscopy can be used in combination with computers in order to obtain three-dimensional images. It makes the three-dimensional structures of biofilms more visible. The confocal laser scanning microscope (CLSM) works by passing a laser beam through the aperture of a light source which is then focused (by an objective lens) on a small area of the sample’s surface [90]. Samples can be scanned point by point or multiple points at once. This allows an image to be built up pixel by pixel by collecting the emitted photons from the fluorophores in the sample. The CLSM has been used to investigate biofilm growth on paints in the field [91]. For this project, painted polycarbonate slides were glued on polycarbonate panels mounted on a test raft in a harbor. The painted slides enabled CLSM analyses of undisturbed biofilms. The results showed that extracellular polymeric substances were the dominating biofilm component. The CLSM has been used to study biofilm structure, composition, and metabolism in various microorganisms, allowing in-depth analysis of biological structures without killing or damaging them. It was used to analyze S. epidermidis biofilms exposed to the antibiotics farnesol and vancomycin [92]. The interesting results showed that even though these antibiotics were not able to effectively kill biofilm bacteria, the

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biofilm biomass was strongly reduced. Also farnesol and vancomycin seemed to damage the cell membrane as determined by the live/dead staining. Stoodley and others designed a flow cell that could be mounted onto the stage of a confocal laser scanning microscope that allowed the biofilm to be observed in situ under flow conditions [93]. Also the elastic properties of biofilms could be clearly seen. Real-time observations of bacterial adhesion, etc. provide a better understanding of industrial problems and situations.

6.2.4 Scanning Electron Microscope (SEM)—Energy-Dispersive X-Ray (EDX) Spectroscopy The scanning electron microscope (SEM) is great for examining biofilm with high resolution. It is suitable for following the adhesion stage and biofilm formation. SEM is a well-established basic method to observe the morphology of bacteria attached to materials’ surfaces, the morphology of the material’s surface, and the relationship between them [94]. It is also used to determine the effect of surface roughness on adhesion. Kouider and others used the SEM to determine the effect of stainless steel surface roughness on Stapylococcus aureus adhesion [95]. The results showed that the adhesion level was found to largely depend on the substrate roughness, with a maximum at Ra = 0.025 μm and a minimum at Ra = 0.8 μm (where Ra stands for roughness average). Due to the use of a very narrow electron beam, SEM micrographs have a large depth of field that provides a characteristic three-dimensional appearance that is useful for understanding the surface structure of a sample. See Fig. 6.14, an image of pollen grains taken on an SEM [96]. The scanning electron microscope produces three-dimensional images of a sample by scanning the surface with a focused beam of electrons. This beam referred to as the primary electron beam, is produced by an electron gun. Its electrons pass through electromagnetic lenses and then travel over the surface of the specimen. As a result, the primary electron beam knocks electrons out of the surface of the specimen. These secondary electrons are transmitted to an electron collector and amplified in order to produce an image (scanning electron micrograph) on the viewing screen. This image displays the topography of a sample’s surface. Objects are generally magnified 1000–10,000 X. In addition, characteristic xrays are emitted when the primary electron beam removes an inner shell electron from the sample. This is because a higher energy electron fills the shell and releases energy. The energy of these characteristic x-rays can be measured by energy-dispersive x-ray spectroscopy (EDX) and used to identify and measure the abundance and distribution of elements in the sample [97]. SEM samples need to be prepared to withstand vacuum conditions and the high energy beam of electrons. They have to be small enough to fit on the specimen stage too. The samples are usually rigidly mounted to a specimen holder or stub using a conductive adhesive. Metal objects require little sample preparation. However,

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Fig. 6.14 Above is an image of pollen grains taken on an SEM. It shows the characteristic depth of field of SEM micrographs

non-conductive materials are usually coated with an ultrathin coating of electrically conducting material that is deposited on the sample either by low-vacuum sputtercoating or by high vacuum evaporation. In regards to biological specimens, they need to be completely dry for SEM testing. Therefore, the SEM has been most successful for imaging samples immune to dehydration such as mineral deposits. Since the extracellular polymers (EPS) of biofilms are highly hydrated (with more than 90% water), they are greatly affected by the dehydration, sample preparation process. This process degrades the initially slimy polymers, so they appear in SEM images as a network of random strands. This is not desirable. However, it provides images of biofilm microorganisms without the polymeric matrix (which if intact would have obstructed the view of these microorganisms). It should be mentioned that a SEM technique is available for overcoming obstacles such as the dehydration of samples. This method (for monitoring biofilm) enables the imaging of hydrated specimens [98]. It is a modified, low-vacuum scanning electron microscopy technique called environmental scanning electron microscopy (ESEM).

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6.2.5 Transmission Electron Microscope (TEM) The transmission electron microscope (TEM) uses a finely focused beam of electrons. The beam is from an electron gun and passes through an ultrathin section of a specially prepared specimen, which is usually placed on a copper mesh grid. An electromagnetic condenser lens is used to focus the beam on a small section of the sample. It directs the beam of electrons in a straight line to illuminate the specimen. After passing through the specimen, the beam goes through an electromagnetic objective lens, which magnifies the image. Finally an image (called a transmission electron micrograph) appears on a fluorescent screen or photographic plate. Dark and light areas are displayed, depending upon the number of electrons absorbed by different parts of the specimen. A TEM is shown in Fig. 6.15 [99] and a TEM image of a cluster of polio virus is provided in Fig. 6.16 [100]. This instrument can generally magnify objects 10,000–100,000 times. Many microscopic specimens are very thin, so there is a weak contrast between the background and their ultra structures. Therefore, the contrast can be improved by using a dye that absorbs electrons to make a darker image in the stained area. Salts of heavy metals are used as stains. Also a microbe can be seen by the method called shadow casting [101]. Here, a heavy metal such as gold is sprayed (at about a 45° angle) to hit the microbe from only one side. The uncoated side of the specimen leaves a shadow. This provides a three-dimensional effect with a general idea of the size and shape of the specimen. The TEM provides high resolution and is valuable for examining different layers of specimens. On the other hand, a disadvantage for using the TEM is the sample preparation process [102]. Since samples must be dehydrated and viewed under a high vacuum, they are killed and distorted and sometimes appear to have additional structures. The additional structures that result from the preparation process are called artifacts. TEM takes place in a high vacuum, with pressures around 10−5 mm Hg. Since biological specimens are usually highly hydrated and need to be dehydrated before being placed in a vacuum chamber, this process may change the structure of the specimen. This problem needs to be addressed, especially if conclusions are to be made based on the shape of the specimen under the microscope. One attempt is to fix the samples by using cross-linking chemicals that help maintain the sample’s original structure after the water has been removed. In work carried out by Hunter and Beveridge, high pressure freeze-substitution and transmission electron microscopy have been used for high resolution imaging of the natural structure of gram-negative biofilm [103]. They found different domains of healthy and lysed cells (cells breaking down) randomly dispersed within a single biofilm. Also particulate matter was suspended within a network of fibers. It seemed to be an integral part of the exopolymeric substance (EPS). Their work shows a structural complexity to biofilms at both the cellular and extracellular matrix levels. It should be mentioned that all aspects of Cryo TEM ultimately rely on the initial freezing step because the cells need to be rapidly frozen so that they are vitrified in amorphous ice [104]. This is to prevent the formation of ice crystals that could damage

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Fig. 6.15 A TEM is displayed. (Key: 1. High tension cable. 2. Electron emitter. 3. Stepper motors for centering the electron beam. 4. Condenser. 5. Aperture controls. 6. Specimen holder. 7. Objective lens. 8. Projector lens. 9. Optical binoculars. 10. Fluorescent screen 11. Vacuum pump leads. 12. Goniometer. 13. Vacuum and magnification control. 14. Focusing control)

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Fig. 6.16 A TEM image of a cluster of polio virus is provided. The polio virus is 30 nm in diameter

cellular structure. The samples then undergo freeze substitution while slowly being brought back to room temperature. The amorphous ice is substituted with organic solvents to preserve the samples’ components. The samples can be embedded in plastic resin (for example) and further processed by a variety of imaging techniques. The TEM shows enhanced preservation of a sample’s components. As mentioned above, Cryo TEM is a technique applied on samples cooled to cryogenic temperatures and embedded in the environment of vitreous water. In 2017, the Nobel Prize in Chemistry was awarded to Jacques Dubochet, Joachim Frank and Richard Henderson for developing Cryo-Electron Microscopy for high-resolution structure determination of biomolecules in solution [105]. Figure 6.17 shows a (Cryo-TEM) image of an intact ARMAN (Archaeal Richmond Mine Acidophilic Nanoorganism) cell from an Iron Mountain biofilm [106, 107]. An ARMAN cell is a novel ultra- small archaeon (a microorganism that resembles bacteria) present in acid mine drainage biofilms.

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Fig. 6.17 A (cryo-TEM) image is displayed for an intact ARMAN cell from an iron mountain biofilm. Image width is 576 nm

6.2.6 Atomic Force Microscope (AFM) Atomic force microscopy is a type of scanning probe microscopy with very high resolution on the order of fractions of a nanometer. It was invented by IBM scientists in 1982. Gerd Binnig and Heinrich Rohrer developed the scanning tunneling microscope (STM), a precursor to the AFM, for which they jointly earned half of the Nobel Prize for Physics in 1986 [108]. The other half of that Nobel Prize went to Ernst Ruska for his work with electron optics and for the design of the first electron microscope. The first AFM was commercially available in 1989. It is one of the foremost tools for imaging, measuring, and manipulating matter at the nanoscale. Specimens do not require special preparation for this type of microscopy. The AFM consists of a flexible cantilever with a sharp tip (probe) at its end that is used to scan the surface of a specimen positioned on a piezoelectric scanner. When the tip is brought close to a sample’s surface, forces between the tip and the sample lead to a deflection of the cantilever (which is typically silicon or silicon nitride). Depending on the situation, forces that are measured in AFM include mechanical contact force, van der Waals forces, capillary forces, chemical bonding, etc. For example, mechanical force is used to measure mechanical properties of the sample like Young’s modulus (which is a measure of stiffness or adhesion strength). The

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AFM is also used for imaging. The reaction of the probe to the forces that the sample imposes on it can be used to form a three-dimensional image of a sample’s surface at a high resolution. In addition, the forces between the tip and sample can be used to change the properties of the sample in a controlled way. Examples include atomic manipulation and local stimulation of cells. Figure 6.18 shows a simple block diagram of an atomic force microscope [109]. The AFM can be operated in three modes: contact, non contact, and the tapping mode. In the contact mode, the cantilever is scanned across the mapped surface of a specimen. For the non contact operation, the cantilever oscillates as it is scanned across the specimen’s surface. The tapping mode is usually used in ambient conditions or in liquids. It is similar to the non contact mode, except that the amplitude of oscillations is bigger and the cantilever movement is affected by different forces. The non contact mode can provide resolution on the atomic scale and is good for measuring soft biological samples. Tapping provides resolution on the scale of individual molecules. The AFM can be used to image any types of surfaces including those of biological specimens. Figure 6.19 shows an AFM image of glass [110]. Figure 6.20

Fig. 6.18 Displayed is a simple block diagram of an atomic force microscope. It uses beam deflection detection. As the cantilever is displaced by its interaction with the surface, the reflection of the laser beam will be displaced on the surface of the photodiode

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Fig. 6.19 This is an AFM topographical scan of a glass surface. It portrays the roughness of the material. The image space is (x, y, z) = (20 um × 20 um × 420 nm)

Fig. 6.20 Single polymer chains (0.4 nm thick) recorded in the AFM tapping mode under aqueous media with different pH

displays single polymer chains [111]. The AFM is used to image individual microorganisms deposited on a variety of conductive and non conductive surfaces in the initial stages of biofilm formation before the entire surface is covered with EPS. Researchers used the AFM to examine biofilm surfaces under ambient conditions and observed distinctive features. They analyzed the height images obtained on biofilms

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of Straphylococcus aureus, Nocardia brasiliensis and Pseudomonas aeruginosa [112]. In another project, scientists were interested in cohesive strength, which affects the balance between growth and detachment of biofilm. Therefore, its quantification is important to understand, predict, and model biofilm development. They developed a novel AFM method to reproducibly measure (in situ) the cohesive energy levels of moist—1- day biofilms (grown from an undefined culture taken from activated sludge). Their results showed that cohesive energy increased with biofilm depth, from 0.10 ± 0.07 to 2.05 ± 0.62 nJ/μm3 . This observation was reproducible, with four different biofilms showing the same behavior [113].

6.2.7 Ultraviolet-Visible (UV-VIS) Spectroscopy Ultraviolet-Visible (UV-Vis) spectroscopy uses light from the UV and visible regions of the spectrum. The visible region includes wavelengths (about 400–700 nm) associated with colors that we can see, while the ultraviolet light has shorter wavelengths (starting at about 10 nm up to about 400 nm) that can cause sunburns and cancer. Figure 6.21 is a display of the electromagnetic spectrum [114]. The instrument used in UV-Vis spectroscopy is called a UV/Vis spectrophotometer. Figure 6.22 is a picture of this equipment [115]. Its basic components are a light source, a holder for the sample, a diffraction grating in a monochromator or a prism to separate wavelengths

Fig. 6.21 A display of the electromagnetic spectrum is provided

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Fig. 6.22 A Beckman DU640 UV/Vis spectrophotometer is displayed

of light, and a detector. This type of spectroscopy is used in analytical chemistry for the quantitative determination of samples such as transition metal ions, highly conjugated organic compounds and biological macromolecules. Samples are often placed in a transparent cell (curvette), which is usually rectangular in shape with an internal width of 1 cm. (This width becomes the length in the Beer-Lambert law.) In a typical experiment, wavelengths (about 200–800 nm) are sent through the sample (analyte) that is to be analyzed. Samples are most often liquids, but can also include gases and solids. The intensity of light that passes through a sample is I and the intensity of the light before it passes through the sample is I o . Using these values, the percentage transmittance (%T ) is determined as follows. %T = I /Io times 100 The absorbance A is based on the transmittance, which is incorporated into the Beer-Lambert law [116]. This law is most often used in a quantitative way to determine the concentrations of absorbing species in solution. It is written below along with the meaning of each symbol. A = log10 (Io / I ) = ε c L A is the measured absorbance, I o is the intensity of the incident light at a given wavelength, I is the intensity of the light that passes through the sample, L is the path length through the sample, c is the concentration of the absorbing species, and

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ε is a constant known as the molar attenuation coefficient (which is sometimes called molar absorptivity or extinction coefficient). A UV-Vis spectrum shows absorbance (y-axis) plotted against wavelength (xaxis). The wavelengths of absorption peaks can be correlated with the types of bonds in a given molecule and are valuable in determining the functional groups within a molecule. An ultraviolet absorption spectrometer system was developed (for the Lydon B. Johnson Space Center in Houston, Texas) for continuous real-time monitoring to detect the growth of biofilms. Continuous monitoring could provide early warning of potentially harmful buildups of bacteria. This ultraviolet absorption spectrometer system is based on a miniature UV-Vis spectrometer with a fiber-optic input and a CCD array detector. The system measures UV absorption spectra of biofilms produced on the inner surface of a quartz window of a flow cell. Tests using this system showed that biofilms produced characteristic absorption spectral bands at wavelengths from 230 to 400 nm [117]. Absorption bands obtained from a single strain of Pseudomonas aeruginosa differed from the absorption bands of biofilms grown from a mixed bacterial population from untreated urban river water.

6.2.8 White Light Interferometer A white light interferometer is an instrument used for a non-contact optical method. It measures the surface height of 3-D structures with surface profiles between tens of nanometers and a few centimeters. This method relies on visible-wavelength light (white light). The interferometer uses the wave superposition principle to combine waves in order to extract special information from them. NOTE: If two waves (that are in phase) combine, then the result is constructive interference. However, the combining of two out of phase waves produces destructive interference. See Fig. 6.23 for a schematic of a white-light interferometer [118]. A CCD image sensor (like one for digital photography) is located at the point where the two images are superimposed. A broad band (white light) source is used to illuminate the test and reference surfaces. A condenser lens collimates the light from the light source and a beam splitter separates the light into reference and measurement beams. The reference beam is reflected by a reference mirror and the measurement beam is reflected or scattered from the test surface. The beam splitter relays the returning beams to the CCD image sensor. As a result, an interference pattern of the test surface topography is formed. This pattern (surface topography) is spatially sampled by individual CCD pixels. Keep in mind that interference occurs for white light when the path lengths of the measurement beam and the reference beam are nearly the same. In order to view microscopic structures, it is necessary to combine an interferometer with the optics of a microscope, such as a standard optical microscope. White light interferometry (WLI) is not usually used to image bacterial biofilms that are immersed in water. This is because there is not enough refractive index

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Fig. 6.23 A schematic layout of a white-light interferometer

contrast to induce reflection from the biofilm’s interface. The properties of hydrated biofilms make it hard to characterize them in situ, especially in a non-destructive way. A method was developed to measure and monitor the topology and thickness of live biofilms using a WLI microscope. The researchers used a microfluidic system to produce a reflective interface on the surface of biofilms [119]. This technique allowed for the measurements of roughness, surface area, etc. Experimental results showed that an increase in biofilm thickness appeared after an increase in surface roughness. The group also developed a flow cell to observe changes in biofilm structure as a result of changes in environmental conditions like flow velocity and the amount of available nutrients. Bacterial biofilms are being monitored with a microfluidic flow chip designed for imaging with white light interferometry. The novel imaging chip enables the use of WLI to capture high resolution three-dimensional profile images of biofilm growth over time [120]. For example, biofilm formation was detected as early as three hours after inoculation of the flow cell with a live bacterial culture (Pseudomonas fluorescens). Researchers are using this setup in order to better understand biofilm development and properties such as anti-microbial resistance. In the future, this work may be used to combat common diseases associated with biofilm formation.

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6.2.9 Fourier-Transform Infrared Spectroscopy (FTIR) FTIR stands for “Fourier-transform infrared spectroscopy” which utilizes infrared light for a probe [121–123]. When the infrared light enters matter (solid, liquid or gas), the matter partially absorbs the light. This technique shines a beam (containing many frequencies of light at once) and measures how much of that beam is absorbed by the sample. Next the beam is modified to contain a different combination of frequencies to give a second data point. This process is repeated many times. Then a computer is used to analyze the data to determine what the absorption is at each wavelength. The energy of light could be used for vibration of binding molecules or their rotation. Therefore, the absorbed light’s energy (wavelength) must correspond to the sample’s own molecular binding. The phenomena could be utilized to analyze the sample’s chemical structures. Since the amount of absorbed light would generally be proportional to the concentration and volume of the samples, they could be utilized to quantitatively measure the samples. The schematic relationship between the wavelength and chemical structure (s) is shown in Fig. 6.24. In the figure, the horizontal axis corresponds to wavelength. The wavelength is defined as the inverse of wavelength and the mathematical relation is shown in Eq. 6.3. v =

Fig. 6.24 Infrared light and chemical structure

1 λ

(6.3)

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Here, ν’ is the wavelength, and λ is the wavelength. As for the wavelength, the following relation exists between wavelength ν’, frequency ν and the speed of light, c. v =

v c

(6.4)

The region of energy required for the stretching of a corresponding chemical structure is shown. It is usually the region from 1500 to 500 cm−1 . This is because the complete set of absorptions for each compound can be found in this region. Using such a chart, one can predict the chemical structures from the experimental data. As described above, FTIR usually uses the absorption of penetrated infrared light. However, the reflection type of FTIR has been developed [124–126]. Figure 6.25 shows this principle. The infrared light is directed onto a dense crystal with a high refractive index. The internal reflectance creates an evanescent wave that goes beyond the surface of the crystal into the sample held in contact with the crystal. If the high reflective index crystal is closely attached to the sample, then the total of the repeated reflective beams would have information about the chemical structures in the vicinity of the surface. In this method, the prism is attached and has firm contact with the specimens. The total amount of reflected light by the specimen’s surface is collected and measured. The molecules in the vicinity of the specimen’s surface absorb the infrared light through reflection. Therefore, we get information about the molecules on a material’s surface. Using this method, we could measure and analyze organic matter existing on materials’ surfaces. We want to analyze biofilms formed on materials’ surfaces, so this method would be very powerful and effective [127–129].

Fig. 6.25 Schematic illustration for the principle of FTIR-ATR (attenuated total reflectance)

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6.2.10 Raman Spectroscopy As described in the previous section (Sect. 6.2.9), light entering matter might be reflected, absorbed, or refracted. In addition, it can be scattered (Fig. 6.26). Scattering usually occurs when the incident light changes its orientation due to an interaction with matter. It is classified into Rayleigh scattering (elastic scattering) and Raman scattering (non-elastic scattering), as shown in Fig. 6.27 [130]. In the case of Rayleigh scattering, the wavelength for the incident light would be the same as that of the scattering one. However, for the case of Raman scattering, the wavelength of incident light would be different from that of the scattering one. Strictly speaking, two Raman scatterings could be observed. One has a higher wavelength than the original one, and the other has a lower wavelength. The former is called Stokes scattering and the latter is Anti-Stokes scattering. Raman scattering occurs through the interaction of incident light and matter, just like FTIR described in the previous section. The phenomenon was first found by Raman and Krishnan [131–133]. They observed the scattering by using mercury as a light source and a photospectroscope. Since the laser was used as a light source, the output power increased [132, 134]. Therefore, the Ruby Laser (694.3 nm) [135] and Helium-Neon Laser (632.8 nm) [136, 137] were developed. In the mean time, monochrometers were developed and stray light was eliminated. In addition, the use of photomultiplier tubes also increased the sensitivity of Raman spectroscopy. In Fig. 6.26 The interaction between light and matter

Fig. 6.27 Rayleigh scattering and Raman scattering

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Fig. 6.28 Schematic illustration of the apparatus for Raman spectroscopy

1970, when Raman spectroscopy was combined with microscopes, research activities increased tremendously [138]. Figure 6.28 shows the general setup for Raman spectroscopy. It detects and analyzes organic compounds and other covalent inorganic compounds on solid surfaces. The general advantages for this spectroscopy are provided. Advantages of Raman Spectroscopy #1: No special preparations are needed for the samples. They can be measured directly without any pre-treatments. #2: Organic and inorganic specimens can be measured. Also specimens that are solutions, powders, crystals, or gasses, can be measured. #3: Structures of matter derived from organisms can be analyzed. #4: If microscopy is combined with it, then the local analysis would be available. #5: Without any special optical elements or detectors, the measurement at lower frequencies (particularly those lower than 400 cm−1 ) would be available. #6: The non-destructive depth analyses would be available. On the other hand, Raman spectroscopy has disadvantages too. Some are mentioned. #1: Fluorescence emitted by laser irradiation often lies in the way to analyze the results correctly. #2: The specimens might be damaged by laser irradiation. #3: The database for Raman spectroscopy is not so abundant, as compared with that for infrared spectroscopy. #4: Generally, the cost of Raman spectroscopic apparatus is higher than that for infrared spectroscopy. As for disadvantages, the fluorescence phenomenon caused by laser radiation is the worst obstacle for Raman spectroscopy. To avoid this phenomenon, it is best to use different laser sources having different wavelengths. Generally, Raman peaks

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remain at the same wavelength regardless of the laser sources. Also the fluorescence problem would soon disappear by using different wavelength laser sources. If the specimens are very delicate and vulnerable to laser irradiation, the output power of the laser should be decreased for the measurement. As already described, Raman spectroscopy and Infrared spectroscopy utilize the common chemical phenomena relating to chemical covalent bonds. Therefore, the peaks derived from the same specimens could be found at the same or very similar wavelengths. However, there are some differences, which could be attributed to the different mechanisms of the two methods. Infrared spectroscopy (particularly FTIR) measures the transition between molecular vibrational energy when the infrared light is irradiating specimens. The transition is related to a resonance situation of electric dipoles. In the case of infrared light irradiation, the symmetric vibration generally would not absorb infrared light energy. Due to this reason, infrared spectroscopy has an advantage to analyze the structure for samples having asymmetric vibrational bonding. On the other hand, Raman spectroscopy is basically a two-photon inelastic light scattering phenomenon. When the energy of the incident photon is larger than the vibrational quantum energy of the sample, then a part of the energy for the incident photon is lost to give the sample molecular vibrations. Therefore, Raman scattering does not have any relation to resonance situations. Even in the symmetric vibration of molecular bonds, Raman shifts could be observed. Based on the differences of the two methods, we can say that Raman spectroscopy is suitable for symmetric molecules, while FTIR is good for asymmetric molecules. We can compare the two methods according to the following points. (A) The size of specimens For Raman spectroscopy, the minimum size of specimens could be around 1 μm, while for FTIR it is about 10 μm. (B) Specimens in glass containers Both the incident and scattered light of Raman spectroscopy can penetrate transparent glass. Therefore, the contents of the glass container could be measured for Raman spectroscopy. On the other hand, infrared light would be absorbed through glass. It is basically impossible to measure the contents of a glass container using FTIR. (C) Aquatic solution Raman spectroscopy is basically not affected by water, while middle infrared light is absorbed into water molecules. Therefore, Raman spectroscopy is suitable for measurements in water. On the other hand, FTIR is not suitable for such a measurement. (D) Database for finger print analyses The finger print method is where the experimental data is compared to databases in order to determine and analyze the structure of samples. As for this matter, FTIR has abundant data and the database is already mature. On the other hand, the database for Raman spectrums is not so abundant.

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(E) The cost of apparatus. The cost for Raman spectroscopy is generally higher than that for FTIR. (F) The damage to specimens (objectives). Since modern Raman spectroscopy generally uses laser light, the energy is very high and might damage the specimens to some extent. On the other hand, infrared spectroscopy does not generally use a high energy source. Therefore, the damage of specimens from FTIR is generally minor. Overall, the authors recommend Raman spectroscopy over FTIR. However, FTIR is important and both methods should be used to obtain the best analytical conclusions.

6.2.11 Nuclear Magnetic Resonance (NMR) Nuclear magnetic resonance (NMR) spectroscopy is a powerful tool for organic chemistry. It is a non invasive technique and provides information about the number of distinct types of hydrogen and carbon atoms in a given molecule and each atom’s environment. This type of imaging (often called MRI) is available in medical facilities to analyze soft tissue such as in the heart or brain. NMR uses radiation from the radio frequency (RF) region of the spectrum (radio waves). NMR spectrometers usually work in the narrow region between 108 and 109 hertz (Hz). They also use a superconducting magnet, which subjects the sample to a very high external magnetic field. Figure 6.29 is a photo of a NMR spectrometer [139]. For a typical NMR experiment, a sample is placed in the hollow bore of a superconducting magnet and irradiated with RF radiation that is emitted from the NMR probe. The sample is placed in a strong magnetic field because no RF radiation can be absorbed in the absence of such a field and no NMR spectrum can be recorded. This is because some atomic nuclei have nuclear spin. The nucleus of a hydrogen atom (example: a proton) has a spin of + ½ or – ½. Nuclear spin generates a small magnetic field that simulates a small bar magnet with north and south poles. A nucleus can be studied by this method if it has an odd number of protons, an odd number of neutrons, or both. A portion of the RF radiation absorbed by the sample at each frequency is reemitted at the same frequency. Each of these re-emitted frequencies (signals) corresponds to a frequency that has been absorbed by the sample. The amplitude of each signal is proportional to the amount of radiation that was originally absorbed. This data is converted into a NMR spectrum that is a plot of signal amplitude (y-axis) against chemical shift (x-axis). (Chemical shift is a measure of the extent to which a signal’s frequency differs from that of a reference compound.) Nuclear magnetic resonance imaging (NMRI) is used to measure flow velocity in biofilms. It delivers profiles of flow velocity for relatively low flow velocities like a few centimeters per second [140]. These profiles provide valuable information about

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Fig. 6.29 This is a photo of a Bruker 700 MHz NMR spectrometer

the performance of biofilm systems under controlled conditions in laboratory biofilm reactors. This technique is valuable for fundamental biofilm research. However, its use with real-life biofilm reactors is limited. For example, the reactor would need to fit into the core of the instrument. Researchers have used pulsed-field gradient nuclear magnetic resonance (PFGNMR) for measuring water diffusion in live biofilms [141]. The reactors used for this application are much smaller than those used for measuring flow velocity. In order to measure rates of diffusion of water using NMR, the sample is subjected to a series of pulsed magnetic field gradients that encode the displacement of each hydrogen atom into its signal. The displacement information is preserved in the magnitude of the NMR signal, which represents the vector sum of all spins within the volume of measurement. The goals of this work were to measure the effective diffusion coefficient for water in live biofilms, monitor how the effective diffusion

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coefficient changes over time under growth conditions and to correlate the effective diffusion coefficient with depth in the biofilm. They used Shewanella oneidensis MR1 biofilms for experiments with pulsed-field gradient nuclear magnetic resonance. This allowed them to measure (in situ) two-dimensional effective diffusion coefficient maps within the biofilms. They used this information to determine surface-averaged relative effective diffusion coefficient (Drs ) profiles. The investigators found that these profiles decreased from the top of the biofilm to the bottom. They also found that they changed with biofilm age.

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79. Lewandowski, Z., & Beyenal, H. (2013). Fundamentals of biofilm research (1466559608). CRC Press. 80. Vikramaditya, B., & Nelson, B. J. (1997). Visually guided microassembly using optical microscopes and active vision techniques. In Proceedings of the 1997 IEEE International Conference on Robotics and Automation (pp. 3172–3177). New Mexico: Albuquerque. 81. Levoy, M., Ng, R., Adams, A., Footer, M., & Horowitz, M. (2006). Light field microscopy. ACM Transactions on Graphics (TOG), 25, 924–934. 82. Nagahara, H., Kuthirummal, S., Zhou, C., & Nayar, S. K. (2011). Flexible depth of field photography. IEEE Transactions on Pattern Analysis and Machine Intelligence, 33(1). 83. Noguchi, M., & Nayar, S. K. (1994, October). Microscopic shape from focus using active illumination. In Proceedings of 12th International Conference on Pattern Recognitio (Vol. 1, pp. 147–152). IEEE. 84. Hermanson, G. T. (2013). Bioconjugate techniques (3rd ed.). 85. Goding, J. W. (1996). Immunofluorescence. In: Monoclonal antibodies (3rd ed., pp. 352–399). 86. File: Bloodcell sun flares pathology.jpeg. Date: October 27, 2005. This work is in the public domain. https://commons.wikimedia.org/wiki/File:Bloodcell_sun_flares_pathology.jpeg. 87. Morrison, L. E., Ramakrishnan, R., & Wilber, K. (2002). Labeling fluorescence in situ hybridization probes for genomic targets. In Methods in molecular biology. https://doi.org/ 10.1385/1-59259-300-3:41. 88. Crivat, G., & Taraska, J. (2012, January 1). Imaging proteins inside cells with fluorescent tags. Trends in Biotechnology, 30(1), 8–16. 89. Vindin, H. File: STD Depth Coded Stack Slices through Cells.png. Date: June 30, 2014. License: Creative Commons Attribution-Share Alike, 4.0 International https://commons. wikimedia.org/wiki/File:STD_Depth_Coded_Stack_Slices_through_Cells.png. 90. Pawley, J. B. (Ed.). (2006). Handbook of biological confocal microscopy (3rd ed.). Berlin: Springer. 91. Kamjunke, Norbert, Spohn, Uwe, Futing, Manfred, Wagner, Georg, Scharf, Eva-Maria, Sandrock, Stefan, et al. (2012). Use of confocal laser scanning microscopy for biofilm investigation on paints under field conditions. International Biodeterioration & Biodegradation, 69, 17–22. https://doi.org/10.1016/j.ibiod.2011.11.015. 92. Cerca, N., Gomes, F., Pereira, S., Teixeira, P., & Oliveira, R. (2012). Confocal laser scanning microscopy analysis of S. epidermidis biofilms exposed to farnesol, vancomycin, and rifampicin. BMC Research Notes, 5, 244. https://doi.org/10.1186/1756-0500-5-244. 93. Hall-Stoodley, L., & Stoodley, P. (2009, June 1). Evolving concepts in biofilm infections. Cellular Microbiology,11(7). https://doi.org/10.1111/j.1462-5822.2009.01323.x. 94. Peters, G., Locci, R., & Pulverer, G. (1982). Adherence and growth of coagulase- negative staphylococci on surfaces of intravenous catheters. Journal of Infectious Diseases, 146(4), 479–482. 95. Kouider, N., Hamadi, F., Mallouki, B., Bengoram, J., Mabrouki, M., Zekraoui, M., et al. (2010). Effect of stainless steel surface roughness on Staphylococcus aureus adhesion. International Journal of Pure and Applied Science, 4(1), 17. 96. File: Misc pollen. jpg. Date: December 9, 2004. This work is in the public domain. https:// commons.wikimedia.org/wiki/File:Misc_pollen.jpg. 97. Modgi, S., McQuaid, M. E., & Englezos, P. (2006). SEM/EDX analysis of Z-direction distribution of mineral content in paper along the cross direction. Pulp and Paper Canada, 48–51. https://www.pulpandpapercanada.com/paptac/PDFs/May06/paperanalysis.pdf. 98. Stokes, D. J. (2001). Characterization of soft condensed matter and delicate materials using environmental scanning electron microscopy (ESEM). Advanced Engineering Materials, 3(3), 126–130. 99. Ricce. File: Simens numeri.jpg. Date: August 19, 2008. This work is in the public domain. https://commons.wikimedia.org/wiki/File:Simens_numeri.jpg. 100. Murphy, F., & Whitfield, S. File: Polio EM PHIL 1875 lores.png. Date: May 27, 2006. The image (a work of the U.S. federal government) is in the public domain. https://commons. wikimedia.org/wiki/File:Polio_EM_PHIL_1875_lores.PNG.

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Chapter 7

Standardization—Current and Future

Abstract To improve and develop industrial products and processes, a standard is always needed. Nowadays, not only for academic activities, but also for industrial ones, various standards are developed beyond borders among nations. Therefore, the international standards are also needed. In light of that, we introduce various industrial standards relating to biofilms. Since the concept of biofilms is relatively new, there are not many existing standards for them. However, we already have some standards in advanced countries. In this chapter, we describe some related national standards for the United States, the European Union (EU), and Japan, as well as the International Organization for Standardization (ISO).

When industrial problems are issued for research and development, standards are needed. These standards provide a common ground for solving problems. The industrial standard is usually established in each country. For example, the United States has the ANSI standard. The United Kingdom has the BS standard, while Japan has the JIS standard. Table 7.1 summarizes various representative examples of industrial standards for many countries. As the table shows, advanced countries must have their own industrial standards. Therefore, it can be said that standardization is needed to drive countries’ industries and make them prosperous. Since there are many industrial fields, each standard has its classification. For example, electrical engineering has its own parts and mechanical engineering has its own parts, etc. However, now that industries’ and companies’ activities are border free, it would be very inconvenient for them to have different rules determined by their own standards. Therefore, we have the International Organization for Standardization (ISO), with international standards that are available all over the world. In 1946, the United National Standards Coordinating Committee proposed to establish the organization for the international standards in Switzerland. The purpose was to facilitate the international exchanges of commodities and cooperation among different countries. However, it was originally oriented toward European philosophy and policies to some extent. Therefore, in this chapter, we classify it into the European

© Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_7

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156 Table 7.1 Summary of various representative examples of industrial standards for many countries

7 Standardization—Current and Future Countries

Standards

Japan

JIS

Germany

DIN

The United States

ANSI

The United Kingdom

BS

Canada

CSA

France

NF

China

GB

Taiwan

CNS

Korea

KS

Indonesia

SNI

Russia

GOST

Vietnam

TCVN

Austria and New Zealand

AS/NZS

Malaysia

MS

Brazil

ABNT

Standard’s section and describe it there. However, it should include all of the countries that belong to the United Nations. The standards for biofilms are presented as they relate to the United States, European countries, and Japan.

7.1 The United States As shown in Table 7.1, the United States has ANSI as its national industrial standard. When we come to think about national standards, the United States has two kinds: the Occupational Safety and Health Administration (OSHA) and the American National Standards Institute (ANSI). OSHA is basically one of the laws, while ANSI was made by Standard Developing Organizations (SDO) and set as the national standard of the United States. When OSHA cites an ANSI standard as its own standard, the ANSI standard becomes a law like other OSHA standards. However, ANSI does not make the standard by itself and just approves the documents made by SDO. Some private organizations belong to SDO, so some of their proposed standards might just be arbitrary ones. Table 7.2 shows the private sectors belonging to SDO. Since biofilm research started in the United States and is full scale, the United States has been the only country with international standards for biofilms. These standards are mainly set in ASTM. The proposal for the standards is led by the Center of Biofilm Engineering (CBE) in Montana State University because this center has led the research activities in the United States and throughout the world. Up to 2017, representative standards proposed by the CBE included five standards in ASTM:

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Table 7.2 Various private sectors belonging to SDO Organizations

Abbreviated names

Contents

American Society for Testing and Materials International

ASTM International

Standards for industrial materials

Underwriters Laboratories, Inc.

UL

Preparation for various safety standards, tests, and recognitions

National Electrical Manufacturers Associations

NEMA

Electric goods and facilities except consumer electronics and their establishments

National Fire Protection Association

NFPA

The establishments of standards for fire controls and safety facilities

ASTM E2196 [1–5], ASTM E2562 [6–9], ASTM E2647 [10–14], ASTM E2871 [15–18], and ASTM E2799 [19–21]. In 2018, two new ASTM standards were added: ASTM 3151 [22] and ASTM E3161 [23].

7.1.1 ASTM E2196 The title of this standard is Standard Test Method for Quantification of Pseudomonas aeruginosa Biofilm Grown with Medium Shear and Continuous Flow Using a Rotating Disk Reactor [1–10]. This specially devised reactor (rotating disk type reactor) is used and the system is stirred continuously. The flow produces medium shear stress on the cylindrical coupon (diameter: 1.27 cm, thickness 1.5–4.0 mm, material: polycarbonate and others) and the biofilm formed on the specimen by Pseudomonas aeruginosa (ATCC70088) under these conditions is measured quantitatively. The concrete procedure is as follows. #1: #2: #3: #4:

#5:

#6:

6 pieces (coupons) are attached to the rotating disk (diameter: 7 cm). The stirrer rotates the disk at 200 rpm in the bacterial solution of Pseudomonas aeruginosa. Firstly, the batch process is carried out for 24 h. This is followed by a continuous operation with the new liquid culture for 24 h. In the case of a batch operation, the reactor is operated for 24 h at 21 °C. It is composed of 200 mL culture media (TSB, 300 mg/L) and 1 mL bacterial solution (108 CFU/mL) is added. (NOTE: TSB is Tryptic Soy Broth.) In the case of a continuous operation, the reactor is operated for 24 h at 21 °C. The solution is also TSB (30 mg/L) and it is provided a speed of 6.7 mL/m and discharged at the same rate. Suspended bacteria remaining on the surface of a coupon is removed by immersing in a buffer solution.

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Fig. 7.1 The schematic illustration for the measurement system for ASTM E2196

#7:

Biofilms produced on the surface of a coupon are scraped by sterilized wooden applicator sticks for 15 s and immersed in 9 mL buffer solution for about three to four times. #8: The surface of a coupon is washed by using pipets with buffer solution (The angle of the pipet is 60°. The volume of buffer solution used is 1 mL for each.) #9: The recovered solution is homogenized (20500 rpm × 30 s). #10: Serial dilution and plate culture. The result is evaluated as a common logarithm of viable bacteria count per area (log (CFU/cm2 )). The system is shown in Fig. 7.1.

7.1.2 ASTM E2562 The title of this standard is Standard Test Method for Quantification of Pseudomonas aeruginosa Biofilm Grown with High Shear and Continuous Flow Using a CDC Biofilm Reactor. A CDC biofilm reactor was devised and developed. It is a continuously stirred tank reactor where high shear stress is produced on the surface of the coupon specimen (cylindrical coupon, the diameter: 1.27 cm, thickness 3.0 mm, polycarbonate and other materials). Its purpose is to obtain the quantitative measurement of biofilms produced by Pseudomonas aeruginosa (ATCC 70088). The procedure is as follows.

7.1 The United States

159

#1:

3 coupons are attached to the polypropylene rod (diameter 1.9 cm, length 21 cm) and they are fixed at the top of the reactor. The total number of rods is 8. #2: The baffle located in the center of the reactor is rotated at 125 rpm in the bacterial solution of Pseudomonas aeruginosa. #3: The batch operation is carried out for 24 h and followed by the continuous operation for 24 h. #4: In the case of a batch operation, the reactor is operated for 24 h at 21 °C. It is composed of 350 mL culture media (TSB, 300 mg/L) and 1 mL bacterial solution (108 CFU/mL) is added. #5: In the case of a continuous operation, the reactor is operated for 24 h at 21 °C. The solution is also TSB (100 mg/L) and has the speed of 11.7 mL/m. It is discharged at the same rate. #6: Suspended bacteria remaining on the surface of a coupon is removed by immersing in a buffer solution. #7: Biofilms produced on the surface of coupon are scraped by sterilized wooden applicator sticks for 15 s and immersed in 9 mL buffer solution for about three to four times. #8: The surface of a coupon is washed by using pipets with buffer solution (The angle of the pipet is 60°. The volume of buffer solution is 1 mL for each.) #9: The recovered solution is homogenized (20500 rpm × 30 s). #10: Serial dilution and plate culture. (Steps #6 to #10 are similar to those for ASTM E2196.) The result is also evaluated as a common logarithm of viable bacteria count per area (log (CFU/cm2 )). The flow chart of the system is shown in Fig. 7.2.

7.1.3 ASTM E2647 The title of this standard is Standard Test Method for Quantification of Pseudomonas aeruginosa Biofilm Grown Using a Drip Flow Biofilm Reactor with Low Shear and Continuous Flow. The reactor used for the standard is a Drip Flow Biofilm Reactor shown in Fig. 7.3. The standard describes the method of biofilm formation and its quantification where Pseudomonas aeruginosa (ATCC 70088) produces biofilms at the interface between air and liquid phases under low shear stress in a Drip Flow Reactor. The concrete procedure is as follows. #1: #2: #3:

The glass slides are prepared (75 × 25 × 1 mm). However, other materials are available. The specimen (glass slide) is set in the chamber (3 cm × 10 cm) of the reactor. A 6 h batch operation is carried out in the horizontal state. Then the reactor is tilted 10°.

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Fig. 7.2 The schematic illustration of the evaluation process for ASTM E2562

Fig. 7.3 The schematic illustration of Drip Flow Biofilm Reactor for ASTM E2647

7.1 The United States

161

#4:

The new bacterial solution is provided and discharged continuously. The flow is laminar at a lower speed. The operation continues for 48 h to produce biofilms on specimens. #5: In the case of a batch operation, the reactor is operated for 6 h at 21 °C. It is composed of 15 mL culture media (TSB, 3000 mg/L) and 1 mL bacterial solution (108 CFU/mL) is added. #6: In the case of a continuous operation, the reactor is operated for 48 h at 21 °C. The solution is also TSB (270 mg/L) and it is provided the speed of 50 mL/h and discharged at the same rate. No stirring and the laminar flow. #7: Suspended bacteria remaining on the surface of a glass slide is removed by immersing in a buffer solution. Then the specimen is gently removed without stirring. #8: Biofilms produced on the surface of a coupon are scraped by a sterilized spatula or scraper for 15 s and immersed in 45 mL buffer solution for about three to four times. #9: The surface of a coupon is washed by using pipets with buffer solution. (The angle of the pipet is 60°. The volume of buffer solution is 1 mL for each.) The process is repeated five times. #10: The recovered solution is homogenized (20500 rpm × 30 s). #11: Serial dilution and plate culture. The result is also evaluated as a common logarithm of viable bacteria count per area (log (CFU/cm2 )).

7.1.4 ASTM E2871 The title of this standard is Standard Test Method for Evaluating Disinfectant Efficacy Against Pseudomonas aeruginosa Biofilm Grown in a CDC Biofilm Reactor Using a Single Tube Method. The CDC Biofilm Reactor and bacterial solution with Pseudomonas aeruginosa (ATCC 15422) are used. The specimen is a cylindrical coupon with the following: diameter: 1.27 cm, the thickness: 3.0 mm, and the material: the borosilicate glass. Biofilms are produced in the same way as with ASTM E2562 (Sect. 7.1.2). The procedure after the biofilm formation is as follows. #1: Suspended bacteria remaining on the surface of a glass slide is removed by immersing coupons with rods in a buffer solution. Then the specimen is gently removed without stirring. #2: The coupon is dropped from rods into a centrifuge tube and spun at a tilt against the bottom. #3: 4 ml of biocides or buffer solution is added to the centrifuge tube and air bubbles are removed by tapping it gently. #4: Still standing at 20 °C for a certain amount of time. #5: The addition of neutralizing agent and stirring.

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#6: Stirring by vortex mixer for 30 s and ultrasonic treatment for 30 s. The latter is repeated twice. #7: Serial dilution and plate culture. The result is also evaluated as a difference between a common logarithm of viable bacteria count per coupon (log (CFU/coupon)).

7.1.5 ASTM E2799 The title of the standard is Standard Test Method for Testing Disinfectant Efficacy against Pseudomonas aeruginosa (ATCC 15422) Biofilm Using the MBEC Assay. This standard describes the performance of biocides against biofilms produced by Pseudomonas aeruginosa, using Minimum Biofilm Eradication Concentration (MBEC) Assay. The purpose is the determination of the minimum effective concentration for disinfectants to control biofilms. The commercial MBED Assay device is used. The specimen is liquid (disinfectant) and the bacteria is Pseudomonas aeruginosa (ATCC 15422). The test procedure is as follows. #1: 96 well plates with peg lids are prepared. #2: 150 µL bacterial solution is added to the well plate. #3: Peg lids are closed and the pegs are immersed into the solutions. Biofilms form on the pegs. #4: On the other hand, the other 96 wells having specimens are prepared. The peg lids producing biofilms are attached to the specimens. Each well has 150 µL bacterial solution (105 CFU/mL in TSB, vibration culture) at 35 °C for 16–18 h. #5: After the attachment of samples with peg-lids, samples are moved to 96 well plates filled with neutralizer (recovery plates) and the reaction is stopped. Biofilms are broken by ultrasonic treatment (in 30 ± 5 min) and their constituents are moved from pegs to wells with neutralizer. #6: After serial diluting, the neutralizer for each well is distributed to TSA (Tryptic Soy Agar) and cultured. #7: The number of viable bacteria is measured and calculated as a common logarithm per area. The value for non-treated solution minus that for treated solution equals a decreasing value. #8: 100 µL TSB is added to the remaining well solution and cultured at 35 ± 2 °C for 24 h. The existence of bacteria is confirmed by a turbidity meter or absorption spectrometer. The quantitative analysis is carried out by the decreasing number defined in #7 and the qualitative analysis is evaluated by turbidity or the detection of bacteria.

7.1 The United States

163

7.1.6 ASTM E3151 The title of this standard is Standard Test Method for Determining Antimicrobial Activity and Biofilm Resistance, Properties of Tube, Yam, or Fiber Specimens. The standard describes the antibiofilm effect and antibacterial one of fibers, tubes, etc. We could say that medical catheters and suture threads might be targets too. The test tube for the culture (6 × 50 mm, 0.7 mL) should be prepared. The length of tubes and threads is up to 40 mm and the width up to 3 mm. As for bacteria, Staphylococcus epidermidis (ATCC 35984) is used. The procedure is as follows. #1: The specimen is cut to a 4 cm length coupon and put into culture test tubes. #2: In the case of tube samples, 0.3 mL bacterial solution is inoculated to the tube like specimens. Then 0.2 mL is inoculated to the inner side of the tube. #3: In the case of thread samples, 0.5 mL bacterial solution is inoculated. #4: They are stirred at 100 rpm by an orbital shaker and cultured for 35 × 24 h. #5: The liquid culture is neutralized, stirred, and treated by ultrasonic waves. The suspended bacteria for worked specimens and non-worked ones are measured. Then the antibacterial effect is evaluated. #6: Tubes and threads are taken out of the liquid culture and suspended planktonic bacteria are removed. Then they are neutralized, stirred, and treated by ultrasonic waves. The number of bacteria derived from biofilms is measured for worked and non-worked specimens. Then the antibiofouling effect is evaluated.

7.1.7 ASTM E3161 The title of this standard is Standard Practice for Preparing a Pseudomonas aeruginosa or Staphylococcus aureus Biofilm using the CDC Biofilm Reactor. This standard describes the concrete procedure for the preparation of P. aeruginosa (ATCC 15422) and S. aureus biofilms (ATCC 6538). The basic flow is almost the same as that for ASTM E2562. In this standard, a CDC biofilm reactor is used that produces high shear stress for specimens. The evaluation process and biofilm evaluation are almost the same as that for ASTM E2562. However, borosilicate glasses (diameter: 1.27 cm, the thickness: 3 mm) are used as specimens and the bacteria are different. Two kinds of bacteria are used. One of them is Pseudomonas aeruginosa, which is the same as that for ASTM E2562. However, the strain is different. In this case, ATCC 15422 is used. This time Staphylococcus aureus (ATCC 6538) is also used. Even though the test procedures are almost the same, the biofilm formation conditions are slightly different because the bacteria were changed. In the case of a batch test, P. aeruginosa are cultured in TSB (300 mg/L and 3 mg/L) at 21 °C for 24 h), while S. aureus is also in TSB (1 g/L and its 1/30 diluted solution) at 36 °C for 24 h. In the case of a continuous operation, 11.7 ml/min new culture solution is continuously provided and discharged. P. aeruginosa is cultured

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in TSB (100 mg/L and 1/300 diluted solution). The rotation rate is 125 rpm and operated at 21 °C for 24 h. On the other hand, S. aureus is cultured in TBS (1 g/L or 1/30 diluted) at 60 rpm. The culturing temperature is 36 °C and the operation time is 24 h.

7.1.8 Characteristics of the ASTM Standard in the USA The standards defined in ASTM, the USA, were prepared as the result of investigations and great effort made by the Center of Biofilm Engineering. This Center’s work (which seems to be the world’s first trials) has led world industries that are related to biofilm engineering. However, one should be careful to check the tests before applying the methods to their own cases. These methods have some common characteristics. #1: Bacteria used for the standard tests are mostly Pseudomonas aeruginosa and Staphylococcus aureus. Actually, Pseudomonas aeruginosa are easy to form biofilms. Also quite a few people in Europe and the USA have generic diseases caused by P. aeruginosa. It should be mentioned that S. aureus are found on our skin and appear in many cases of biofilms. However, one needs to think about his/her particular case and determine if the two bacteria would be appropriate model bacteria for applications to a particular industry. #2: The evaluation method is based on the bacterial number. There are many ways to evaluate biofilms, as explained in Chap. 6. The most difficult method is for evaluating biofilms quantitatively. From that viewpoint, the bacterial number could easily lead to the quantification. However, it would be hard to evaluate biofilms only by a bacterial number. As we have already said in the previous chapters, the biofilm process is composed of many steps. At the beginning, bacterial activity is the main factor and bacteria play an important role in forming biofilm. However, EPSs seem to be more important in affecting materials and components after a biofilm’s structure is established. In addition, not only viable bacteria, but also the dead or collapsed bacteria need to be considered in regards to biofilm problems. From the viewpoint, it is questionable how important and effective the bacterial number would be for specific individual cases. We feel that an evaluation method (based on bacterial number) would be available only to analyze cases that focus on the effect of bacteria on biofilms. In regards to a specific case, one should carefully check on the type of LBR to use and the availability of an appropriate standard (testing method). The evaluation for biofilms always depends on the type of problem that needs to be solved and on the environmental conditions involved.

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7.2 European Union 7.2.1 European Standards (EN) While each country in Europe has its own standard, the EU has its united standard among the member countries. It is called European Norm or European Standards. The standard aims to smoothly accelerate trade within the European Union and to arrange for certain industrial levels. It is mainly classified into two groups: the viewpoints of electrical engineering fields and those of the non-electrical engineering fields. The European Committee for Electrotechnical Standardization (CENLEC) is the organization for the former group [24]. The latter group has the European Committee for Standardization (CEN) [25]. The EU countries introduced the standards set by CENLEC and CEN into their own countries and set them to their nations’ standards. They had to properly adjust the balance between the united standards and their countries’ standards. As for biofilms, they don’t have any standards yet. However, they already have the disinfection standards [26]. Disinfection is related to biofilm problems, since bacteria in biofilms generally have high resistance to biocides. Basically, the European Standards relating to biofilms are those for disinfectants and antibacterial effects. In the EU, they don’t have any direct standards for materials’ biofilms. Their standards are related to disinfectants, which are connected to biofilms. Firstly, the quantitative suspension test was proposed in 1997 (BS EN 1276) and other standards have been established since then. The basic idea for the European standard for disinfection is shown in Fig. 7.4. In this concept, the suspension test is the antibacterial or antifungal effects of disinfectants against microbial organisms in solutions. Therefore, the standard might describe the antimicrobial effects against planktonic bacteria. On the other hand, the surface test is carried out on samples’ surfaces. Since the antimicrobial effect in this case is related to interfaces and the bacteria there, it includes the concept of biofilms. Fig. 7.4 The concept of European antibacterial (and antifungal) effects

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7.2.2 Phase 1—Basic Suspension Test This corresponds to EN 1040 and EN 1275. Both are suspension tests and describe the antimicrobial effects against planktonic organisms. EN 1040 checks antibacterial effects and EN 1040 describes antifungal effects. In EN 1040, two bacteria (Pseudomonas aeruginosa and Staphylococcus aureus) are generally used. The developed disinfectant and its diluted solution are used and the temperature should be kept at 20 °C. The number of viable bacteria and the change with time are measured. In EN 1275, Aspergillus niger or Candida albicans is usually used as the test organism. Basically, fungicidal agents (less than 80% of the near test solution) are used and the temperature should be kept at 20 °C. The model fungi are added to the solution and the number of viable fungi is measured with time.

7.2.3 Phase 2 Part 1—Quantitative Suspension Test This part includes two classes. One of them is for antibacterial disinfectants and the corresponding standard is EN 13717. It is basically a qualitative surface test. On the other hand, EN 13697 is a quantitative surface test. The standards could determine the variable parameters to interfere with substances, temperature, and contact times. The bacteria and fungi used for these tests are Pseudomonas aeruginosa, Staphylococcus aureus, Enterococcus hirae, Aspergillus niger and Candida albicans. A certain amount of bacteria and developed disinfectants are placed on some materials’ surfaces. Then the change of viable organisms’ numbers is measured to determine the efficacy of the disinfectants.

7.2.4 Phase 3—Field Test An example of the field test is mentioned here. It is EN 1500 and corresponds to hand sanitization. The test anticipates that the result would be applied to skin and glove covered hands. Escherichia coli or Staphylococci are used as the bacteria. The developed hand sanitizer and the control solution such as 60% propanol are put on materials used for gloves. Then the change of bacterial numbers with time is measured. Usually, the five log reduction of the bacterial number is needed to determine the efficacy.

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7.2.5 International Standards (ISOs) Standards should be international, even if they were mainly established in European countries. (NOTE: ISO stands for the International Organization for Standardization.) A representative standard is the one prepared for dental treatment. In dental technology and science, biofilms are often present. Pellicle (a protein film that forms on the surface enamel of teeth) protects the teeth from acids produced in the mouth. However, biofilm can grow on this film. Therefore, it is very natural for the standard to be released for dental fields. The standard is ISO 16954 [27, 28] and the title is “Dentistry - Test methods for dental unit waterline biofilm treatment.” The original purpose for this standard was to evaluate the efficacy of cleaning methods for biofilms formed on dental cleaning machines provided with non-sterilized water. The apparatus for the cleaning machine used for real treatments or simulated ones is used for the test. The specimens are water for the apparatus and the water line tubes. As for bacteria, Pseudomonas aeruginosa (ATCC 700888) and Klebsiella pneumonia (ATCC 13882) are used. The apparatus is operated at 23 ± 3 °C for seven days. The operation cycle is the combination of 30 s on and 9 min off. One day’s operation is composed of 30 cycles. The capability of controlling biofilms is evaluated by the number of viable bacteria and the SEM observation results for the antibacterial treated apparatus as compared with the nontreated one. The test period should be more than 4 weeks, where SEM observation could confirm a certain amount of biofilm. For measuring the number of viable bacteria, bacteria are cultured on R2A agar plates at 23 ± 3 °C for 7 days. For SEM observations, the surface is observed on three places with at least 1000x to 5000x magnification. The results are evaluated by four step scales: Scale 1; no biofilms, and no bacteria, Scale 2; biofilms and bacteria are observed in amounts less than 10%, Scale 3; biofilms and bacteria are observed in amounts less than 50%, and Scale 4; biofilms and bacteria are observed in amounts more than 50%. As for the antibacterial effect test for materials, the standard ISO 22196 [29–33] is used, which is derived from the Japanese standard (JIS) Z2801 [34]. It is usually called the Film Covering Method. The specimens are placed in a plastic Petri dish, while the bacterial solution is prepared as follows: the bacteria are incubated in 10 mL of a nutrient broth for 24 h at 35 °C, and then diluted two-thousand fold with sterilized water and established as a bacterial solution. The diluted bacterial solution is applied to the specimen (16 µL/cm) and then a polymer film is laid over the solution. The sample is kept in an incubator for 24 h at 35 °C. After the incubation, a solution of 10 mL of sterilized water containing 200 µL of Tween® 80 (a nonionic surfactant and emulsifier) is introduced into the plastic Petri dish. The bacteria attached to the specimen and polyethylene film are washed with the aqueous solution. To determine the number of viable cells, serial decimal dilutions of the cell suspension are made, a 0.1 mL portion of which is uniformly spread on an agar medium. The plate is incubated at 35 °C for 24 h and the colonies formed are counted. The viable cell count is represented as a colony forming unit per milliliter (CFU/mL). The final

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colony formation number is measured to evaluate the antibacterial properties. This method is explained once again in the following section.

7.3 Japan In Japan, the concept for antibacterial effect has been traditionally well established. Overall, one could say that the Japanese are very sensitive to the antibacterial effect. Therefore, there are some standards for the antibacterial effect of materials. The representative method is the Film Covering Method. As explained in the previous section, the Japanese industrial standard for the antibacterial effect is summarized in JIS Z2801 that was established in 2000 [34]. This standard was further developed and resulted in the ISO. The test is characterized as an interaction between bacteria and materials’ surfaces (standard: 50 mm × 50 mm) covered with polyethylene (PE) film (standard: 40 mm × 40 mm). Since the size of covered films and types of materials, could be adjusted and changed, the method has applications to versatile practical cases. In addition, the covering films would control the evaporation to some extent. Figure 7.5 shows the procedure schematically.

Fig. 7.5 The outline for the Film Covering Method

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The specimens are sterilized by ethanol (3 non-treated specimens and three antibacterial treated specimens). They are placed in a sterilized Petri dish, respectively. A 0.4 mL (2.5 to 10 × 105 CFU/ml) bacterial solution is inoculated on the specimen’s surface. Then non-antibacterial film such as PE is put on the bacterial solution, so that the contact area is covered and closed with the polymer films. The Petri dish is covered with a lid and cultured in an incubator at 35 ± 1 °C, with a relative humidity more than 90% for 24 ± 1 h. After the culture takes place, specimens and covered films are washed with 10 mL of SCDLP (Soybean-Casein Digest broth with Lecithin and Polysorbate 80). The washed-out-solution is diluted 10 times and inoculated on an agar plate at 35 ± 1 °C for 40–48 h. Then the number of viable bacteria is measured from the colony formation unit. The antibacterial activity is defined as follows. Antibacterial activity = log(Nc/Np)

(7.1)

Here Nc is the average bacterial number on the control, while Np is the average bacterial number on the target specimen’s surface. When the antibacterial activity is more than 2, one could say that the target materials have an antibacterial effect. As for the antibacterial effect, there are other standards. The standard for fibers is described in JIS L1902 [35] and the standard for photocatalytic TiO2 is JIS R1702 [36, 37]. These standards are basically for the antibacterial effect. For dental purposes, JIS T5111 was proposed [38]. However, this standard is just a transplant from ISO 16954. Therefore, the detailed information is omitted here. For readers, who are interested in the details, please refer to the references. As described above, the direct standards have only been proposed by the United States. In light of that, the USA has led the world for the standard (for biofilm). In Europe, all of the standards are related to disinfectants. Japanese standards are oriented toward the antibacterial effect. However, the antibacterial effect is different from antibiofilm properties. The contribution of materials to biofilm control is still lacking for all of the industrial standards. Now in Japan, SIAA (the Society of International Sustaining Growth for Antimicrobial Articles) which proposed the Film Covering Method for JIS and ISO is accelerating the pace to establish a biofilm standard. According to their recent report, they are seeking for the method to evaluate materials’ antibiofilm properties by the staining of crystal violet. The proposal is greatly anticipated [39, 40].

References 1. ASTM International. Standard test method for quantification of Pseudomonas aeruginosa biofilm grown with medium shear and continuous flow using rotating disk reactor. https:// www.astm.org/Standards/E2196.htm. 2. Garo, E., Eldridge, G. R., Goering, M. G., Pulcini, E. D., Hamilton, M. A., Costerton, J. W., et al. (2007). Asiatic acid and corosolic acid enhance the susceptibility of Pseudomonas aeruginosa biofilms to tobramycin. Antimicrobial Agents and Chemotherapy, 51(5), 1813–1817.

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3. Rayner, J., Veeh, R., & Flood, J. (2004). Prevalence of microbial biofilms on selected fresh produce and household surfaces. International Journal of Food Microbiology, 95, 29–39. 4. Buckingham-Meyer, K., Goeres, D. M., & Hamilton, M. A. (2007). Comparative evaluation of biofilm disinfectant efficacy tests. Journal of Microbiological Methods, 70, 236–244. 5. Kanematsu, H., Barry, D. M., Ikegai, H., Yoshitake, M., & Mizunoe, Y. (2017). Biofilm evaluation methods outside body to inside—Problem presentations for the future. Medical Research Archives, 5, 1–17. 6. ASTM International. (2012). ASTM E2562-12 standard test method for quantification of Pseudomonas Aeruginosa biofilm grown with high shear and continuous flow using CDC biofilm reactor. West Conshohocken, PA: ASTM International. 7. Garvey, M., Rabbitt, D., Stocca, A., & Rowan, N. (2015). Pulsed ultraviolet light inactivation of Pseudomonas aeruginosa and Staphylococcus aureus biofilms. Water and Environment Journal, 29, 36–42. 8. Martinez-Gutierrez, F., Boegli, L., Agostinho, A., Sánchez, E. M., Bach, H., Ruiz, F., et al. (2013). Anti-biofilm activity of silver nanoparticles against different microorganisms. Biofouling, 29, 651–660. 9. ASTM Testing Materials. ASTM E2562-12: standard test method for quantification of Pseudomonas Aeruginosa biofilm grown with high shear and continuous flow using CDC biofilm reactor. 10. Standard, A. (2008). ASTM E2647-08 standard test method for quantification of a Pseudomonas aeruginosa biofilm grown using a drip flow biofilm reactor with low shear and continuous flow. ASTM International. 11. Woods, J., Boegli, L., Kirker, K. R., Agostinho, A. M., Durch, A. M., Delancey Pulcini, E., et al. (2012). Development and application of a polymicrobial, in vitro, wound biofilm model. Journal of Applied Microbiology, 112, 998–1006. 12. Alvarado-Gomez, E., Perez-Diaz, M., Valdez-Perez, D., Ruiz-Garcia, J., Magaña-Aquino, M., Martinez-Castañon, G., et al. (2018). Adhesion forces of biofilms developed in vitro from clinical strains of skin wounds. Materials Science and Engineering: C, 82, 336–344. 13. Velázquez-Velázquez, J. L., Santos-Flores, A., Araujo-Meléndez, J., Sanchez-Sanchez, R., Velasquillo, C., González, C., et al. (2015). Anti-biofilm and cytotoxicity activity of impregnated dressings with silver nanoparticles. Materials Science and Engineering: C, 49, 604–611. 14. Pérez-Díaz, M., Alvarado-Gomez, E., Magaña-Aquino, M., Sánchez-Sánchez, R., Velasquillo, C., Gonzalez, C., et al. (2016). Anti-biofilm activity of chitosan gels formulated with silver nanoparticles and their cytotoxic effect on human fibroblasts. Materials Science and Engineering: C, 60, 317–323. 15. ASTM International. (2013). ASTM E2871-13 standard test method for evaluating disinfectant efficacy against Pseudomonas aeruginosa biofilm grown in CDC biofilm reactor using single tube method. West Conshohocken, PA: ASTM International. 16. Fritz, B., Walker, D. K., Goveia, D., Parker, A. E., & Goeres, D. M. (2015). Evaluation of Petrifilm™ Aerobic Count Plates as an equivalent alternative to drop plating on R2A Agar Plates in a biofilm disinfectant efficacy test. Current Microbiology, 70, 450–456. 17. Saha, R., Saha, N., Atwain, A., & Donofrio, R. S. (2014). Evaluation of disinfection efficacy of ozone and chlorinated disinfectant against the biofilm of Klebsiella michiganensis and Pseudomonas aeruginosa. Annals of Microbiology, 64, 1607–1613. 18. Wahlen, L., Parker, A., Walker, D., Pasmore, M., & Sturman, P. (2016). Predictive modeling for hot water inactivation of planktonic and biofilm-associated Sphingomonas parapaucimobilis to support hot water sanitization programs. Biofouling, 32, 751–761. 19. Allan, N., Omar, A., Harding, M., & Olson, M. (2011). A rapid, high-throughput method for culturing, characterizing and biocide efficacy testing of both planktonic cells and biofilms. In A. Mendez-Vilas (Ed.), Science against microbial pathogens: communicating current research and technological advances. Microbiology book series (pp. 864–871). Formatex. 20. Percival, S. L., Mayer, D., & Salisbury, A. M. (2017). Efficacy of a surfactant-based wound dressing on biofilm control. Wound Repair and Regeneration, 25, 767–773.

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21. Howard, R., Harding, M., Daniels, G., Mobbs, S., Lisowski, S., & De Boer, S. (2015). Efficacy of agricultural disinfectants on biofilms of the bacterial ring rot pathogen, Clavibacter michiganensis subsp. sepedonicus. Canadian Journal of Plant Pathology, 37, 273–284. 22. ASTM International. (2018). Standard test method for determining antimicrobial activity and biofilm resistance properties of tube, yarn, or fiber specimens. In E35.15 (Vol. ASTM E315118). 23. ASTM International. (2018). Standard practice for preparing a Pseudomonas aeruginosa or Staphylococcus aureus biofilm using the CDC biofilm reactor (Vol. ASTM E3161-18). ASTM International. 24. CENELEC. European Committee for Electrotechnical Standardization. https://www. cenelec.eu/. 25. CEN. European Committee for Standardization. https://www.cen.eu/Pages/default.aspx. 26. Sandle, J. T. (2017). The European approach to disinfectant qualification. La Vague, N52, 45–48. 27. International Standard. (2019). Dentistry—Test methods for dental unit waterline biofilm treatment. 28. Offner, D., Fioretti, F., & Musset, A.-M. (2016). Contamination of dental unit waterlines: Assessment of three continuous water disinfection systems. BDJ Open, 2, 16007. 29. International Standard. (2019). Measurement of antibacterial activity on plastics and other non-porous surfaces. 30. Ando, Y., Miyamoto, H., Noda, I., Sakurai, N., Akiyama, T., Yonekura, Y., et al. (2010). Calcium phosphate coating containing silver shows high antibacterial activity and low cytotoxicity and inhibits bacterial adhesion. Materials Science and Engineering: C, 30, 175–180. 31. Torlak, E., & Sert, D. (2013). Antibacterial effectiveness of chitosan–propolis coated polypropylene films against foodborne pathogens. International Journal of Biological Macromolecules, 60, 52–55. 32. Vincent, M., Hartemann, P., & Engels-Deutsch, M. (2016). Antimicrobial applications of copper. International Journal of Hygiene and Environmental Health, 219, 585–591. 33. Chiu, T.-W., Yang, Y.-C., Yeh, A.-C., Wang, Y.-P., & Feng, Y.-W. (2013). Antibacterial property of CuCrO2 thin films prepared by RF magnetron sputtering deposition. Vacuum, 87, 174–177. 34. ICROCHEM. JIS Z 2801 test for antimicrobial activity of plastics. https://microchemlab.com/ test/jis-z-2801-test-antimicrobial-activity-plastics. 35. Center, M. S. Antimicrobial activity JIS 1902 & ISO 20743. http://www. manufacturingsolutionscenter.org/antibacterial-activity-JIS-L1902-ISO-20743.html. 36. Yao, Y., Ochiai, T., Ishiguro, H., Nakano, R., & Kubota, Y. (2011). Antibacterial performance of a novel photocatalytic-coated cordierite foam for use in air cleaners. Applied Catalysis B: Environmental, 106, 592–599. 37. Nakano, R., Hara, M., Ishiguro, H., Yao, Y., Ochiai, T., Nakata, K., et al. (2013). Broad spectrum microbicidal activity of photocatalysis by TiO2. Catalysts, 3, 310–323. 38. Japanese Industrial Standard. Dentistry—Test methods for dental unit waterline biofilm treatment http://www.kikakurui.com/t5/T5111-2018-01.html (in Japanese). 39. Kanematsu, H. (2017). A new international standard for testing antibacterial effects. Advanced Materials & Processing, 175, 26–29. 40. SIAA. SIAA home page. https://www.kohkin.net/en_index.html.

Chapter 8

Biofilm Problems and Environments

Abstract This chapter describes biofilm problems in various environments. It starts by introducing the marine environment and goes on to discuss biofilm problems like biofouling and corrosion that damage structures used in the building of sea vessels and ports. These problems are very expensive for the shipping industry. Next, the topic of soil and its bacteria are discussed. An emphasis is placed on pathogens that cause food contamination and plant diseases. The reader is also made aware of the biofilms colonizing in our households, especially in the bathrooms and kitchens. In this section, biofilms are mentioned that contaminate food, spread illness, stain counters, etc. Tips are provided too for effective cleaning to control and prevent biofilms in our homes and elsewhere. The next topic presents pathogens that cause problems for the food processing industries. This is followed by a description of how biofilms contribute to the slime and scale buildup in pipes and heat exchangers, which result in a reduced ability for liquid flow and heat exchange that might increase a company’s operational cost and energy consumption. The final section of this chapter is about biofilm problems in hospitals and other medical areas.

8.1 Marine Environments Biofilms (a form of biofouling) cause problems in the marine (seawater) environment. Biofouling is when microorganisms, plants, algae, or animals accumulate on wetted or submerged surfaces. It includes microfouling (biofilm formation and the attachment of bacteria) and macrofouling (the attachment of larger organisms). The fouling community includes hard fouling organisms like barnacles and soft fouling organisms like seaweed and biofilm (slime). Microfouling leads to macrofouling. MICROFOULING refers to the degrading of material by the bacteria attached to it. Bacteria (including marine bacteria) tend to attach to surfaces containing conditioning films composed of carbon compounds [1]. These carbon compounds serve as a form of needed nutrition for bacteria. When the number of bacteria (attached to a material’s surface) reaches a certain threshold value, then the bacteria simultaneously excrete organic polymers (EPS) outside of their cells through a process

© Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_8

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Fig. 8.1 This diagram shows the process for biofilm formation

called quorum sensing [2]. These sticky, organic polymers actually encase the bacteria and protect them from temperature and pH fluctuations, biocides, detergents, etc. The film-like encasing material is a biofilm (a product of microfouling). Figure 8.1 displays the biofilm formation process [3]. The marine environment experiences corrosion that damages structures used in the building of ships, oil pipelines, ports, etc. In addition to regular chemical corrosion (such as that from the salt in seawater), studies show that microorganisms (in the form of complex biofilms) directly participate in the corrosion process or accelerate/influence the corrosion action. These microorganisms degrade metal surfaces and cause crevices and pitting. Corrosion is a real concern that is very costly, about 2.5 trillion U.S. dollars annually. The Worldwide Corrosion Authority (NACE International) estimated that the global cost of corrosion was this amount in 2016 [4]. Metal corrosion is a spontaneous process in which electrons (from a neutral metal) are transferred to a final acceptor. In aerobic conditions, oxygen is the main final electron acceptor. Therefore, for the case of iron, one can see the rust (iron oxide) that forms. For biofilms on iron and steel, different microbial, metabolic groups perform various actions on the metal. Environmental conditions (such as temperature variation, pH, and oxygen levels, etc.) are important too. Despite the difficulty in establishing a relationship between the presence of a bacterial strain and its role in the corrosion of a metal surface, it is widely accepted that members of the Delaproteobacteria class play a direct corrosive role on metal alloys [5]. MACROFOULING is the attachment of larger organisms to materials’ surfaces. In the marine environment, oysters and barnacles, etc. generally live on solid surfaces

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like wood and the bottom of ships. These larger organisms attach to materials’ surfaces to get food, just as bacteria attach to surfaces for nourishment. Since barnacles and oysters use plankton for nutrition, it is natural for them to get the plankton in biofilms formed on solid surfaces. Therefore, we can say that microfouling leads to macrofouling. Marine colonization processes are generally sequential steps for the formation of microbial biofilms which precede the attachment of larger organisms. Since fouling by secondary organisms can restrict light, nutrition, etc. to a settled organism, marine organisms have developed defenses against such problems. Evidence suggests that chemical cues from marine organisms affect the colonization processes of secondary colonizers [6]. Biofouling occurs everywhere, but is most significant economically to the shipping industry. The buildup of biofoulers on marine vessels increases drag, fuel consumption, cost, and the length of the journey. At the same time, it can deteriorate materials, affect the function of various structures, and reduce the overall hydrodynamic performance of the ship. Figure 8.2 displays an instrument, for making measurements, Fig. 8.2 An instrument for making measurements that is covered with zebra mussels

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Fig. 8.3 Organisms that lived, grew, and died, attached to the bottom of a wooden fishing boat. They can be removed by mechanical methods

covered with zebra mussels [7]. Also Fig. 8.3 shows dead organisms (biofouling) attached to the bottom of a wooden fishing boat [8]. BIOFOULING NEEDS TO BE CONTROLLED. Since microfouling leads to macrofouling, an initial approach is to control biofilm formation. Biocides (chemical substances incorporated into anti-fouling surface coatings) are used to prevent/control biofilm formation by repelling/killing microorganisms. They often target bacteria, the microorganisms that produce biofilm. Non-toxic anti-sticking coatings are also used to prevent the attachment of microorganisms. These coatings are generally based on organic polymers, in order to allow for the addition of extra functions such as antimicrobial activities. The most common class of these coatings depends on friction and low surface energy, which result in hydrophobic surfaces that are smooth. Fluoropolymers and silicone coatings are often used [9]. The second class of non-toxic anti-fouling coatings is called hydrophilic coatings. They rely on high amounts of hydration [10]. An example of these coatings is based on highly hydrated zwitterions (formerly called dipolar ions). These coatings prevent the attachment of bacteria, which prevent the formation of biofilm, which prevents the attachment of macrofoulers. It is wise to avoid decreasing the lifetime of coatings. Therefore, use the minimum amount of force required to remove marine organisms attached to the hulls of ships. Figure 8.4 shows a typical reaction of a microorganism (like bacteria) to various types of surfaces [11]. Ultrasonic, electrical, and mechanical technologies are also used to control biofilm formation [12]. These methods, along with surface modification, are used to control macrofoulers too. For example, zebra mussels are stunned or killed by energizing the water with high voltage (electrical energy).

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Fig. 8.4 A microorganism’s reaction to various surfaces. a This is an untreated surface. b This is a surface with a coating that contains a biocide that repels or kills. c This is a surface with a non toxic/non stick coating

8.2 Soil Environments Soil is a mixture that includes organic matter, minerals, organisms, gases, and liquids. Bacteria are abundant microbes in soil and mostly live in the top 10 cm where organic matter is present. One gram of soil could contain up to one billion bacteria. Soil is a source of carbon, a habitat for organisms, a recycling system for nutrients and organic wastes, a medium for water storage, and a place for plants to grow. It undergoes physical and chemical changes due to climate, its original parent materials, weathering, etc. Some parent mineral materials for soil are quartz (SiO2 ), calcite (CaCO3 ), feldspar (KAlSi3 O8 ), and mica [K(Mg, Fe)3 AlSi3 O10 (OH)2 ]. A soil’s texture depends on particles of clay, sand, and silt that make it up. Clay is made up of very small particles of aluminum silicate. Sand is basically silicon dioxide and silt is considered to be very fine sand, clay, etc. Figure 8.5 shows a typical soil profile [13]. Soils that have developed for a long period of time are considered to be mature

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Fig. 8.5 A typical soil profile is displayed with dark topsoil and reddish layers of subsoil

and contain a few horizons (horizontal layers that vary with different types of soils). Horizon O represents the surface and displays grass, weeds, sticks, etc. that have not completely decayed. Horizon A is made of topsoil, a thin layer of organic matter and bacteria. The B horizon is called subsoil. It has roots and mainly includes inorganic material like rock particles, sand, and gravel. The C horizon lies on bedrock. Figure 8.6 shows various types of soil [14]. Note that loam is a soil composed of about 40% sand, 40% silt, and 20% clay. These proportions can vary and result in different types of loam such as sandy loam. Soil texture affects the diversity of bacterial communities [15]. As previously mentioned, bacteria are abundant microbes in soil. Also one gram of soil could hold up to a billion bacteria. Some bacteria are fragile and can die because of small changes in the soil. Others are strong and can withstand the cold and heat and lie dormant for years until favorable conditions are available. Bacteria in soil belong to four functional groups. Most of them are decomposers that consume simple carbon compounds. Another group is called the mutualists, which form partnerships with plants. A good example is nitrogen-fixing bacteria that form symbiotic relationships with the roots of legumes like clover. Pathogens form the group of harmful bacteria in soil. The fourth group includes lithotrophs or chemoautotrophs. They get energy from compounds of iron, sulfur, or hydrogen instead of from carbon compounds. Harmful bacteria in soil can cause potential harm to humans, trees, and plants. Some forms of bacteria produce poisonous toxins, which can be fatal if the spores of

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Fig. 8.6 This diagram shows soil types based on clay, silt, and sand composition as used by the United States Department of Agriculture (USDA)

such bacteria are inhaled, ingested, or transferred through a wound. Bacillus cereus is a harmful bacteria commonly found in soil. It is a Gram-positive, rod-shaped, spore forming bacteria. Some strains are harmful to humans and cause poisoning. Symptoms include vomiting, fever, and diarrhea. Researchers investigated various types of soil in Georgia and found that bacteria of the genus Bacillus prevailed in most of the studied soils [16]. Figure 8.7 shows a scanning electron micrograph of Bacillus cereus [17]. Keep in mind that these bacteria (like others) produce biofilms (sticky, thin film outer coverings) to protect them from antibiotics, detergents, etc. Bacillus anthracis is a Gram-positive, spore-forming, rod-shaped bacterium that synthesizes a protein capsule, poly-D-gamma-glutamic acid: a polysaccharide layer, [18]. It can survive for years in the soil. However, the inhalation of its spores from contaminated soil can result in illness. Anthrax produces a toxin that can result in skin ulcers, respiratory distress, and possible death. Agrobacterium tumefaciens (which is also called Agrobacterium radiobacter) is a rod-shaped Gram-negative bacterium (found in soil) that causes disease in plants. It enters them (from the soil) through their roots or stems and results in crown gall disease. The host tree or plant dies as a result of tumor development and metabolism changes. Figure 8.8 shows sunflowers infected with the crown gall disease [19]. Clostridium perfringens is an anaerobic, Gram-positive, rod-shaped, bacteria mostly found in soil and water. It forms spores and is responsible for food-borne

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Fig. 8.7 Above is a SEM (scanning electron microscope) image of Bacillus cereus Fig. 8.8 Sun flowers infected with crown gall disease

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Fig. 8.9 Above is a display of Clostridium perfringens using Gram stain. As you can see, the Gram-positive bacteria stain purple

illnesses. C. perfringens is a leading cause of food poisoning in the United States and Canada [20]. Its infections (for example) show evidence of tissue necrosis (the presence of dead cells), gas gangrene (a situation where tissue gas is produced in gangrene), and bacteremia (the presence of bacteria in the blood). Treatment of severe symptoms includes the use of antimicrobial drugs, surgery, etc. To prevent the growth of Clostridium perfringens spores, one should cook food thoroughly, refrigerate leftover food, and reheat leftovers before serving them. Figure 8.9 displays Clostridium perfringens using Gram stain [21]. Clostridium tetani is a rod-shaped, anaerobic, Gram-positive bacterium that is common in soil. It moves by flagella that surround its body. Under certain conditions, the flagella are shed and spores form. C. tetani sometimes causes the disease tetanus, which generally starts when spores enter the body through a puncture wound. Tetanus (which is also known as lockjaw) affects the nervous system. It can be prevented with the tetanus vaccine. Figure 8.10 is a micrograph of a group of C. tetani bacteria that cause tetanus in humans [22].

8.3 Household Environments Biofilms are produced by bacteria and can be found everywhere. It is important to be aware of biofilms in our households, especially in the bathrooms and kitchens. Figure 8.11 shows a typical bathroom in the United States [23]. Biofilm containing

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Fig. 8.10 Above is a micrograph of a group of C. tetani bacteria that cause tetanus in humans

the bacteria Serratia marcescens produces a pinkish color and is commonly found in bathtubs, toilet bowls, sinks, tiles, and showers. It grows in moist areas where fatty substances (like soap scum) and materials with minerals are located. The biofilm containing this type of bacteria looks like a pinkish red to orange slime. It can infect open wounds and the respiratory and urinary tracts of immuno-compromised individuals. Figure 8.12 shows Serratia marcescens growing on a slice of bread [24]. It is a species of rod-shaped Gram-negative bacteria. Investigators discovered that pink biofilms develop rapidly in bathrooms (of Japan), and are difficult to remove, and recur quickly. They tested a number of pink biofilms in various bathrooms and found Methylobacterium strains to be the major genus in all of the samples, which were analyzed by fluorescence in situ hybridization [25]. The strains exhibited high tolerance to cleaning agents and to desiccation. Some of the strains cause infections in humans. A picture of Methylobacterium is displayed in Fig. 8.13 [26]. The most common types of bacteria in bathrooms include Bacteroidaceae, Escherichia coli (E. coli), Streptococcus and Salmonella [1]. Bacteroidaceae is a family of anaerobic, non-spore forming bacteria that contain Gram negative rods. They may be disease causing (pathogenic). See Fig. 8.14 [27]. E. coli is Gramnegative, rod-shaped bacteria. Most strains of this type of bacteria are harmless. However, some can cause serious food poisoning in humans. Figure 8.15 shows a photo of E. coli [28]. Streptococcus is Gram positive bacteria. It has a spherical shape and tends to grow in pairs or chains. Some of its species cause illnesses like pneumonia, meningitis, pink eye, etc. Salmonella is rod-shaped Gram negative bacteria.

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Fig. 8.11 This is a picture of a typical American bathroom

Fig. 8.12 Serratia marcescens growing on a slice of bread

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Fig. 8.13 Methylobacterium

Most infections caused by them are due to ingestion of food contaminated by animal or human feces. Figure 8.16 shows a photo of Salmonella [29]. The toilet (including the seat, bowl, flush handle, etc.) has millions of bacteria. Most of them are located deep inside the toilet bowl. Some airborne microbes can be released when the toilet is flushed. It is best to keep the lid down when flushing the toilet. Also the floor, counters, sink, faucets, and the light switch contain bacteria. However, the worst item is our toothbrush with thousands of bacteria per square inch. It can contain Staphylococci bacteria and E. coli. There is no need to panic because we have similar types of bacteria and more of them in our mouth. Biofilms colonize in our kitchens. A common place to find biofilms is in kitchen cutting boards, especially if they have deep cuts or cracks that may provide a home for bacteria. Bacteria that cause food borne illnesses and form biofilms include Listeria monocytogenes, E. coli, and Salmonella. If these bacteria are on a cutting board (as a result of cutting raw chicken, etc.), then they can get into our food and make us sick. For example, if the cutting board is not properly cleaned and other food like bread is cut on it, then the bacteria can be transferred to the bread. E. coli and Salmonella were previously mentioned. Listeria monocytogenes is a Gram-positive, rod-shaped bacterium that causes listeriosis through the ingestion of contaminated food. It can

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Fig. 8.14 A member of the Bacteroidaceae family: Bacteroides biacutis anaerobically cultured in blood agar medium

Fig. 8.15 Low temperature electron micrograph of E. coli magnified 10,000 times

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Fig. 8.16 A color enhanced scanning electron micrograph shows Salmonella typhimurium (in red) invading cultured human cells. This type of Salmonella causes diarrhea and can be transferred to humans through raw or under cooked food (especially meat)

result in gastrointestinal problems, sepsis, etc. Figure 8.17 shows a picture of this bacterium [30]. Scientists analyzed appliances and surfaces in kitchens and found various types of bacteria. A picture of a typical kitchen is displayed in Fig. 8.18 [31]. They recruited 20 volunteer families to swab 14 common items in a kitchen such as a can opener, a microwave keypad, a coffee maker, etc. Their project had the following results [32]. Twenty-five percent of the tested items (examples: refrigerator’s meat compartment, can opener, blender gasket) contained E. coli bacteria. This bacterium causes diarrhea, urinary tract infections, pneumonia, etc. Salmonella was found on twenty-five percent of the tested items which included the meat and vegetable compartments in the refrigerator and the refrigerator ice and water dispensers. This type of bacteria causes salmonellosis, a disease that causes diarrhea, fever, stomach pain, etc. Ten percent of the tested items contained Listeria monocytogenes, bacteria that causes fever, muscle aches, and gastrointestinal symptoms. This type of bacteria was found in the refrigerator’s vegetable compartment.

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Fig. 8.17 Above is an electron micrograph of a flagellated Listeria monocytogenes bacterium

Other researchers studied 80 surfaces in kitchens of four households. They used high-throughput sequencing of the 16S rRNA gene to explore biogeographical patterns of bacteria [33]. Most sequences belonged to Actinobacteria, Bacteriodetes, Firmicutes, and Proteobacteria. Actinobacteria are Gram-positive bacteria and Bacteriodetes are Gram-negative, rod-shaped bacteria. Firmicutes form a phylum of bacteria that are mostly Gram-positive with a cell wall structure. Proteobacteria form a phylum of Gram-negative bacteria that include a variety of pathogens. These researchers found that the major sources of bacteria in the kitchen are humans and raw foods brought into the kitchen. Individuals are exposed to microorganisms by handling, preparing, and eating food. Indirect exposure results by contacting contaminated surfaces, dish cloths, dish towels, sponges, etc. The investigators determined that Gram-negative Proteobacteria were more abundant on moist surfaces, while Gram-positive spore-forming Firmicutes appeared on dry surfaces (example: floor) and cold surfaces (example: refrigerator). NOTE: It should be mentioned that results for experiments such as this one are influenced by variables like the kitchen design, the counter material, the cleaning agents used, the frequency of cleaning the kitchen, the types of food prepared there, etc. In addition to spreading illness, biofilms can stain surfaces, produce rust, and create expenses for remediation in the household. The best method for controlling biofilms is to prevent them from colonizing. Once established, they are hard to get rid

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Fig. 8.18 A typical kitchen

of. One should clean frequently and keep surfaces dry and free of residue (a form of nutrition for bacteria). In order for an antimicrobial chemical to be effective, biofilms must first be agitated, broken up, and removed before the chemical is added. The chemical should remain on the surface a while before rinsing it away. (This process resembles the removal of dental plaque: a hardened form of biofilm.)

8.4 Food Processing Industries Food is a basic necessity for life. Therefore, food processing is an important procedure in which food is prepared for consumption. The food processing industry tends to specialize in the areas of food preparation and food packaging. Some processing plants carry out both tasks. For example, they turn raw ingredients into prepared food and then package it for shipment to stores, etc. They use preservatives to extend the shelf-life of many food items. Also in order to promote food safety, regulatory agencies (like the Food and Drug Administration: FDA) have standards for preparing food, testing for contamination, workplace hygiene, etc. Figure 8.19 is a display of processed food in a supermarket [34]. Biofilms impact the food processing industries. They can exist on all types of surfaces in food plants. Some surface examples include plastic, glass, metal, food

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Fig. 8.19 Processed food in a supermarket

products, etc. If bacteria attach to the food product or to surfaces that come in contact with the product, then biofilm formation is promoted and the result could be economic loss (due to food spoilage) and hygienic problems. Contamination of food and pathogenic microorganisms cost the food industry millions of dollars each year [35]. One of the main problems in the food industry is the survival of foodborne pathogens due to insufficient cleaning of surfaces and/or instruments that come in contact with the food. Food contact surfaces must be clean and sanitized to avoid biofilm formation, contamination of the final product, and food poisoning. Some pathogens that cause problems for the food industry include Listeria monocytogenes, Salmonella, Escherichia coli, Pseudomonas, Campylobacter jejuni, and Bacillus. Listeria can create biofilm in slicers and other steel utensils and contribute to cross contamination [36]. It has also appeared in milk and dairy products. Studies show that Salmonella can attach to and form biofilms on plastic, cement, and stainless steel [37]. Pseudomonas are found on drains, floors, fruits, vegetables, and meat surfaces [38]. They can form biofilms on stainless steel surfaces and coexist within biofilms with other pathogens. Bacillus is found in dairy processing plants. Campylobacter can produce biofilms on stainless steel and glass surfaces. It is Gramnegative bacteria with s-shaped rods and often found in poultry. C. jejuni is one of the main causes of bacterial foodborne diseases. See Fig. 8.20 [39]. Food processing industries must have clean and sanitary apparatus in order to provide safe food items for consumers. They are wise in using stainless steel for most equipment because it is durable, corrosion resistant, and has other good qualities. However, they need an effective and reliable method to prevent and remove biofilm.

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Fig. 8.20 Campylobacter jejuni (SEM image above) is a main cause of gastrointestinal disease

GENERAL METHOD TO PREVENT AND REMOVE BIOFILM 1. Develop, implement, and stick to a good cleaning and sanitizing program. 2. Use well-designed and properly installed processing equipment. Note: Biofilms occur in curves and joints of equipment and are difficult to remove. Keep in mind that surface characteristics such as roughness promote biofilm formation. Therefore, roughness is measured by the R factor. R factor limits should be considered when purchasing equipment. 3. Cleaning is very important because it prepares the surface for sanitizing. Mechanical methods for removing biofilms include scraping, and the use of scrubbers, brushes, and various cleaning tools. If these methods don’t work, then detergents and chemical treatments are required. For example, one study showed that a hydrogen peroxide/peroxyacetic acid-based compound has the ability to penetrate biofilms, so they can easily be removed [40]. A promising alternative treatment for killing bacterial spores and pathogens (without the use of toxic chemicals) is the electrostatic space charge system (ESCS). The United States Department of Agriculture (USDA) used the ESCS against bacterial spores of Bacillus stearothermophilus and reduced them by 99.8%. The USDA also treated

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the pathogens Campylobacter jejuni, Listeria monocytogenes, Salmonella enteritidis, Staphylococcus aureus, and Escherichia coli, and achieved up to 99.9% reduction efficiency [41]. The problem of biofilms is not going away because the food industry is moving toward longer processing runs with a minimum number of cleanings between runs.

8.5 Pipes and Heat Exchangers Biofilms contribute to slime and scale buildup in clogged pipes and heat exchangers. These clogged items (which have a smaller diameter) display a reduced ability for liquid flow and heat exchange. Companies with such problems could experience increased operational costs and energy consumption. Also if these situations are not corrected in time, the industries may be shut down for a long period in order to clean, repair, or replace the defective pipes. Biofilms also cause corrosion in metal pipes and are potential health hazards if they contain pathogenic bacteria. Therefore, it is essential to have well-designed equipment and to test it regularly for the presence of biofilms, chemicals, and biological content. It is also important to clean the system and apparatus frequently using physical and chemical treatments as needed. A heat exchanger is a system for transferring heat between two or more fluids. It can be used in both heating and cooling processes. The fluids may be separated by a solid wall or in direct contact like water and air (as exhibited in a cooling tower). A double pipe heat exchanger is the simplest type used in industry. See Fig. 8.21 [42]. The two fluids can enter the exchanger at the same end and travel in parallel or they can enter from opposite ends (which is a more efficient way). A cooling water system is used to remove unwanted heat. A simple type is an open re-circulating system. It generally includes pumps, heat exchangers, and a cooling tower. The pumps keep the water re-circulating through heat exchangers. After the cooling water picks up heat, it moves to the cooling tower where heat is released by the process of evaporation. See Fig. 8.22 [43]. The leftover water collects at the base of the cooling tower for reuse. There is an intake of water to make up for the loss of water through evaporation. Biofilms are problematic for cooling water systems. They develop on the inside of metal pipes and cause clogging and corrosion. Clogging reduces a pipe’s ability to exchange heat. The biofilm’s layers form barriers between re-circulating cooling water and the surfaces of the heat exchangers. Also some biofilms, like those with anaerobic sulfate-reducing bacteria, cause corrosion [44]. In addition, biofilms containing pathogenic bacteria can exist in cooling tower systems and cause health concerns. Biofilms appear slimy on water system surfaces and can be detected by wet chemistry tests and online monitoring devices like biofouling sensors. Also one can assess microbial levels in water systems by passing a portion of the water through a membrane filter to capture any colonies present so they can be tested and evaluated [45].

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Fig. 8.21 Simple tubular heat exchanger

Fig. 8.22 Above is a cooling tower where heat is released by the process of evaporation

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Biofilms can negatively affect the quality of drinking water too. After waste water is treated, it flows through clean pipes before arriving at our faucets. Sometimes biofilms form in the pipes that carry clean water. This might cause the clean water to become contaminated. Scientists found that by removing organic carbon (nutrition for bacteria) from processed water, the amount of biofilm formed can be limited. WATER DISTRIBUTION SYSTEMS eventually develop biofilms. These selfproduced biofilms protect bacteria (that formed them) including pathogenic types that are potential threats to public health. One possible concern is Legionella, small Gramnegative, rod-shaped bacteria that cause Legionnaires’ disease [46]. See Fig. 8.23 [47]. Symptoms for this disease resemble those for pneumonias. Biofilms can also discolor water and cause taste and odor problems. They can clog and corrode metal pipes too. Non ferrous pipes like PVC (polyvinyl chloride) do not corrode. Biofim development depends on many variables such as temperature, nutrient levels, stagnant periods, pipe materials, etc. Chemical and physical options are available for controlling biofilms, especially in pipes and aging drinking water systems. Companies should reduce nutrient levels (bacterial food) in the water to help decrease the growth of biofilm and paint/coat metal to protect it from corrosion. Also they can disinfect the water. Chemicals are generally used for microbial control. For example, biodispersants help detach microbial deposits from metal surfaces. Chemical cleaning for biofilms depends on the actions of detergent ingredients like alkaline constituents, oxidizing agents, surface active agents, etc. [48].

Fig. 8.23 Transmission electron microscope image of Legionella pneumophila, bacteria responsible for most cases of Legionnaires’ disease

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Investigators are interested in biofilms on pipe walls in water distribution systems. Several made field observations of biofilm using biofilm potential monitors. The monitor results were compared with pipe samples taken from the distribution system and with laboratory pipe reactors. They found that water having total organic carbon concentrations in the range 1.5–3.9 mg/l, a free chlorine residual of 0.2 mg/l was needed to reduce biofilm concentration to below 50 pg ATP/cm2 [49]. (This represents 50 pg of adenosine triphosphate, ATP, the essential energy molecule existing in all organisms. One can quickly judge whether living organisms or biofilms are present by testing the amount of ATP.) Other researchers recommend the use of an enzyme cleaning technique to eliminate the formation of biofilm in industrial pipes. They mention that the traditional cleaning in place (CIP) technique for the water treatment process usually removes or sterilizes microbes on the surface of pipes. However, the method does not work well when biofilms form in pipelines. The scientists carried out a study that compared the two methods: the CIP and the enzyme approach. They found that using muramidase to remove biofilm in pipes is more effective and in-depth than CIP [50]. NOTE: Muramidase is an enzyme (present in tears) that destroys the cell walls of certain bacteria. Some physical cleaning methods to remove biofilm from pipes include flushing water through pipes at high velocities, scaling (like that to remove plaque on teeth), pigging, and the use of brushes, scrappers, and swabs. A swab is a compressible foam cylinder that is forced through a pipe using water pressure. Pigging is actually a technique that uses a pig (a bullet-shaped cylinder) to clean pipelines. Pigs, which tend to be harder and less flexible than swabs, are blown through a pipe. See Fig. 8.24 [51]. The pig fits tightly against the interior wall of the pipe and scrapes the edges to remove biofim and debris. Brushes and blades are sometimes added to a pig to improve its effectiveness for cleaning. This method requires launching and retrieval of the pig.

8.6 Hospital and Medical Fronts Biofilms are the cause of many medical problems. They are produced by bacteria and encase them to protect the bacteria from antibiotics, etc. Biofilms form at the interface of different phases by bacterial activity. From the viewpoint of structure, bacteria at the interface are surrounded by sticky, external polymeric substances (EPS). It should be mentioned that bacteria in biofilms are different from those outside of the biofilm. Bacteria outside of the biofilm are called planktonic bacteria. The details of these bacteria have already been described in a previous chapter (Chap. 3). However, we would like to mention some important characteristics to remind you of the typical differences. When bacteria exist in oligotrophic (nutrient-poor) environments, their average size in biofilms is generally smaller than those in nutrient environments. Therefore, we can say that the bacteria shrink in the biofilms. After biofilms grow for a certain

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Fig. 8.24 Above is a pig within a cut out section of pipe

amount of time, some of them rupture, due to the balance between changes of EPS and shear stress on the interface. Eventually the bacteria in biofilms become bigger and the average size is like that of the original planktonic type, as shown in Fig. 8.25. Bacteria generally shrink their sizes in oligotrophic environments. Since biofilms are

Fig. 8.25 Size differences between bacteria in biofilms and the planktonic type

196 Table 8.1 Several common examples for biofilm-induced chronic diseases/infections

8 Biofilm Problems and Environments Chronic diseases/infections 1. Sinusitis 2. Otitus media 3. Urinary tract infection 4. Periodontitis 5. Rosacea

Possible bacteria responsible for the condition 1. Streptococcus pneumonia 2. Streptococcus pneumonia 3. Escherichia coli 4. Periodontal bacteria and Streptococcus mutans 5. Helicobacter pylori and Bacillus oleronius

a sort of oligotrophic environment, it is very natural for bacteria to shrink in biofilms [52–54]. We have also observed the shrinking, using FIB-SEM [55]. The shrinking of bacterial size in biofilms means that the bacteria in biofilms would not need so much nutrition. In addition to that, the rate of cell division would generally slow down in the biofilms. From this view point, we could say that bacteria in biofilms would lower their life activities. Such a situation is called being “dormant” [56–60]. Therefore, the bacteria in the biofilms would be saving energy. People often say that biofilms provide a way for bacteria to survive in oligotrophic environments. In regards to medical science, one can say that bacteria in biofilms have a high drug tolerance (drug resistance). Inside our bodies, bacteria in biofilms have high resistance to antibiotics, while those outside of the body also have high resistance to disinfectants and biocides. When we come to think about our bodies and medical fronts, these characteristics would lead to chronic diseases. Actually, many doctors insist that most of the chronic diseases could be attributed to biofilms. Table 8.1 (prepared by the authors) shows a few concrete examples for chronic diseases/infections caused by biofilms. Costerton et al. wrote a paper that relates biofilms to diseases [61]. In particular, a serious problem caused by biofilms in hospitals is biomaterial associated infections like those on stents, hip replacement parts, etc. It is hard to establish the countermeasure for this situation and to solve the problem. One needs to control biofilms on biomaterials. Many diseases are caused by and closely related to biofilms [62]. To solve these problems, many countermeasures have already been proposed. The biofilm problem should be solved not only from the environmental viewpoint, but also from that of materials science, since biofilms are one of the interfacial problems. In light of that, the approach and effort from the area of materials science has not been enough. There have been many versatile countermeasures so far. However, most of them are pharmaceutical measures. Such an approach from materials science and interactions between biofilms and materials is now expected.

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38. Chmielewsky, R. A. N., & Frank, J. F. (2003). Biofilm formation and control in food processing facilities. Comprehensive Reviews in Food Science and Food Safety, 2, 22–32. 39. De Wood, P. File: ARS Campylobacter jejuni.jpg. Date: January 2, 2008. License: This image is in the public domain. It is the work of the United States Department of Agriculture. https:// commons.wikimedia.org/wiki/File:ARS_Campylobacter_jejuni.jpg. 40. Humm, B. J. (1992). A research update on the effects of cleaners and sanitizers on food processing biofilms. Food Protection Report, 8, 5–6. 41. United States Department of Agriculture: Research, Education, & Economics Information System. https://portal.nifa.usda.gov/web/crisprojectpages/0404877-reduction-and-control-ofpathogens-associated-with-food-processing-surfaces.html. 42. Koen B. File: Tubular heat exchanger.png. Date: April 7, 2007. License: This work is in the public domain. https://commons.wikimedia.org/wiki/File:Tubular_heat_exchanger.png. 43. Malfoy. File: Cooling tower power station Dresden. Jpg. Date: June 29, 2007. License: Creative Commons Attribution-Share Alike 3.0 https://commons.wikimedia.org/wiki/File: Cooling_tower_power_station_Dresden.jpg. 44. Earthman, J. C., & Wood, T. K. (1997). Corrosion inhibition by aerobic biofilms on SAE 1018 steel. Applied Microbial Biotechnology, 47, 62–68. 45. Sandle, T. (2017). The problem of biofilms and pharmaceutical water systems. American Pharmaceutical Review. https://www.americanpharmaceuticalreview.com/Featured-Articles/ 345440-The-Problem-of-Biofilms-and-Pharmaceutical-Water-Systems/. 46. United States Environmental Protection Agency, Office of Water. (2000, September). Legionella: Drinking water fact sheet. EPA. https://www.epa.gov/sites/production/files/201510/documents/legionella-factsheet.pdf. 47. CDC Public Health Image Library #1187. File: Legionella pneumophila 01.jpg. Date: June 5, 2006. License: This work is in the public domain. The image is the work of the Centers for Disease Control and Prevention, part of the U.S. Department of Health and Human Services. https://commons.wikimedia.org/wiki/File:Legionella_pneumophila_01.jpg. 48. Fukuzaki, S. (2015). Chemical cleaning. In H. Kanematsu & D. M. Barry (Eds.), Biofilm and materials science (pp. 155–162). Switzerland: Springer. 49. Hallam, N. B., West, J. R., Forster, C. F., & Simms, J. (2001). The potential for biofilm growth in water distribution systems. Water Research, 35(17), 4063–4071. https://doi.org/10.1016/ S0043-1354(01)00248-2. 50. Liu, X., Tang, B., Gu, Q., & Yu, X. (2014). Elimination of the formation of biofilm in industrial pipes using enzyme cleaning technique. MethodsX, 1, 130–136. https://doi.org/10.1016/j.mex. 2014.08.008. 51. Barrison, H. File: Pipeline PIG.jpg. Date: February 24, 2009. License: Creative Commons Attribution-Share Alike 2.0 Generic https://commons.wikimedia.org/wiki/File:PipelinePIG. jpg. 52. Colwell, R. R., & Grimes, D. J. (2000). Nonculturable microorganisms in the environment: ASM Press. 53. Kieft, T. L. (2000). Size matters: Dwarf cells in soil and subsurface terrestrial environments. In Nonculturable microorganisms in the environment (pp. 19–46). Springer. 54. Byrd, J. J. (2000). Morphological changes leading to the nonculturable state. In Nonculturable microorganisms in the environment (pp. 7–18). Springer. 55. Kim, J., Hahn, J.-S., Franklin, M. J., Stewart, P. S., & Yoon, J. (2008). Tolerance of dormant and active cells in Pseudomonas aeruginosa PA01 biofilm to antimicrobial agents. Journal of Antimicrobial Chemotherapy, 63(1), 129–135. 56. García-Contreras, R., Zhang, X.-S., Kim, Y., & Wood, T. K. (2008). Protein translation and cell death: The role of rare tRNAs in biofilm formation and in activating dormant phage killer genes. PLoS ONE, 3(6), e2394. 57. Chihara, K., Matsumoto, S., Kagawa, Y., & Tsuneda, S. (2015). Mathematical modeling of dormant cell formation in growing biofilm. Frontiers in Microbiology, 6, 534. 58. Kim, J., Park, H.-J., Lee, J.-H., Hahn, J.-S., Gu, M. B., & Yoon, J. (2009). Differential effect of chlorine on the oxidative stress generation in dormant and active cells within colony biofilm. Water Research, 43(20), 5252–5259.

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59. Burkharta, C. N., Burkhart, C. G., & Gupta, A. K. (2002). Dermatophytoma: Recalcitrance to treatment because of existence of fungal biofilm. Journal of the American Academy of Dermatology, 47(4), 629–631. 60. Olsen, I. (2015). Biofilm-specific antibiotic tolerance and resistance. European Journal of Clinical Microbiology and Infectious Diseases, 34(5), 877–886. 61. Costerton, J. W., Cheng, K. J., Geesey, G. G., Ladd, T. I., Nickel, J. C., Dasgupta, M., et al. (1987). Bacterial biofilms in nature and disease: Annual review of. Microbiology, 41(1), 435– 464. 62. Jamal, M., Tasneem, U., Hussain, T., & Andleeb, S. (2015). Bacterial biofilm: Its composition, formation and role in human infections. Journal of Microbiology and Biotechnology, 4, 1–14.

Chapter 9

Biofilm Usefulness

Abstract This chapter describes the usefulness of biofilms. It starts by discussing the importance of biofilms for producing electricity and the process which is known as microbial fuel cell (MFC) technology. The next topic presents beneficial applications of biofilms to the environment. It includes bioremediation, nitrogen-fixing bacteria that make atmospheric nitrogen available to plants, and biofilm/bacteria use for recycling elements vital to life. Biofilms are also used to immobilize harmful materials, as biological pesticides, and for bioleaching to extract metals from their ores. The final section provides a description of how biofilms contribute to water treatment applications for pollutants such as plastics in our oceans, heavy metals, industrial wastes, oil spills, and sewage.

9.1 Energy Applications Biofilms are useful for producing energy in the form of electricity. This process, which relies on bacteria to make electricity, is called microbial fuel cell (MFC) technology. Conventional fuel cells generally depend on hydrogen gas for the fuel source, while the microbial fuel cell uses various water-based organic fuels such as sewage. A fuel cell is an electrochemical cell that converts the chemical energy of a fuel into electricity through redox reactions. Fuel cells can produce electricity continuously for as long as fuel and oxygen are supplied. They contain an anode, cathode, an electrolyte, and an external circuit. At the anode a catalyst causes the fuel to undergo oxidation reactions that generate ions and electrons. The ions move from the anode to the cathode through the electrolyte. At the same time, electrons flow from the anode to the cathode through an external circuit, producing electricity. At the cathode, ions and oxygen combine to form water. A typical fuel cell diagram is displayed in Fig. 9.1 [1]. The microbial fuel cell (MFC) uses various, water-based organic fuels such as sewage. A lot of biomass (waste) is available. It provides nutrition for bacteria, which in turn convert it to energy. A variety of bacteria exist, so researchers are able to find a bacterium that can handle almost any waste compound. Bacteria can © Springer Nature Singapore Pte Ltd. 2020 H. Kanematsu and D. M. Barry, Formation and Control of Biofilm in Various Environments, https://doi.org/10.1007/978-981-15-2240-6_9

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Fig. 9.1 Above is a typical display of a fuel cell that produces an electric current. The fuel is hydrogen, the source of oxygen is the air, and water is a product

provide a cheap source of electricity. There are many types of MFCs throughout the world, but they basically have the same operating system. They have a pair of battery-like terminals: an anode and cathode electrode. The electrodes are connected by an external circuit and an electrolyte solution to help conduct electricity. To be more specific, an anode respiring bacterium breaks down the organic waste to carbon dioxide and transfers the released electrons to the anode. Then, the electrons travel from the anode through an external circuit to generate electrical energy. The electrons complete the circuit by traveling to the cathode. Here, ions and oxygen form water. (NOTE: Anode respiring bacteria are able to transfer electrons in organic substrates to a solid electrode.) You may wonder how bacteria get electrons to the anode. The bacteria form a biofilm on the anode’s surface. The sticky biofilm matrix is rich with material that can potentially transport electrons. This possibility allows scientists to consider that the biofilm is acting as an anode. Previous studies showed that the rate of bacterial metabolism at the anode increases when the electrical potential of the anode increases. Therefore, a team of researchers determined that the electrical potential is equal to the concentration of electrons and it is the electrons that the bacteria transfer to the anode [2]. This team also found that the biofilm produced more current when the biofilm is of medium thickness. Other researchers coated individual bacterial cells with an electron-conducting polymer to provide a high-performance anode for microbial fuel cell applications. They relied on the polymer polypyrrole to improve the electrical conductivity of bacterial cells without reducing their viability. The scientists used iron ions as the oxidative initiator to make pyrrole monomers polymerize on the bacterial surface. The organism used was proteobacterium Shewanella oneidenis, which tolerates metal and both aerobic and anaerobic lifestyles. The coated bacteria were tested for biocurrent

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generation with a carbon anode. Compared to their unmodified counterparts, they displayed a fivefold increase in electricity generation [3]. Microbial fuel cells belong to either the mediated or unmediated group. The mediated group uses a mediator, a chemical that transfers electrons from the bacteria in the cell to the anode. Unmediated MFCs typically have electrochemically active bacteria with redox proteins on their outer membrane that can directly transfer electrons to the anode. A special type of MFC is soil-based. In this case, soil acts as the nutrient-rich anodic media. The anode is placed at a certain depth within the soil, while the cathode is on top of the soil and exposed to air. The soil includes various microbes, including bacteria needed for MFCs, and nutrients from plant decay, etc. A soil-based MFC is displayed in Fig. 9.2 [4]. An example of an unmediated (mediated-free) MFC is the plant microbial fuel cell. This fuel cell can run on wastewater and get energy directly from certain plants. A diagram for the plant microbial fuel cell can be seen in Fig. 9.3 [5]. MFCs are attractive for power generation, especially since most organic material can be used as nutrition for the fuel cell. They can be used for applications that require only low power. Also they can work at a small scale. Electrodes in some Fig. 9.2 A soil-based microbial fuel cell

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Fig. 9.3 Plant microbial fuel cell

cases need only be 7 µm thick by 2 cm long [6]. This allows an MFC to replace a battery. It provides a renewable form of energy and does not need to be recharged. MFCs operate well in mild conditions with temperatures of 20–40 °C and at a pH of about 7 [7]. Higher power production has been observed too, with a biofilm-covered graphite anode [8, 9]. Another biofilm application for energy is the microbial electrolysis cell (MEC) that was introduced in 2005 [10]. The MEC requires an external source of electricity for electrolysis to produce hydrogen at the cathode. However, this external energy supply is small since most of the energy comes from the chemical energy of substrates oxidized at the anode [11]. Therefore, hydrogen can be produced by using a small amount of energy. MEC is of special interest because it produces hydrogen that is needed for the upcoming hydrogen-energy economy [12, 13]. Hydrogen can serve as an alternative to fossil fuels in internal combustion engines and for power generation. It is also used for rocket fuel, refining petroleum, treating metals, and processing food. Figure 9.4 shows a microbial electrolysis cell [14]. Biofilms also have potential for producing precursors of desired liquid fuels through the process of microbial electrosynthesis. An option is to reduce carbon dioxide to produce multicarbon organic compounds that are precursors for desirable liquid transportation fuels. In one study, researchers found that biofilms of Sporomusa

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Fig. 9.4 A diagram of a microbial electrolysis cell is displayed

ovate growing on graphite cathode surfaces consumed electrons with the reduction of carbon dioxide to acetate and small amounts of 2-oxobutyrate [15].

9.2 Environmental Applications Microbes make up most of the living biomass on Earth and help recycle elements vital to life. They often live in colonies of biofilms on surfaces. Biofilms and bacteria play important roles in the environment. They decompose dead and complex organic matter. Bacteria use this organic matter as a source of nutrients. In turn, they recycle organic compounds trapped in dead matter so other organisms can benefit from them. Bacteria make nitrogen in the atmosphere (an important element for plants) available to plants through the process of nitrogen fixation. Figure 9.5 shows the nitrogen cycle [16]. The process of nitrogen fixation takes place with the help of nitrogen fixing bacteria like Rhizobium and Cyanobacteria. A diagram of the cyanobacterium is provided in Fig. 9.6 [17]. These bacteria change atmospheric nitrogen into ammonium, nitrites, and nitrates. The nitrates are the predominant form absorbed by the roots of plants in the soil. The plants convert the nitrates to proteins. Some leguminous plants are able to have a mutualistic association with the bacteria living in their tissues. The bacteria live in nodules (small growths on the roots) and carry out nitrogen fixation. On the other hand, the plant supplies nutrients and energy to the bacteria. BACTERIA ARE KEY PLAYERS IN BIOREMEDIATION. This is the process of depleting/degrading toxic compounds in the natural environment [18]. Bacteria like Pseudomonas have been well-known for degrading oil spills on soils and oceans. Contamination, caused by toxic heavy metals in the environment, can be controlled to some extent by bacteria. Bacterial cell walls play an important role in accumulating heavy metals by bacterial cells. Pseudomonas fluorescens and Enterobacter cloacae

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Fig. 9.5 A diagram of the nitrogen cycle is displayed

Fig. 9.6 This is a detailed diagram for cyanobacterium

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are able to uptake heavy metals such as arsenic and cadmium [19, 20]. Figure 9.7 shows colony growth of Enterobacter cloacae bacteria on Tryptic Soy Broth agar [21]. This is a Gram-negative, rod shaped bacterium. BACTERIA IMMOBILIZE HARMFUL MATERIALS. Nanowires grown by certain types of bacteria can immobilize uranium and other harmful materials. These bacteria can be brought into uranium contamination sites (like nuclear plants) to contain the radiation. A research team at Michigan State University learned that Geobacter bacteria (found naturally in soil) essentially electroplates uranium, causing it to be insoluble so it can’t contaminate groundwater. This team, led by Dr. Gemma Reguera, demonstrated that the long arms (known as pili) of Geobacter give them the ability to connect to uranium particles and yank them out of water. These arms can be compared to wires, since they are able to conduct electricity [22]. THE BACTERIA Bacillus thuringiensis (Bt) IS COMMONLY USED AS A BIOLOGICAL PESTICIDE. Bt is a Gram-positive soil dwelling bacterium. During the

Fig. 9.7 Rough and smooth colony growth of Enterobacter cloacae bacteria on Tryptic Soy Broth agar

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formation of spores, many Bt strains produce crystal proteins that express insecticidal properties. These proteins are toxic to the digestive system of insects. Therefore, the toxin has been used in pesticides to control pests such as various types of worms, etc. [23]. It is incorporated into dusting powder applied to plants that insects eat. Also the toxin gene has been incorporated into plants to make them insect resistant. Insects spread disease and cause devastating damage to crops. Therefore, insect control is important for agriculture and to prevent human disease. Using microbial insect control is safer than the use of chemicals. Figure 9.8 shows crystals of Bt-toxin from Bacillus thuringiensis [24]. BIOFILMS SUPPORT THE PROCESS CALLED BIOLEACHING [25, 26]. This process uses bacterial microorganisms to extract metals from their ores. The bacteria feed on nutrients in the minerals, causing metals to separate from their ores. Examples of these metals include gold, silver, zinc, and copper. Bioleaching is generally carried out by iron and sulfide oxidizing bacteria. Iron ions (Fe+3 ) are used to oxidize the ore. The role of the bacteria is the further oxidation of the ore and the regeneration of the chemical oxidant Fe+3 from Fe+2 . Some bacteria used in this process include Leptospirillum ferrooxidans, Thiobacillus ferrooxidans, etc. Using microorganisms is environmentally friendly and cheaper than chemical extractions. Some bacteria can clean pollution, while others generate electricity. However, the bacteria Desulfitobacterium can simultaneously breakdown pollutants like printed circuit boards and produce electricity [27]. This bacterium is a Gram-positive, anaerobic, rod-shaped, spore forming bacteria.

Fig. 9.8 Crystals of Bt-toxin from Bacillus thuringiensis

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BACTERIA (TU-103) CONVERT NEWSPAPER TO BIOFUEL. In 2011, Tulane University’s alternative fuel research scientists discovered a strain of Clostridium called TU-103. It can change nearly any form of cellulose into butanol, a fuel that may be used in an internal combustion engine [28]. Actually a strain of this type of bacteria (called Clostridium acetobutylicum) was used in 1916 to jointly produce acetone, ethanol, and butanol from starch. This bacterium can digest cellulose too (as previously mentioned). The bacterium is sometimes called Weizmann Organism, after a lecturer at the University of Manchester in England, who used it in 1916. NOTE: Cellulose is an organic compound with the formula (C6 H10 O5 )n . It is the structural component of the primary wall of green plants and is mainly used to make paperboard and paper. BIOSENSORS: Bacteria are also useful as biosensors [29]. This is because they are able to detect pollutants and pathogens. Each year industries generate tons of waste which ends up in landfills. Therefore, scientists are using biosensors (bacteria) that can locate biologically active pollutants. Bacterial biosensors have a receptor that is activated in the presence of pollutants and a reporter that records changes. They use the lux operon from Aliivibrio or Phototbactrium as a reporter. The lux operon includes an inducer and structural genes for the enzyme luciferase, which produces bioluminescence similar to that seen in fireflies. Simply stated, a bacterium containing the lux operon will emit visible light when the receptor is activated. The lux operon can be transferred to many bacteria.

9.3 Water Treatment Applications Water is an important resource that we need for survival. Almost three-fourths of the Earth is covered with water. Humans and other forms of life are primarily made of water. Water is needed to produce food and keep plants alive. Water is sometimes called the universal solvent because it can dissolve a number of certain compounds. Therefore, it is good for cleaning surfaces, removing wastes, and transporting dissolved nutrients into the tissues of our bodies. Water also plays a key role in controlling climate. Some places get too much water and have floods, while others do not get enough water and experience droughts. The planet’s water supply is constantly recycled and purified by the water cycle, whenever it is not disrupted by human activities. The water cycle (also known as the hydrological cycle) describes the continuous movement of water on, above, and below the surface of the Earth. It is driven by solar energy and leads to temperature changes. When water evaporates, it takes up energy from its surroundings and cools the environment. When it condenses, it releases energy and warms the environment. The water cycle includes three main processes. One is called evaporation, which is the conversion of liquid water in oceans, rivers, etc. into water vapor in the atmosphere. Another is transpiration, the evaporation of water from leaves of plants into the atmosphere. The third one is precipitation. This is when water returns from the

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Fig. 9.9 The water cycle

atmosphere (in the form of snow, rain, etc.) to the Earth’s surface. (NOTE: Sublimation is when a substance directly changes from a solid to a gas.) Precipitation that falls on land and remains on its surface is called surface water. Surface water that seeps into underground bodies of water stored in layers of rock, etc. is called groundwater. Figures 9.9 and 9.10 provide detailed diagrams of the water cycle [30, 31]. Water is a very important resource for the global community. Therefore, it is essential to maintain a high quality water supply, free of pollutants. Water pollution is any change in water quality that causes harm to humans and other living organisms that use the water. Polluted water is also defined as being unsuitable for swimming, fishing, and boating. Every year thousands of people get sick from drinking polluted well water and from swimming in water that contains disease-causing organisms. Various forms of water pollutants exist like sewage, oil spills and an abundance of plastics in our oceans, heavy metals, etc. Biofilms are important for water treatment applications. SEWAGE TREATMENT (also called wastewater treatment) is the method for removing contaminants from municipal wastewater to produce treated effluent that is safe to release into the environment. Wastewater from homes, businesses, etc. flows through sewer pipes to sewage treatment plants. It generally passes through several levels of treatment. The first process involves a settling tank, where most solids and sediments are removed. The next step is a biological process in which bacteria remove as much as 90% of degradable, oxygen-demanding organic wastes. Many treatment systems include a disinfection process that uses chlorine to kill bacteria.

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Fig. 9.10 Another view of the water cycle that contains the transpiration process

In rural areas, homeowners have on-site septic systems. Household sewage and wastewater is pumped into a settling tank where bacteria decompose the solid wastes. This partially treated wastewater seeps from the tank to a drainage field made of gravel or crushed stone located just below the topsoil. Then it percolates through the soil beneath the gravel, where most of the potential groundwater pollutants are filtered out. Also the soil bacteria present decompose other degradable materials. To be more specific, sewage treatment generally involves three stages: the primary, secondary, and tertiary stages. During the primary stage, sewage is held in a basin where heavy solids can settle to the bottom, while lighter ones like grease and oil float on the surface. The floating materials are removed by using screens and skimmers. After the settled and floating items are removed, then the remaining liquid is subjected to secondary treatment. NOTE: Sewage solids collected on the bottom of the settling tank are called sludge, which is pumped to sludge treatment facilities. Secondary treatment is designed to substantially degrade the sewage’s biological content derived from human waste, food waste, etc. This stage is used to remove the organic matter and reduce the BOD (biochemical oxygen demand, a measure of the biologically degradable organic matter in water). BOD is determined by the amount of oxygen that bacteria need to metabolize organic matter. During secondary treatment, the sewage undergoes aeration to enhance the growth of aerobic bacteria and other microorganisms that oxidize the dissolved organic matter to carbon dioxide and water. Two methods for this stage of treatment include activated sludge systems

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and trickling filters. Activated sludge contains large numbers of sewage-metabolizing microbes, which oxidize much of the sewage’s organic matter. An important microbe is Zoogloea bacteria that form floc, which is allowed to settle out in a settling tank. Zoogloea is a genus of gram-negative, aerobic, rod-shaped bacteria. Figure 9.11 displays floc formed by this bacterium [32]. Sometimes the sludge floc floats instead of settling out. This is called bulking and is caused by filamentous bacteria like Sphaerotilus natans and Nocardia species. Trickling filters are used for secondary treatment. Here sewage is sprayed over a bed of rocks or molded plastic that is aerated. A biofilm of aerobic microbes grows on the rock or plastic surfaces and oxidizes the organic matter trickling over the surfaces to carbon dioxide and water. Another biofilm-based design for sewage treatment is the rotational biological contactor system. This system allows biomass to grow on various media and then sewage passes over these surfaces. For further treatment, sludge is pumped to anaerobic sludge digesters, which encourage the growth of anaerobic bacteria, especially methane-producing bacteria. These bacteria decrease the organic solids by degrading them to soluble substances and gases, most of which are methane and some carbon dioxide. Finally, the sludge can be used for landfill or

Fig. 9.11 This photo shows floc (in the container on the left) formed by Zoogloea resiniphila

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Fig. 9.12 This diagram shows a simple flow process for a typical large-scale treatment plant

as a soil conditioner. This is possible when the number of pathogens present is below a certain number. Tertiary sewage treatment is designed to essentially remove all BOD, nitrogen, and phosphorus. Some systems use denitrifying bacteria to form volatile nitrogen gas. Finally the purified water is chlorinated. This water is suitable for drinking. Figure 9.12 shows a simple diagram of the flow process for a typical large-scale treatment plant [33]. Figure 9.13 is of a typical treatment plant using subsurface constructed wetlands [34]. It is interesting to add that runoff sewage and farm wastes in coastal waters generally contain large quantities of nitrates and phosphates. These nutrients can result in large algae populations. Harmful algae are sometimes called brown, green, or red toxic tides. They release poison that kills some fish-eating birds as well as other animals (including dogs) that swallow the water while swimming. They also poison many humans who eat shellfish contaminated by the algae. The good news is that oxygen-using bacteria are able to decompose algae. However, this depletes the water of dissolved oxygen. OIL SPILLS are a form of pollution. They are usually the release of liquid petroleum hydrocarbons into the environment (especially the sea), as a result of human activity. Oil pollution comes from many sources like oil tanker spills and leaks, ruptures from offshore drilling rigs, etc. Spilled oil adversely affects plants and animals. Volatile organic hydrocarbons in oil kill many aquatic organisms. Other chemicals in oil form tarlike globs that coat the feathers of swimming birds and the fur of marine mammals. The oil reduces their insulating ability and makes them less

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Fig. 9.13 Displayed is a typical treatment plant using subsurface constructed wetlands

buoyant in the water. This tarlike material causes many of these birds and mammals to drown or die from loss of body heat. Figure 9.14 shows a bird covered in oil, as a result of a spill in the Black Sea [35]. Also some components of oil sink to the ocean floor and smother crabs, oysters, clams, and more. Oil consuming bacteria are used to remove oil from an ecosystem. These bacteria rely on oil for energy. One such rod-shaped bacterium (without flagella) is Alcanivorax borkumenis [36, 37]. It is an alkane-degrading marine bacterium, which becomes predominant in seawater containing crude-oil when nitrogen and phosphorus nutrients are available. NOTE: An alkane is a saturated hydrocarbon composed of single bonds and the elements hydrogen and carbon. The genome of A. borkumenis is a singular, circular chromosome containing many base pairs. A certain sequence on the genome codes for the degradation of a certain range of alkanes. It is interesting to note that this bacterium can’t consume sugars or amino acids as a source of energy. This is because it lacks the required genes to do so [38]. Its genome also contains instructions for the formation of biosurfactants that add in the process of degradation. The genome also codes for several defensive mechanisms [39]. Cycloclasticus pugetii is an aerobic, gram-negative, rod-shaped bacterium that degrades aromatic hydrocarbons like naphthalenes [40]. Researchers confirmed that the addition of nitrogen and phosphorus fertilizers enhanced the biodegradation of spilled oil and the growth of Cycloclasticus strains. Their investigation used beach simulating tanks that mimicked an oil-polluted beach. They added natural seawater to both tanks, but nitrogen and phosphorus fertilizers were only added to one tank. The tank with the added fertilizers displayed more biodegradation of oil and more growth of the Cycloclasticus bacterium.

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Fig. 9.14 A bird covered with oil

Oceanospirillales bacteria get their energy from the breakdown of various organic products, those containing carbon [41]. The bacteria are an order of Proteobacteria and prefer high salt concentrations to grow in their marine environment. They are displayed in Fig. 9.15 [42]. PLASTIC POLLUTION is the accumulation of plastic objects in the Earth’s environment that causes potential harm to humans and wildlife. It can affect land, waterways, and oceans. Marine animals can become entangled in plastic objects, ingest plastic waste, and be exposed to toxic chemicals within the plastics. To somewhat control the over abundance of plastics, humans reduce, reuse, and recycle them. Figure 9.16 is an example of plastic pollution in Ghana [43]. Plastics take a long time to degrade. Therefore, plastic eating bacteria would be helpful for this task. Plastics are used worldwide and are made from a variety of structural polymers. Therefore, important research is being carried out to obtain methods for plastic biodegradation. One example is provided about researchers in Japan. Scientists in Japan discovered a strange species of bacteria in samples of soil near a bottle recycling facility in Sakai, Japan. These bacteria were able to breakdown and metabolize plastic. Yoshida et al. showed that biodegradation of plastics by these specialized bacteria could be a viable bioremediation strategy [44]. The new species, Ideonella sakaiensis, breaks down plastic by using two enzymes to hydrolyze poly (ethylene terephthalate) PET (used in plastic products worldwide) and a primary reaction intermediate. Both enzymes are needed to efficiently convert

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Fig. 9.15 A display of Oceanospirillales bacteria grown on arsenic

Fig. 9.16 This is a photo of plastic pollution on a beach in Ghana in 2018

PET into its monomers: terephthalic acid and ethylene glycol [45]. The new bacterium is able to use PET as its major source of energy and carbon. It is Gram-negative, aerobic, and rod-shaped. HEAVY METALS ARE WATER POLLUTANTS TOO. In general, heavy metals are considered to be metallic elements with an atomic mass greater than 40 [46]. They occur naturally in the Earth’s crust and enter our environment as a result of air pollution, industrial discharge, and other ways. Here we will focus on cadmium,

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Fig. 9.17 Cadmium metal is displayed next to a U.S. penny

mercury, and lead. These toxic metals are on the World Health Organization’s (WHO) list of chemicals of major public concern. Cadmium is an element with the symbol Cd and the atomic number 48. It is a soft, silvery-white metal and occurs as a minor component in most zinc ores and in zinc production. It is resistant to corrosion and has been used as a protective plate for other metals. Cadmium is mainly produced as a byproduct of mining. It has been used in pigments, coatings, electric batteries, and electroplating. Exposure to cadmium is associated with a large number of illnesses including kidney disease, cardiovascular diseases, lung cancer, etc. Figure 9.17 displays the metal cadmium [47]. Heavy metals are highly toxic elements that affect human health and contaminate our food and water supply. Therefore, methods are needed to reduce heavy metal bioaccumulation. In a previous study, Lactobacillus rhamnosus GR-1 (LGR-1) reduced heavy metal bioaccumulation in a Tanzanian group of women and children. It was hypothesized that LGR-1 could sequester the heavy metals lead and cadmium by reducing their absorption across intestinal epithelium, and other Lactobacilli reduced the amount of lead and cadmium in solutions tested at concentrations of 0.5–50 mg/L [48]. The results of this work offer a simple and effective way to reduce the amount of heavy metals absorbed from foods in contaminated locations. In another study, the results showed that Pseudomonas aeruginosa can be used as a suitable biosorbent material for the removal of cadmium and other heavy metals from contaminated water, etc. [49]. Mercury has the symbol Hg and the atomic number 80. It is a heavy, silvery-white metal in liquid form. This element mostly occurs in deposits of cinnabar (mercuric sulfide). Mercury is mainly used for the manufacture of industrial chemicals or

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Fig. 9.18 A display of liquid mercury

for electrical and electronic applications. Mercury and most of its compounds are extremely toxic and must be handled with care. Mercury can be absorbed through the skin and mucous membranes and its vapors can be inhaled. Symptoms of mercury poisoning include muscle weakness, nausea, poor coordination, numbness in hands and feet, kidney disorders, etc. Figure 9.18 displays liquid mercury [50]. Bacteria have developed a variety of resistance mechanisms to counteract heavy metal stress. Several examples include the formation and sequestration of heavy metals in complexes and reduction of the metal to a less toxic form. Pseudomonas aeruginosa use the mer operon that reduces toxic Hg+2 to volatile Hg0 , which diffuses out of the cell [51]. Researchers have also reported that biofilms are capable of removing heavy metals from wastewater [52–54]. Electron microscopy showed that mercury-reducing Pseudomonas putida biofilms were found to accumulate elemental mercury on the exterior of the biofilms [55]. Lead is soft and malleable and has the symbol Pb and atomic number 82. It is denser than most materials, is gray in color, and is generally combined with sulfur. Lead has been used in batteries, bullets, gasoline, radiation shielding, etc. Over the years its use has decreased, because it is a highly toxic metal that can be inhaled or swallowed. Lead can cause severe damage to the brain and kidneys, and even result in death. Lead ions (Pb+2 ) are incorporated into teeth and bones because they are similar to calcium (Ca+2 ) ions. Lead interferes with the function of enzymes that require calcium ions. It also causes chronic neurological problems and blood-based disorders. A recent problem with lead was experienced in 2014 by residents in Flint, Michigan (U.S.A.). This city had lead contamination in its drinking water that was

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Fig. 9.19 A display of lead solidified from the molten state

attributed to corrosion in the lead and iron pipes that distributed water to the people. Figure 9.19 shows lead solidified from the molten state [56]. As already mentioned, heavy metals contaminate drinking water. Examples of known mechanisms of heavy metal toxicity include oxidative stress (such as an imbalance between free radicals and antioxidants in the body) and interference with protein folding and functions [57]. Researchers found that Burkholderia cepacia on hematite and alumina surfaces preferred to accumulate lead ions (Pb+2 ) at concentrations higher than 1 µM [58]. This suggests that the chemical nature of the attachment surface affects metal sequestration. It has also been found that for a biofilm, EPS (the polysaccharide components) binds heavy metals [59].

References 1. Dervisoglu, R. File: Solid oxide fuel cell protonic.svg. Date: May 2012. License: This work is in the public domain. https://commons.wikimedia.org/wiki/File:Solid_oxide_fuel_ cell_protonic.svg. 2. Arizona State University (Jan 7, 2008) Fuel cell that uses bacteria to generate electricity. Science Daily. https://www.sciencedaily.com/releases/2008/01/080103101137.htm. 3. Wiley (June 27, 2017) Coating bacteria with electron-conducting polymer for microbial fuelcells: Coating of individual bacterial cells with an electron-conducting polymer provides for a high-performance anode for microbial fuel-cell applications. Science Daily. https://www. sciencedaily.com/releases/2017/06/170627105323.htm.

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4. Guy, M. F. C. (2010). File: Soil MFC.png. Date: September 1, 2010. License: Creative Commons Attribution-Share Alike 3.0. https://commons.wikimedia.org/wiki/File:SoilMFC. png. 5. KVDP. File: Plant microbial fuel cell.png. Date: April 23, 2010. License: This work is in the public domain. https://commons.wikimedia.org/wiki/File:Plant_Microbial_Fuel_Cell.png. 6. Chen, T., Barton, S. C., Binyamin, G., Gao, Z., Zhang, Y., Kim, H.-H., et al. (2001). A miniature biofuel cell. Journal of the American Chemical Society, 123(35), 8630–8631. 7. Bullen, R. A., Arnot, T. C., Lakeman, J. B., & Walsh, F. C. (2006). Biofuel cells and their development. Biosensors & Bioelectronics, 21(11), 2015–2045. 8. Venkata Mohan, S., Veer Raghavulu, S., & Sarma, P. N. (2008). Biochemical evaluation of bioelectricity production process from anaerobic wastewater treatment in a single chambered microbial fuel cell (MFC) employing glass wool membrane. Biosensors & Bioelectronics, 23(9), 1326–1332. 9. Venkata Mohan, S., Veer Raghavulu, S., & Sarma, P. N. (2008). Influence of anodic biofilm growth on bioelectricity production in single chambered mediatorless microbial fuel cell using mixed anaerobic consortia. Biosensors & Bioelectronics, 24(1), 41–47. 10. Liu, H., Grot, S., & Logan, B. E. (2005). Electrochemically assisted microbial production of hydrogen from acetate. Environmental Science and Technology, 39(11), 4317–4320. 11. Sleutels, T. H. J. A., Lodder, R., Hamelers, H. V. M., & Buisman, C. J. N. (2009). Improved performance of porous bio-anodes in microbial electrolysis cells by enhancing mass and charge transport. International Journal of Hydrogen Energy, 34(24), 9655–9661. 12. Winter, C.-J. (2005). Into the hydrogen energy economy-Milestones. International Journal of Hydrogen Energy, 30(7), 681–685. 13. Rizzi, F., Annunziata, E., Liberati, G., & Frey, M. (2014). Technological trajectories in the automotive industry: Are hydrogen technologies still a possibility? Journal of Cleaner Production, 66, 328–336. 14. Deretsky, Z. (National Science Foundation). File: Microbial electrolysis cell.png. Date: April 22, 2010. License: This work is in the public domain. It is a work of the U.S. federal government. https://commons.wikimedia.org/wiki/File:Microbial_electrolysis_cell.png. 15. Nevin, K. P., Woodard, T. L., Franks, A. E., Summers, Z. M., & Lovley, D. R. (2010). Microbial electrosynthesis: Feeding microbes electricity to convert carbon dioxide and water to multicarbon extracellular organic compounds. mBio, 1(2), e00103–e00110. https://doi.org/10.1128/ mbio.00103-10. 16. Johann Dreo, E. P. A. File: Nitrogen Cycle.svg. Date: September 27, 2009. License: Creative Commons Attribution-Share Alike 3.0. https://commons.wikimedia.org/wiki/File:Nitrogen_ Cycle.svg. 17. Kelvinsong. File: Cyanobacterium—inline.svg. Date: January 23, 2013. License: Creative Commons Attribution-Share Alike 3.0. https://commons.wikimedia.org/wiki/File: Cyanobacterium-inline.svg. 18. Chandra, S., Sharma, R., Singh, K., et al. (2013). Application of bioremediation technology in the environment contaminated with petroleum hydrocarbon. Annals Microbiology, 63(2), 417–431. 19. Lopez, A., Lazaro, N., Priego, J. M., & Marques, A. M. (2000). Effect of pH on the biosorption of nickel and other heavy metals by Pseudomonas fluorescens 4F39. Journal of Industrial Microbiology and Biotechnology, 24, 146–151. 20. Nanda, M., Kumar, V., & Sharma, D. K. (2019). Multimetal tolerance mechanisms in bacteria: The resistance strategies acquired by bacteria that can be exploited to clean-up heavy metal contaminants from water. Aquatic Toxicology, 212, 1–10. 21. File: Enterobacter cloacae 01.png. License: This is the work of the Centers for Disease Control and Prevention. It is in the public domain. https://commons.wikimedia.org/wiki/File: Enterobacter_cloacae_01.png. 22. Rousseaux, C. (2011). Geobacter: The junk food connoisseurs of the bacterial kingdom. Department of Energy. https://www.energy.gov/articles/geobacter-junk-food-connoisseursbacterial-kingdom.

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44. Yoshida, S., Hiraga, K., Takehana, T., et al. (2016). A bacterium that degrades and assimilates poly (ethylene terephthalate). Science, 351(6278), 1196–1199. https://doi.org/10.1126/science. aad6359. 45. Tanasupawat, S., Takehana, T., Yoshida, S., Hiraga, K., Oda, K. (2016). Ideonella sakaiensis sp. nov., isolated from a microbial consortium that degrades poly (ethylene terephthalate). International Journal of Systematic and Evolutionary Microbiology, 66(8), 2813–2818. https:// doi.org/10.1099/ijsem.0.001058. 46. Zhang, H., Walker, T. R., Davis, E., Ma, G. (September 2019). Ecological risk assessment of metals in small craft harbour sediments in Nova Scotia, Canada. Marine Pollution Bulletin, 146, 466–475. https://www.sciencedirect.com/science/article/abs/pii/S0025326X19305144? via%3Dihub. 47. File: CadmiumMetalUSGOV.jpg. License: This work is in the public domain. https://commons. wikimedia.org/wiki/File:CadmiumMetalUSGOV.jpg. 48. Daisley, B. A., Monachese, M., Trinder, M., Bisanz, J. E., Chmiel, J. A., Burton, J. P., et al. (2019). Immobilization of cadmium and lead by Lactobacillus rhamnosus GR-1 mitigates apical-to-basolateral heavy metal translocation in a Caco-2 model of the intestinal epithelium. Gut Microbes, 10(3), 321–333. https://doi.org/10.1080/19490976.2018.1526581. 49. Chelliaiah, E. R. (2018). Cadmium (heavy metals) bioremediation by Pseudomonas aerugnosa: A minireview. Applied Water Science, 8(154). https://doi.org/10.1007/s13201-018-0796-5. 50. Bionerd. File: Pouring liquid mercury bionerd.jpg. Date: 2008. License: Creative Commons Attribution 3.0 unported. https://en.wikipedia.org/wiki/File:Pouring_liquid_mercury_bionerd. jpg. 51. Outten, F. W., Outten, C. E., & O’Halloran, T. (2000). Metalloregulatory systems at the interface between bacterial metal homeostasis and resistance. In G. Storz & R. Hengge-Aronis (Eds.), Bacterial stress responses (pp. 145–157). Washington, D.C: ASM Press. 52. von Canstein, H., Li, Y., Timmis, K. N., Deckwer, W.-D., & Wagner-Döbler, I. (1999). Removal of mercury from chloralkali electrolysis wastewater by a mercury-resistant Pseudomonas putida strain. Applied and Environment Microbiology, 65, 5279–5284. 53. White, C., & Gadd, G. M. (1998). Accumulation and effects of cadmium on sulphate-reducing bacterial biofilms. Microbiology, 144, 1407–1415. 54. Azizi, S., Kamika, I., & Tekere, M. (2016). Evaluation of heavy metal removal from wastewater in a modified packed bed biofilm reactor. PLoS ONE, 11(5), e0155462. https://doi.org/10.1371/ journal.pone.0155462. 55. Wagner-Döbler, I., Lünsdorf, H., von Lübbenhüsen, T., Canstein, H. F., & Li, Y. (2000). Structure and species composition of mercury-reducing biofilms. Applied and Environment Microbiology, 66, 4559–4563. 56. Chemical Elements. File: Lead-2.jpg. Date: March 5, 2016. License: Creative Commons Attribution 3.0 unported. https://commons.wikimedia.org/wiki/File:Lead-2.jpg. 57. Nies, D. H. (1999). Microbial heavy-metal resistance. Applied Microbiology and Biotechnology, 51, 730–750. 58. Templeton, A. S., Trainor, T. P., Traina, S. J., Spormann, A. M., & Brown, G. E., Jr. (2001). Pb (II) distributions at biofilm-metal oxide interfaces. Proceedings of the National Academy of Sciences of the United States of America, 98, 11897–11901. 59. Kazy, S. K., Sar, P., Singh, S. P., Sen, A. K., & D’Souza, S. F. (2002). Extracellular polysaccharides of a copper-sensitive and a copper-resistant Pseudomonas aeruginosa strain: Synthesis, chemical nature and copper binding. World Journal of Microbiology & Biotechnology, 18, 583–588.

Chapter 10

Biofilm Control and Thoughts for the Future

Abstract Biofilms affect many industries and our daily lives. Their formation and growth processes have common essential factors, but some of their components differ from one environment to another. Biofilms have a negative side in that they cause many problems. However, on a positive note, they provide benefits if used properly and effectively. Therefore, it is very important to be able to control biofilm formation and growth. In this chapter, we describe future trends for the research and applications of biofilms and the types of results that we can expect.

We have already discussed many current aspects of biofilms in various environments. So what is the future? Where are we going and what kind of results can we expect? This chapter describes topics of the future and our expectations for biofilms. To start, we would like to mention the theoretical development of biofilms. This topic relates to Chap. 2. On the theoretical side, the most interesting topic involves various reactions among components of biofilms and the following changes of chemical and physical properties in them with time. Since there are many components and complicated situations, this information has been hard to get. The schematic illustration of Redox reactions for biofilms is shown in Fig. 10.1 As the figure suggests, there are a couple of questions to answer. When the substrate is a metallic material, the dissolution of substrate metals into metallic ions in biofilms must occur. For these biofilms, some interactions between the metallic ions and EPSs must play a key role for one of the Redox reactions. What would it look like? We presume that the metallic ions would bind to some coordination positions of proteins to produce chelates or ligands. We need to clarify the phenomenon and to analyze it. On the other hand, bacteria are still an important component of biofilms. As was discussed in several chapters, bacteria in biofilms are different from those in the planktonic state. Even though they might be in dormant states, they would still be active to some extent. They have a special site of protein on their surfaces, called transporters. Some metals would be captured by the transporters. Therefore, the interaction between metallic ions and transporters on bacterial membranes must be considered. Also the mutual interactions among EPSs should be taken into account

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Fig. 10.1 Reactions among components in biofilms

and the interactions between EPSs and bacteria must be analyzed. All of these reactions and interactions must be analyzed in detail. Then we will get a chance to utilize the results to solve many problems related to biofilms in the future. To answer questions that are suggested in Fig. 10.1, electrochemistry seems to provide us with some good tools. Many reactions and interactions in biofilms are Redox reactions. Therefore, an electrochemical approach would be a good way to solve problems. Such electrochemical investigations have been carried out in various ways so far [1–20]. The electrochemical approach to biofilm investigation leads to four categories, as shown in Fig. 10.2. This approach would improve our understanding of corrosion [5–7, 13, 16, 19, 20]. The investigations about this topic have been focused on the analyses for the electrochemical dissolution of metallic materials. However, they should be more devoted to the interactions between metal ions and EPSs, even though the topic has been investigated to some extent [6, 16, 17, 19, 21–23]. This type of understanding would lead to the development of biofilm sensors. Various sensors have already been investigated and invented [24–33]. Lewandowski and Beyenal wrote good books about this topic [34, 35]. In particular, their application of microelectrodes to sensors was excellent, since microelectrodes would lead to small and practical sensors. Such an electrochemical study about sensors could be classified into two types: the potentiometric type [24, 36–40], and amperometry [24, 41–45]. However, better sensors are expected in the future as a result of electrochemical investigations. In particular, the impedance technique should be developed much more [2, 12, 27, 32, 46–51]. All of these investigations need information about

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Fig. 10.2 An electrochemical approach to investigate biofilms and the expected results

EPSs and the interactions among components in biofilms. Unfortunately, there is not enough data at this point. To fill the gap, structural analyses obtained by using advanced apparatus, such as NMR, Mass-spectroscopy, Raman Spectroscopy, FTIR, etc. will work well. The combination of electrochemical methods with advanced analytical methods will allow sensor research to advance much more. The development and findings for new electrode materials is needed too. Also highly sensitive sensor materials like graphene are expected [52–55]. Electrochemical investigations could clarify and control quorum sensing [3, 56– 65]. Even though we did not include much information about quorum sensing in this book, the process will be one of the important factors for understanding biofilms and solving problems related to them. For example, the quorum sensing signal transmission process controls biofilm formation. Therefore, the control of this transmission process could lead to a new medical treatment and to some new countermeasures for solving biofilm problems in various fields. To achieve this goal more effectively, the structural and molecular analyses should be combined with the electrochemical approach. Structural analyses are the promising topics. Unfortunately, electrochemical methods do not provide us with that information. This may be one of the limitations for electrochemistry. However, its combination with microscopic techniques could produce very powerful tools for analyzing biofilms [2, 10, 44, 57, 66–71]. We expect that these efforts and their results will clarify the structures of biofilms in the near future. In addition to electrochemical methods, we need new countermeasures to control the negative aspects of biofilms. These harmful effects have already been discussed and many countermeasures have been proposed. Since bacteria survive in severe environments and have very strong resistance to antibiotics, etc. biofilms can’t be completely eliminated. From the viewpoint, biofilms should be mitigated to some extent. Effective mitigating methods should be sought for in the future. Such an effort

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might lead to a never-ending story. However, we should continue our research and investigations. As for a countermeasure, let us introduce our unique proposal. For the past couple of years, we have investigated the application of alternate electromagnetic fields to control biofilms. We found that biofilm was controlled at some frequencies of the alternate electromagnetic field where the magnetic flux densities were a couple of militesla [72–75]. We are now trying to clarify the mechanism and to apply this fundamental finding to control biofilms. Our work consists of repeated experiments and investigations on the laboratory scale as well as some field experiments under a Japanese national project (Research and Development for physical treatment of scales in geothermal power generation utilizing hot spring heat. New Energy and Industrial Technology Development Organization: Research and Development of controlling scales for geothermal generation 2014–2018). According to our results and findings, biofilms were controlled at some discrete frequencies under a weak alternate electromagnetic field like 4, 8 kHz, etc. under the electromagnetic field of 1 mT. Conventionally, the relation between electromagnetic fields and biofilms has been discussed and investigated [76–84]. However, we believe that our findings have important characteristics. #1: The phenomenon was found repeatedly, when the alternate electromagnetic field was applied. #2: It occurred at some discrete frequencies just like the resonance phenomenon. #3: The frequencies affecting the biofilm formation and growth are restricted to several kHz to tens of KHz, the radio frequency region. #4: The frequency depends on the combination of substrate materials and the bacterial flora or species. Now we are struggling to clarify the mechanisms, so that the development would lead to a better countermeasure for the control of biofilms. One of the potential directions to clarify the mechanism is to study the correlation between the applied electromagnetism and the bacterial activities. DNA must be affected by the electromagnetic fields at radio frequencies [85–94]. On the other hand, quorum sensing might be affected by electromagnetism. Actually, we could find many investigations about it [95–104]. The mechanism differs from one case to another. However, now our focus is on the effect that electromagnetic waves have on polymeric substances at radio frequencies. Schnabel pointed out in his book [105] that electromagnetic waves would generally be absorbed into organic matter at high frequencies. Non-polarizable organic matter absorbs the electromagnetic field reversibly. It absorbs the electromagnetic wave once. However, it would be radiated after removing the electromagnetic field. We presume that this relates to the spin activation and relaxation processes. However, polarizable polymers absorb the electromagnetic wave irreversibly. It means that the absorbed energy from electromagnetic fields would be transformed to other energy types such as thermal energy, etc. The excitation is called oscillation and the disappearance is called relaxation. Figure 10.3 is a schematic illustration for the phenomenon. The horizontal

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Fig. 10.3 The effect of electromagnetic waves on polymeric substances

axis corresponds to the frequency and the vertical axis corresponds to the dielectric constant. The non-polarizable polymer shows the solid line with the dielectric constant decreasing with increasing frequency, while the polarizable polymer shows the decrease with some discrete peaks (the dotted line). Those peaks correspond to oscillation. We presume that such a phenomenon would apply to EPSs and to the signal transmission process by proteins or polysaccharides including the quorum sensing process. This is only an example. However, such an investigation (the application of electromagnetism to biofilm research) could be coupled with proteomics and metabolomics to develop biofilm investigations theoretically and also practically. Finally, we would like to mention and stress the importance of industrial standards for the testing of biofilms. This topic was discussed in Chap. 7 of the book. It is our impression and feeling that biofilm research has been restricted to a narrow geographical area, even though the impact, effect, and needs might be found in very broad areas. Therefore, in order to solve the many problems related to biofilms, we need various types of professionals and investigative approaches. We should have certain common testing methods available that are determined by an international standard. As shown in Chap. 7, we already have some. However, most of these methods have been based on the count (number) of bacteria. It should be mentioned that biofilms have many different stages, even though bacteria might have played an important and exclusive role initially. No. The bacterial count can’t be used to properly evaluate biofilms. In biofilms, there are lots of dead bacteria as well as viable bacteria. Also biofilms contain lots of VBNC (viable but non-culturable) bacteria, since many of them would be in the dormant state. The composition of biofilms is very complicated, so the counting of bacteria is not a good method for the standard. On the other hand, EPSs would affect materials at the middle or later stages of biofilm growth and formation. Therefore, the substrate material should be factored into the standard somehow. As mentioned in Chap. 7, the SIAA (Society of International Sustaining Growth for Antimicrobial Articles), a non-profit organization in Japan,

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is now going to establish a standard. The authors are involved in this project, which includes a materials’ factor so that anti-biofilm materials might be made more easily. We expect this new standard to be available in the near future. As mentioned in this chapter, there are still more problems and topics for biofilms. We hope that many researchers, engineers, doctors, medical experts, officers, financial businessmen, etc. will cooperate and work together to solve the existing biofilm problems and lead us in a better direction. Hopefully our book will be used as a guide to inspire this work for a better future.

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