Bacteriophage-based biocontrol of the biofilm formed by antibiotic-resistant bacterium

Antibiotics are not only essential for human health but also for the well-being of plants and animals. However, the use of antibiotics has led to the evolution of antibiotic-resistant bacteria (ARB) in the environment. Antibiotic resistance in bacteria is a serious public health concern. ARBs are not only a predicament in clinical scenarios and daily life but also existing in many other environments. Wastewater treatment plants have been acknowledged as breeding grounds for such bacteria. As part of this research, a kanamycin-resistant bacterium, Chryseobacterium taeanense strain K-2, was isolated from the Central Valley Water Reclamation Facility and chosen as a model organism for the study. The bacterium was Gram negative, 0.5 μm long with coccoid morphology, and belonged to the Flavobacteriaceae family. The doubling time was observed to be ~2 h. Some species of bacteria under this genus have been associated with pathogenicity in human. This is, however, the first report on an antibiotic-resistant strain of Chryseobacterium taeanense and its presence in wastewater can be a serious concern. A lytic bacteriophage infecting Chryseobacterium taeanense strain K-2 was isolated from Central Valley Water Reclamation Facility. The lytic infection was evident from distinct clear plaques of 1-2 mm diameter obtained on top agar plates. According to the TEM analysis, the phage belongs to the Podoviridae family with a short tail, hexagonal head, and a total length of 90-100 nm. The latent and eclipse period of lytic infection were found to be the same (10 min). The burst size was around 18±2 PFUs/infected cell. Significantly short latent period and relatively high burst size indicate the exceptional potential of the lytic phage to regulate even very high populations of the host. The optimal phage-to-host ratio for infection was found to be 100:1, causing 98.7% of host death after 9 h. The lytic phage was also found to infect the host cells within the biofilm. The biofilm disintegration and degradation of the extra-polymeric substance within the matrix of the biofilm was evident after overnight infection with the phage. In conclusion, the lytic phage isolated in the present study can be attributed with significant potential for biocontrol of both biofilm-forming and planktonic forms of antibiotic-resistant bacterium, Chryseobacterium taeanense strain K-2.

BACTERIOPHAGE-BASED BIOCONTROL OF THE BIOFILM FORMED BY ANTIBIOTIC-RESISTANT BACTERIUM by Ran Jing A thesis submitted to the faculty of The University of Utah in partial fulfillment of the requirements for the degree of Master of Science Department of Civil and Environmental Engineering The University of Utah August 2012 Copyright © Ran Jing 2012 All Rights Reserved The Univers i ty of Utah Graduate School STATEMENT OF THESIS APPROVAL The thesis of Ran Jing has been approved by the following supervisory committee members: Ramesh Goel , Chair 4/24/2012 Date Approved Andy Hong , Member 4/24/2012 Date Approved Otakuye Conroy-Ben , Member 4/24/2012 Date Approved and by Paul Tikalsky , Chair of the Department of Civil and Environmental Engineering and by Charles A. Wight, Dean of The Graduate School. ABSTRACT Antibiotics are not only essential for human health but also for the well-being of plants and animals. However, the use of antibiotics has led to the evolution of antibiotic-resistant bacteria (ARB) in the environment. Antibiotic resistance in bacteria is a serious public health concern. ARBs are not only a predicament in clinical scenarios and daily life but also existing in many other environments. Wastewater treatment plants have been acknowledged as breeding grounds for such bacteria. As part of this research, a kanamycin-resistant bacterium, Chryseobacterium taeanense strain K-2, was isolated from the Central Valley Water Reclamation Facility and chosen as a model organism for the study. The bacterium was Gram negative, 0.5 μm long with coccoid morphology, and belonged to the Flavobacteriaceae family. The doubling time was observed to be ~2 h. Some species of bacteria under this genus have been associated with pathogenicity in human. This is, however, the first report on an antibiotic-resistant strain of Chryseobacterium taeanense and its presence in wastewater can be a serious concern. A lytic bacteriophage infecting Chryseobacterium taeanense strain K-2 was isolated from Central Valley Water Reclamation Facility. The lytic infection was evident from distinct clear plaques of 1-2 mm diameter obtained on top agar plates. According to the TEM analysis, the phage belongs to the Podoviridae family with a short tail, iv hexagonal head, and a total length of 90-100 nm. The latent and eclipse period of lytic infection were found to be the same (10 min). The burst size was around 18±2 PFUs/infected cell. Significantly short latent period and relatively high burst size indicate the exceptional potential of the lytic phage to regulate even very high populations of the host. The optimal phage-to-host ratio for infection was found to be 100:1, causing 98.7% of host death after 9 h. The lytic phage was also found to infect the host cells within the biofilm. The biofilm disintegration and degradation of the extra-polymeric substance within the matrix of the biofilm was evident after overnight infection with the phage. In conclusion, the lytic phage isolated in the present study can be attributed with significant potential for biocontrol of both biofilm-forming and planktonic forms of antibiotic-resistant bacterium, Chryseobacterium taeanense strain K-2. TABLE OF CONTENTS ABSTRACT ....................................................................................................................... iii LIST OF FIGURES .......................................................................................................... vii LIST OF TABLES ............................................................................................................. ix ACKNOWLEDGMENTS .................................................................................................. x CHAPTERS 1 INTRODUCTION ...................................................................................................... 1 1.1 Antibiotic-resistant bacteria (ARB) in the environment ....................................... 1 1.2 Biofilms and biofilms formed by ARBs ............................................................... 3 1.3 Current therapeutics for ARB biofilms ................................................................. 7 1.4 Bacteriophages ...................................................................................................... 8 1.5 Phage therapy for ARB biofilms ......................................................................... 12 1.6 Objectives ........................................................................................................... 14 2 MATERIALS AND METHODS .............................................................................. 15 2.1 Isolation and characterization of a model antibiotic-resistant bacterium from wastewater ........................................................................................................... 15 2.1.1 Isolation of antibiotic-resistant bacteria (ARB) ......................................... 15 2.1.2 Phylogenetic characterization of ARB using molecular tools ................... 15 2.1.3 Biochemical characterization of isolated ARB using Gram staining ........ 16 2.1.4 DAPI (4', 6-diamidino-2-phenylindole) staining analysis ......................... 16 2.1.5 Growth curve ............................................................................................. 17 2.2 Isolation and characterization of a lytic bacteriophage infecting the isolated ARB .................................................................................................................... 18 2.2.1 Isolation of lytic bacteriophage .................................................................. 18 2.2.2 Viral enumeration using epifluorescence microscopy (EFM) ................... 20 2.2.3 Characterization of the isolated phage ....................................................... 21 2.2.4 Live and dead bacterial enumeration (EFM) ............................................. 22 2.2.5 Phage-to-host ratio analysis ....................................................................... 23 vi 2.2.6 One-step growth curve ............................................................................... 24 2.3 Biofilm quantification and analysis .................................................................... 25 2.3.1 Biofilms formation and phage therapy ...................................................... 25 2.3.2 Biofilms quantification using by epifluorescence microscopy .................. 25 2.3.3 Scanning electron microscope images analysis ......................................... 26 3 RESULTS AND DISCUSSION ................................................................................. 28 3.1 Characterization of Chryseobacterium taeanense K-2 ......................................... 28 3.1.1 Isolation of antibiotic-resistant bacterium, Chryseobacterium taeanense K-2 ......................................................................................................................... 28 3.1.2 16S rRNA gene phylogenetic analysis ........................................................ 30 3.1.3 Growth analysis of Chryseobacterium taeanense K-2 ................................. 31 3.1.4 DAPI (4',6-diamidino-2-phenylindole) staining analysis ............................ 34 3.1.5 Gram staining ............................................................................................... 34 3.2 Characterization of bacteriophage of the host of Chryseobacterium taeanense k-2 ......................................................................................................................... 37 3.2.1 Isolation of bacteriophage ............................................................................ 37 3.2.2 Viral enumeration using epifluorescence microscopy (EFM) ..................... 37 3.2.3 Purification of lytic bacteriophage ............................................................... 39 3.2.4 Transmission electron microscopy (TEM) analysis .................................... 40 3.2.5 One-step growth curve ................................................................................. 42 3.2.6 Phage-to-host ratio analysis ......................................................................... 44 3.3 Biofilms disintegration using phage therapy ........................................................ 47 3.3.1 Phage therapy for biofilm ............................................................................ 47 3.3.2 Biofilms quantification using epifluorescence micrographs and BioImageL ............................................................................................................................... 47 3.3.3 Scanning electron microscope images analysis ........................................... 53 4 CONCLUSIONS ......................................................................................................... 56 4.1 Conclusions from study .................................................................................. 56 APPENDIX ....................................................................................................................... 57 REFERENCES ................................................................................................................. 63 LIST OF FIGURES page 1. Processes governing biofilm formation……….………….……………...…….…….....4 2. Lytic process of the bacteriophage infection process ................................................... 10 3. Lysogenic process of the bacteriophage infection process ........................................... 11 4. Cloudy plaque and clear plaque .................................................................................... 19 5. Chryseobacterium taeanense k-2 screening with 40 ug/ml Kanamycin and without Kanamycin ........................................................................................................................ 24 6. Phylogram depicting the phylogenetic relationship of Chryseobacterium taeanense K-2 .......................................................................................................................................... 3􀀕􁔠 8. Growth curve of Chryseobacterium taeanense K-2 .......................................................... 7. Micrographs obtained from DAPI analysis of Chryseobacterium taeanense K-2 from the log-phage culture ......................................................................................................... 3􀀖􁘠 9. Phase-contrast micrographs of Gram-negatively stained Chryseobacterium taeanense K-2 .................................................................................................................................... 3􀀘􁠠 10. Agar plate containing clear plaques ............................................................................ 3􀀙􁤠 11. Virus like particles (green dots) pointed out by yellow arrows in the epifluorescent micrographs ....................................................................................................................... 3􀀛􁬠 12. The bacteriophages' band after Cesium chloride density gradient purification ......... 􀀗􁜀􀀓􁌠 13. TEM micrographs of the lytic bacteriophage infecting Chryseobacterium taeanense K-2 .................................................................................................................................... 4􀀔􁐠 14. One-step growth curve of lytic bacteriophage infecting Chryseobacterium taeanense K-2 .................................................................................................................................... 4􀀖􁘠 viii 15. Profile of VLPs at different PHR ................................................................................ 4􀀘􁠠 16. Profile of Live/Dead Cells ratios at different PHR ..................................................... 4􀀘􁠠 17. Epifluorescent micrographs of 16-day biofilm (a1& a2) before and (b1 & b2) after infection with phage .......................................................................................................... 4􀀛􁬠 18. Epifluorescent micrographs of 54-day biofilm (a1& a2) before and (b1 & b2) after infection with phage .......................................................................................................... 4􀀜􁰠 19. Epifluorescent micrographs of 84-day biofilm (a1& a2) before and (b1 & b2) after infection with phage .......................................................................................................... 􀀘􁠀􀀓􁌠 20. Effect of phage therapy on % Live cells in the biofilm of different ages ................... 5􀀔􁐠 21. Effect of phage therapy on % Dead cells in the biofilm of different ages .................. 5􀀔􁐠 22. Effect of phage therapy on % EPS in the biofilm of different ages ............................ 5􀀕􁔠 23. SEM micrographs of 16-day-old biofilm (a1&a2) before and (b1 & b2) after phage infection ............................................................................................................................ 5􀀗􁜠 24. SEM micrographs of 54-day-old biofilm (a1& a2) before and (b1 & b2) after phage infection ............................................................................................................................ 54 25. SEM micrographs of 84-day-old biofilm (a1&a2) before and (b1 & b2) after phage infection ............................................................................................................................ 55 LIST OF TABLES Table Page 1: The bacteriophage's size obtained from TEM images ................................................. 42 2: Effect of initial phage/host ratio on bacteriolysis and VLP count ................................ 46 3: Live/dead cells and EPS distribution of the biofilm of the age of 16 days without phages infected .................................................................................................................. 57 4: Live/dead cells and EPS distribution of the biofilm of the age of 16 days with phages infected .............................................................................................................................. 58 5: Live/dead cells and EPS distribution of the biofilm of the age of 54 days without phages infected .................................................................................................................. 59 6: Live/dead cells and EPS distribution of the biofilm of the age of 54 days with phages infected .............................................................................................................................. 60 7: Live/dead cells and EPS distribution of the biofilm of the age of the 84 days without phages infected .................................................................................................................. 61 8: Live/dead cells and EPS distribution of the biofilm of the age of the 84 days with phages infected .................................................................................................................. 62 ACKNOWLEDGMENTS Thank you, in particular, to the chair of my committee, Dr. Ramesh Goel. He always encouraged me and gave a constructive criticism, and spent time to answer my questions with great patience. His advice and guidance were completely invaluable. Also, I want to thank Dr. Shireen Meher Kotay. He has given significant insight and future vision to my project. I would like to thank Choi Jeongdong, Amir M. Motlagh and Ananda Shankar Bhattacharjee for providing advice for the project. Moreover, I am grateful for Dr. Sherwood Casjens's research group for allowing and helping me to do the bacteriophage CsCl purification. CHAPTER 1 INTRODUCTION 1.1 Antibiotic-resistant bacteria (ARB) in the environment Antibiotics are drugs which can be used to treat or prevent disease and infections caused by bacteria and other organisms. During the past 60 years, antibiotics played a significant role in fighting against infections caused by bacteria and other microbes. Also, antimicrobial therapy is the most possible explanation of the increase in the average life expectancy. Due to the invention of antibiotics, some diseases and infections can now be easily cured within a few days. However, recently, the unrestrained usage of antibiotics has become a public health problem. Due to excessive use, large amounts of antibiotics are discarded and added to human medicine, agriculture, aquatic and several other natural environments (Huang et al. 2001). Some bacteria present in these environments resist antibiotics by modifying their genetic makeup, a process termed as mutation. These mutations are carried across generations and eventually develop into potent (sometimes multi-) antibiotic-resistant populations. In 1998，in the United States, 12,500 tons of antibiotic prescriptions were used for human beings (Kutateladze et al. 2010). Also, over 60% of antibiotics usage in the U.S was for agricultural practice, adding an additional 18,000 tons per year to the antibiotic burden in the environment. 􀀕􁔠 It is said that about 70 % of the bacteria that caused infections in hospitals can be resistant to more than one common antibiotic drug used for the treatment. In addition, some bacteria can be resistant to all approved antibiotics and the infection caused by these bacteria can only be treated using potentially toxic drugs. In case of community acquired infections, some bacteria have been reported to have antibiotic resistance, such as the Staphylococci and Pneumococci. One of the recent studies (Bisht et al. 2009) indicated that 25% of bacterial pneumonia cases were resistant to penicillin, and 25% of other cases were reported to be resistant to multiple antibiotics. Antibiotic-resistant bacteria and genes have been detected in wastewaters, and the concentrations are higher than that found in surface waters (Börjesson 2009). Antibiotics and their products can be introduced into wastewater or natural environment by humans or animals (Levy 2002). The antibiotics can become a selective pressure for maintaining resistance among microorganisms. Due to the favorable conditions (high nutrients), wastewater treatment plants can support luxuriant growth of ARBs. Furthermore, the antibiotic resistance can be easy to spread across bacteria due to diverse bacterial communities that harbor the wastewater systems. These ARBs can enter/re-enter the ecosystem through receiving water-bodies (rivers, streams, oceans, lakes etc.). Thus, the wastewater system has a major role to play in the emergence and spread of antibiotic resistance within bacterial populations. It also has a potential for working as a model in the development and biocontrol of the antibiotic-resistant bacteria. Bacteria can gain resistance through two primary means: mutation, and swapping 􀀖􁘠 DNA by using built-in design features (horizontal gene transfer) to share resistant genes from other microorganisms. Antibiotics can kill bacterial cells by disrupting a critical function. The antibiotics specifically bind to some bacterial cell proteins, which are involved in copying the DNA, making proteins, or making the bacterial cell wall, important for the bacteria to grow and reproduce, causing the failure of the protein's functions. If there are mutations in the bacterial genomic DNA, which code for these proteins, the antibiotic cannot bind to the proteins and the bacteria will survive. Therefore, in the presence of antibiotic, the mutant bacterial cells will predominantly survive. On the other hand, bacteria can get antibiotic resistance by gaining the mutant DNA from other bacteria; this phenomenon is termed horizontal gene transfer (Kay et al. 2002). The horizontal gene transfer often involves plasmids (Naik et al. 1994) where the genes specifically coded for antibiotic resistance in one species of bacteria can be transferred to another species of bacteria. In this case, the process requires an environment with genetic diversity and natural selection as the driving forces guiding bacteria resistance to antibiotics. 1.2 Biofilms and biofilms formed by ARBs Biofilms are microbial communities in polysaccharide-rich extracellular matrices and living in association with surfaces (Pace 2006). As a normal mechanism for bacteria to survive in the environment, when they are exposed to surfaces with a sufficient amount of nutrients, biofilms are formed. The polysaccharide layer is created to protect from the 􀀗􁜠 harmful environments and provide the optimal growth conditions for the survival of bacterial cells. For example, some external biofilms, namely chronic wounds and dental plaque, can be manually removed. Because of their inaccessibility and heightened resistance to certain antibiotic combinations and dosages, internal biofilms are more difficult to eradicate. Researchers have estimated that 60-80 % of microbial infections in the body are caused by bacteria growing as a biofilm - as opposed to planktonic (free-floating) bacteria. Biofilms spreading along implanted tubes/wires, prosthetic limbs or catheters can lead to pernicious infections in patients. In industrial environments, biofilms can develop on the interiors of pipes and lead to clogs and corrosion. Biofilms on floors and counters can make sanitation difficult in food preparation areas. The mechanism of biofilm-formation is a complex process; in general, the whole process consists of nine steps, as depicted in Figure 1 (Simoes et al. 2009): Figure 1: Processes governing biofilm formation (Breyers & Ratner, 2004). 􀀘􁠠 i) Preconditioning of the adhesion surface; ii) Transport of planktonic cells from the bulk liquid to the surface; iii) Adsorption of cells at the surface; iv) Desorption of reversibly adsorbed cells; v) Irreversible adsorption of bacterial cells at a surface; vi) Production of cell-cell signaling molecules; vii) Transport of substrates to and within the biofilm; viii) Substrate metabolism by the biofilm-bound cells and transport of products out of the biofilm. These processes are accompanied by cell growth, replication, and EPS production; ix) Biofilm removal by detachment or sloughing. Initially, free-floating antibiotic resistant bacterial cells attach to a surface through weak van-der-Waals forces and anchor to the surface permanently using cell adhesion molecules. These first colonists as initial adhesion sites will promote more and other bacterial cells to arrive and begin to build the matrix that holds the biofilm together. The matrices can provide a protection and promote communication among the cells through chemical and physical signals. It is known that some biofilms' matrix contain water channels that help distribute nutrients and signaling molecules (Haggag 2010). Some bacterial species can attach to surfaces on their own. Others are able to attach to the matrix or earlier colonists. Once colonization finishes, these bacterial layers will develop into larger cell clusters and create the structures of biofilm by combination of cell 􀀙􁤠 division and recruitment. It is noticed that bacteria in biofilms have significantly different properties from free-floating bacteria. The cells are dense as a protected environment that allows them to cooperate and interact. The benefit of dense cells is that the outer layer of cells and extracellular matrix protect the interior of the community to increase resistance to antimicrobials and antibiotics. The benefit of dense cells is that the outer layer of cells and extracellular matrix protect the interior of the community to increase resistance to antimicrobials and antibiotics. In other words, it is not all biofilm cells represent highly resistant, biofilms' antibiotic resistance can be determined by the susceptibility of the most resistant cell. On the other hand, ARBs that grow as adherent biofilms are more difficult to eliminate. ARBs can possibly reside in biofilms and lead to enhanced tolerance to adverse environmental conditions, causing serious infectious diseases (Ngwai et al. 2006). ARBs can resist even high doses of antimicrobial drugs, making traditional therapeutics obsolete. Moreover, the antibiotic-resistant pathogens can exist in the form of biofilms, leading to cross-resistances to other environmental stresses (Gilbert et al. 2002). However, there is a lack of information on the biofilm-associated infections involved in altered virulence properties of antibiotic-resistant bacteria. Relatively few studies have been focused on the biofilm-forming abilities of multiple antibiotic-resistant pathogens. The biofilm formation by multidrug-resistant pathogens may increase the risk of severe infections related to food processing facilities and medical devices. 􀀚􁨠 1.3 Current therapeutics for ARB biofilms Prevention strategies for antibiotic resistance are essential to control the spread of antibiotic-resistant pathogens. However, the discovery and development of novel antibiotics has lagged behind the emergence of antibiotic-resistant pathogens because of the lengthy and expensive processes, requiring phases of clinical investigation trials to obtain approval, and the lack of information on the antibiotic resistance mechanisms. Drugs like linezolid and daptomycin with novel modes of action were developed and approved (Fischbach and Walsh 2009). However, just a few years after their introduction into the market, clinical strains resistant to those antibiotics were reported. These observations imply that the traditional approach of antibiotic discovery, though highly successful in the introduction of numerous drugs, cannot sustain the high demand for development of novel compounds (Breithaupt 1999). Therefore, a more efficient antibiotic discovery platform is essential for allowing us to compete with the evolution of microbial resistance (Davies 1994). In the post-genomic era, with the availability of various genomics-based platforms including whole-genome sequencing, genotyping, and gene expression profiling, a new horizon opens that could revolutionize our pursuit of novel antimicrobial agents (Amini and Tavazoie 2011). However，there are still many limitations when these therapeutics are applied to control biofilms formed by antibiotic-resistant bacteria. For example, chemicals can pass over the surface of a biofilm and only affect bacteria at that surface instead of penetrating the biofilm and the EPS matrix. Biofilms can form layers and matrices that can decrease 􀀛􁬠 the activity of chemical treatment to the point where effective concentrations cannot be achieved. Hence, it is required that biocides must be mobile so they can migrate to the film surface and across the bacteria cell membrane to destroy bacteria. Moreover, the emergence of resistant bacteria to conventional antimicrobials results in a large amount of antibiotics use, demonstrating that novel biofilm control strategies are required. Novel drug delivery technologies have been studied as targets for controlling the process of biofilm formation. To prevent colonization, this technology combines devices of surface modification with biofilm development along with antimicrobials. Electrical approaches are used to release antimicrobials from device surfaces or introduce antimicrobials through the biofilm (Smith 2005). Other therapeutics, such as liposomal systems, have been used to aim antibiotics onto the biofilms' surface, or to target antibiotics towards intracellular bacteria. In addition, new concepts in case of a polymer-based carrier such as biodegradable polymers has been proposed. Phage therapy can help to control biofilm by killing bacteria either before they are able to attach to a surface or before they proliferate. Hence, bacteriophages might provide a possible natural, highly specific, nontoxic way for controlling biofilm formation (Kudva, Jelacic, Tarr, Youderian, and Hovde 1999). 1.4 Bacteriophages Bacteriophages are viruses which can infect and kill a specific bacterial host. Bacteriophages cannot replicate independently without a living host cell. Therefore, 􀀃􀌀􀀃􀌀􀀃􀌀􀀜􁰠 bacteriophages enter into a suitable living cell and utilize energy, metabolic intermediates, and protein for their replication cycle. In general, there are two kinds of cycles, one called the lytic cycle and the other the lysogenic cycle. Figures 2 and 3 depict two kinds of infection processes. The lytic cycle is typically considered the main method of viral replication, since it results in the destruction of the infected cell. In this process, the first step of the lytic infection in its host cell is called attachment (adsorption). The bacteriophages adhere to the bacterial surface using their tail fibers by a chance collision at a chemically complementary site. After adsorption, the bacteriophage injects the DNA into the host cell through a process called penetration. In this process, the tail sheath contracts and the core inside the bacteriophage is driven through the wall to the membrane under mechanical and enzymatic conditions. Immediately after injection of the DNA, synthesis of early proteins, transcription, and translation of a section of the phage DNA begins. In the transcription process, the early proteins as the repair enzyme are produced in order to repair the broken bacterial cell wall. On the other hand, in the translation process, the DNase can degrade the host DNA into precursors of phage DNA; the phage-specific DNA polymerase will copy and replicate phage DNA. During this period, the final result is the synthesis of several copies of the phage DNA. The next step is called synthesis of late proteins. The replicated copies of phage DNA in the previous step can be used for is the synthesis of several copies of the phage DNA. The next step is called synthesis of 1􀀓􁌠 Figure 2: Lytic process of the bacteriophage infection process late proteins. The replicated copies of phage DNA in the previous step can be used for transcription and translation of late proteins. The structure of late protein is made up of capsomeres and the various components of the tail assembly. The lysozyme production is the last step of the replication process. It assembles in the tail and is used to escape from the host cell by breaking the cell wall. Thus, the host cell is lysed, releasing mature viruses. Lysogenic cycle is another method of viral repro Lysogenic cycle is another method of viral reproduction. During the lysogenic cycle duction. During the lysogenic cycle Phages attach receptor site and penetrate DNA Proteins production and copies of phage DNA Empty phage heads are synthesized Head are packed with DNA Collars, sheaths, and base plates assembling Bacterial cell lysis and releasing matured phages 􀀃􀌀􀀃􀌱1􀀔􁐠 Phage's chromosome integrated into specific section of the bacterial chromosome Phage's chromosome replicated along with bacterial DNA replication Binary fission is completed, and each cell has phage's DNA Phages attach receptor site and penetrate DNA Figure 3: Lysogenic process of the bacteriophage infection process process, bacteriophages' chromosomes will become integrated into a specific section(s) of the host bacterial cell instead of killing the bacteria. In addition, the phage genes can code for synthesis to prevent synthesis of phage enzymes and proteins required for the lytic cycle. This phage DNA is called prophage and the host bacteria are said to be lysogenized. These new integrated genetic materials can be transferred to each subsequent generation and released via a new lytic cycle caused by environmental stress (such as UV radiation or PH changing). This process is very rare; however, it also can 1􀀕􁔠 assure new phages are formed which can proceed to infect other cells. Bacteriophages are ubiquitous and are very important for freshwater and marine bacterial communities (Suttle 2006; Wommack and Colwell 2000). Bacteriophages have wide applications in environmental issues including controlling cyanobacterial blooms, and reducing the sludge bulking formation of filamentous bacteria. 1.5 Phage therapy for ARB biofilms Phage therapy is the therapeutic use of bacteriophages to treat pathogenic bacterial infections. It is one approach that has great potential as a solution to the serious worldwide problem of drug-resistant bacteria (Parisien et al. 2008; Sulakvelidze 2005). While they were administered as antibacterial agents as early as 1919, before the discovery of antibiotics, inadequate understanding of phage biology and genetics reduced the efficacy of phage therapy (Gill and Hyman 2009). When bacteriophages infect biofilms, the interactions will occur. The polysaccharide polymerase enzyme degrade the EPS and cause biofilm slough off and the biofilm's integrity might be destroyed, if the phage can produce polysaccharide-degrading enzymes. The interaction depends on the susceptibility of the biofilm cells to the phage and to the availability of receptor sites (Simoes 2010). It is known that fluorescent pseudomonas cells can be eliminated using bacteriophages and in the experiment, about 80% of biofilm was removed in the early stage of development and were 5 days old under optimal conditions (Sillankorva et al. 2004). In addition, there is some evidence that 1􀀖􁘠 indicates that bacteriophages can express biofilm-degrading enzymes which attack the bacterial cells in the biofilm and the biofilm matrix (Lu and Collins 2007). There are two advantages of the phage therapy in biofilm reduction. The first one is that they can multiply in direct correlation to an available growing host; the second one is the bacteriophages' concentration will increase as long as its host is present in the enclosed environment. However, bacteriophages that infect cells at the surface will replicate, generating a high concentration of phage locally at the site of the infection, which can then progressively destroy the biofilm. In other words, bacteriophages can infect bacterial cells and destroy the newly revived cells continuously. In addition, bacteriophages can carry or induce the expression by the bacterial host of enzymes that dissolve the biofilm matrix (Pearl et al. 2008). One study shows that 84% of biofilms forming by Pseudomonas fluorescens strain were reduced by 109 PFU/ml of bacteriophages treated after 200 min and at 26􀀀 (Sillankorva et al. 2008). In addition, another study indicated that 28-hour biofilm formed from E.coli 3000 was 6-log reduction after treating with 109-1010PFU/ml phage. One of the drawbacks of phage therapy is the narrow host range of bacteriophages. The phage receptors are different in gram-negative and gram-positive bacteria (Kutter et al. 2005), and therefore, some phages have specificity at the strain level. The diversity of bacterial infections implies that it may be difficult for any particular engineered phage to be a therapeutic solution for a wide range of biofilms. Because of the high specificity of phages, many negative results may have been obtained because of the failure to select 1􀀗􁜠 lytic phages for the targeted bacterial species. As a result, it is advantageous to design a phages cocktail, a mixture of phages with differing host ranges, which can degrade multiple extracellular polymers to cover a range of biofilm and target biofilms containing multiple strains or species. All in all, this study focused on the single strain of Chryseobacterium taeanense K-2 biofilm development and the implications and performance of biocontrol based on bacteriophage. The results will be applied for modeling of biofilm systems researching for full-scale wastewater treatment facilities. 1.6 Objectives The overarching goal of my research was to demonstrate the application of phage therapy for ARB biofilms. The following specific objectives accomplished my research goal. Objective 1: Isolation and characterization of a model antibiotic resistant bacterium from wastewater. Objective 2: Isolation and characterization of a lytic bacteriophage infecting the isolated ARB. Objective 3: Demonstration of phage therapy for the isolated ARB biofilm. CHAPTER 2 MATERIALS AND METHODS 2.1 Isolation and characterization of a model antibiotic-resistant bacterium from wastewater 2.1.1 Isolation of antibiotic-resistant bacteria (ARB) The mixed liquor was collected from a local wastewater treatment plant and serially diluted in the ratio of 10-1, 10-2, 10-3, 10-4, and 10-5 using 0.85% (w/v) sterile saline solution. Subsequently, each diluted sample was plated uniformly onto the 2% modified R2A agar (Peptone 0.5g, Yeast Extract 0.25g, Agar 10g in 1 L deionized water) plate containing 40μg/ml kanamycin final concentration (Fisher Scientific, PA). The pH of the media was adjusted to 7.2 - 8.7. A method control was prepared in a similar way without antibiotic. The plates were then incubated at 37􀀀 overnight. Single isolated colonies were picked up and screened on antibiotic plates 4 times (subcultures) for obtaining pure culture of isolates. Purified subcultures were picked and grown in modified R2A medium (Peptone 0.5g, Yeast Extract 0.25g, in 1 L deionized water). Glycerol stocks of the pure culture were made from 12-h-old culture and store in -80°C until further use. Fresh liquid cultures of the isolate were made by inoculating a 16 glycerol stock in modified R2A medium each time. 2.1.2 Phylogenetic characterization of ARB using molecular tools Genomic DNA from fresh 10 h culture was extracted using the Soil DNA extraction kit (MoBio Labs, Solana Beach, CA) following manufacturer's protocol and extracted DNA was verified on a 1 %(w/v) agarose gel. PCR amplication of 16S rRNA gene was performed using universal primer, 8f (5'-AGAGTTTGATCMTGGCTCAG-3') and reverse universal primer, 1492r (5'-GGYTACCTTGTTACGACTT-3'), 2X GoTaq Mastermix and genomic DNA as a template. PCR program: initial denaturation at 95ºC for 5 min, 30 cycles of 90 sec denaturation at 95ºC, 120 sec of annealing at 52 ºC, and 3 min of extension at 72 ºC, with a final 10 min extension at 72º was used. Amplicon obtained from PCR was verified on 1 %(w/v) agarose gel and purified using a gel extraction kit (Qiagen, CA). The purified amplicon was sequenced using universal 8f and 1492r primers on an ABI PRISM 377 automated DNA sequencer (Applied Biosystems, USA). The sequences obtained were analyzed for homology using a web-programs BLAST and seqmatch. MEGA version 5 (http://www.megasoftware.net/) was used to align the sequences and generate the phylogenetic tree. 2.1.3 Biochemical characterization of isolated ARB using Gram staining Gram staining is widely used to distinguish two large groups of bacteria based on different cell wall constituents. The Gram positive bacteria stain violet due to the presence of more peptidoglycan in their cell walls which can retain the crystal violet. 17 Gram negative bacteria stain pink (by secondary stain-saffranin), because of their thinner peptidoglycan wall which cannot retain the primary stain, crystal violet. First, bacteria (from 10 h old fresh culture) was mounted onto a clean glass slide using a sterile inoculation loop and heat-fixed. The slide is then flooded with primary stain Crystal Violet for 1 min, following which the mordent, Gram's Iodine, was added. After 1 min, the excess stain was drained and slides were rinsed with deionized water. Slides were then decolorized with 100% ethanol for 1 min and rinsed with deionized water. Sunsequently, the slides were treated with secondary counter stain, safranin. 2.1.4 DAPI (4', 6-diamidino-2-phenylindole) staining analysis DAPI (4', 6-diamidino-2-phenylindole) is a fluorescent stain which is used extensively in fluorescence microscopy because it binds strongly with DNA. Fresh 10 h old bacterial culture was diluted 20 times in 1X Phosphate Buffer Saline (PBS), and fixed for 20 min in 4% paraformaldehyde. The fixed cells were collected on 0.22 μm polycarbonate membrane filter (GE Water & Process Technologies). Cells collected on the filter paper were carefully transfered on a clean microscope slide (Fisher Scientific). The cells on the slide were stained with 100 μL of 5μg/mL DAPI solutiuon and incubated in the dark for 20 minutes. The slides were then washed with sterile deinoized water a few times, and air dried for 5 min. A clean cover slide was put onto the microscope slide after the antifade solution was added on the sample and sealed with nail polish. The slides were then viewed under the appropriate filter of an epifluorescent microscope (Olympus 18 BX51). 2.1.5 Growth curve In microbiology, growth is defined as an increase in the number of cells. Microbial cells have a finite life span, and a species is maintained only as a result of continued growth of its population. A viable count measures the cells in the culture that are capable of reproducing. Optical density (turbidity), a quantitative measure of light scattering by a liquid culture, increases with the increase in cell number (Madigan et al. 2010). To determine the growth of C. taeanense K-2, 100ml of log-phage culture was inoculated into 1 L R2A medium and incubated at 37􀀀 with agitation provided. An aliquot of 1ml bacterial culture was collected at an interval of 30 min and the optical density of the culture was measured at 600nm. Sterile R2A media was used as blank. Growth-curve was deduced by plotting OD vs time (Sigmaplot). 2.2: Isolation and characterization of a lytic bacteriophage infecting the isolated ARB 2.2.1 Isolation of lytic bacteriophage The principle for determination of infection of the host bacterium by a bacteriophage is based on the single phage infecting the host, multiplying, and releasing their progeny, resulting in host cell lysis. The continuous infection causes the host cell lysis on agar plates. This process can be achieved by the double layer agar technique. The double layer agar technique is also known as the "soft agar overlay," or "top-agar" method of 19 plaque assay (Walker et al. 2009). The plaques appear on the 1% top agar after overnight incubation. The bacteriophage and the bacteria host are mixed with 1% top agar and spread on the base agar plate. The bacteriophage progeny will inhibit the host growth and result in the formation of clear zone (Figure 4). As shown in the figure, clear and turbid (cloudy) plaques may appear, representing lysogenic and lytic phage infection, respectively. Mixed liquor from the local full-scale wastewater treatment plant were obtained. In order to remove suspended particles and microorganisms, the mixed liquor sample was filtrated through 0.45 and 0.22 μm filters (Millipore, CA). The bacteriophages present in the filtrate were concentrated using Amicon Ultra-4 with MWCO 30000 (Millipore, CA). The concentrate obtained from the above centrifugation was used as phage stock in the subsequent studies. Figure 4: Cloudy plaque and clear plaque. [http://www.wwnorton.com/college/biology/microbiology2/ch/10/etopics.aspx] 20 From a fresh 10 h C. taeanense culture, 1ml aliquot was infected with 1μl of the above phage stock, mixed thoroughly on a vortex mixer and incubated at room temperature for 15-30 min. To this, 3ml presterilized molten 1% top agar (10g bacto-agar, 1g peptone, 0.5g yeast extract in 1L deionized water) was added and mixed thoroughly on a vortex mixer. The mixture was then spread on a premade 2% base agar plate. A negative control was prepared using the similar protocol without adding the phage stock. The plates were incubated at 37°C overnight. The plates that showed clear plaques were selected to make the lysate stock. These plaques were then picked using a sterilized pasture glass pipette, after which, the plaque with agar was transferred into a fresh and sterile polypropylene tube and resuspended in the SMG buffer (5.8 g NaCl, 2.46 g MgSO4, 7.9 g Tris-HCl, 0.1 g Gelatin in 1L deionized water). The phage solution in SM buffer was stored at 4°C until further use. The isolated phage-stocks were screened using the repeated subculture method to obtain a pure phage-stock. 2.2.2 Viral enumeration using epifluorescence microscopy (EFM) To indentify and enumerate the virus-like particles existing in the phage stock, the bacteriophage samples were filtered through 0.22 μm filter (Millipore,CA) to remove the suspended solids. In order to exclude the free bacterial DNA, 900 μL of the above filtrate was added to 100 μL of RQ1 reaction buffer and 2 μL of RNase free DNase I (Invitrogen) in a fresh polypropylene tube. After incubation for 20 min at 37 􀀀, 35 μL of 0.5 M EDTA was used to stop the DNase activity. The DNase treated sample was 21 vacuum filtered through a stack of 25mm filters consisting of 0.02 μm Anodisc (Whatman Int'l Ltd., Maidstone, England), a 0.22 μm Durapore membrane filters (Millipore, CA), and a glass fiber prefilter (Millipore, CA). Virus-like particles (VLPs) collected on the Anodisc filter were stained using 4 μl SYBR Gold stain 800 μl SMbuffer for 20 min. Finally, the Anodisc filter was transferred onto the slide containing a drop of freshly prepared antifade solution (Kotay et al. 2010) and sealed with a clean cover glass. The slides were observed under an Olympus BX 51 epifluorescence microscope (Olympus, Japan) using a Cy3 filter and VLPs were enumerated using the micrographs captured. 2.2.3 Characterization of the isolated phage The phage titer defines the concentration of infectious viral particles per milliliter of growth medium. By counting the number of plaques and multiplying by the serial dilution factor, we can determine the titer, which is the concentration of phage in the phage stock. To determine the titer, the phage stock was serial diluted 10-1, 10-2, 10-3, 10-4, 10-5, 10-6 times using sterile prefiltered SMG buffer. From each dilution of phage stock, 1μl was added into 1 ml of fresh 10 h C. taeanense K-2 culture and incubated at room-temperature for 1 h. To the mixture, 3ml 1% top agar was added, mixed thoroughly, and poured onto 2% base agar plates. Triplicates of each dilution were plated and plates were incubated at 37°C overnight. 22 CsCl density gradient method was used to purify the phage. First, 2 ml of cesium chloride with a density (ρ) of 1.6g/l was added into the bottom of an ultracentrifuge tube (Beckman, CA). On top of that, 2 ml of 1.4 g/l of cesium chloride was carefully overlaid using a syringe. On top of that, 2 ml 10% sucrose and 4 ml of sample were carefully overlaid without disturbing the layers below. Samples were centrifuged, at 36,000 rpm, at 20 °C for 2-3 h. The purified phage appeared as a distinct band at the interface of the ρ=1.4 g/l layer and ρ=1.6 g/l layer. The phage band was carefully siphoned from the side of the tube by using a syringe, transferred into a clean dialysis tube (Fisherbrand) and dialyzed overnight in SMG buffer to remove the CsCl and extra ions. On a clean 400 formvar-coated copper grid (Fisherbrand), 3.5 uL of the CsCl purified phage was loaded and incubated at room temperature for 2 mins. The excess liquid was soaked from the side of the grid using bibulous paper (Fisherbrand), and the sample was stained using 2 mL 1% uranyl acetate for 2 min (Kotay et al. 2010). The grids were observed under a Tecnai T12 Transition Electron Microscope (FEI, Japan). An accelerating voltage of 80 kV was used and micrographs revealing bacteriophage morphology were recorded. 2.2.4 Live and dead bacterial enumeration (EFM) Live/Dead BacLight™ (Molecular Probes Inc., CA) staining is one of the most important fluorescence assays used to determine and mark living bacteria. The major advantage of this method is it is easy to classify bacterial cells as living or dead in a few 23 minutes. There are two kinds of stains, green fluorescent SYTO® 9 and red fluorescent propidium iodide nucleic acid stains. These stains have the ability to label live and dead bacterial cells due to the diversity of their spectral characteristics. During the staining processing, the SYTO® 9 could stain both live and dead bacteria cells. The propidium iodide was just able to label live bacterial cells due to the ability of penetrating damage of the bacterial cell membrane and displacing SYTO® 9 fluorescence. The emission maxima of the SYTO® 9 and propidium iodide nucleic acid are 480 nm/500 nm and 480 nm/625 nm. Therefore, live and dead bacteria can be viewed separately or simultaneously by fluorescence microscopy with suitable optical filter sets. A BacLight bacterial viability kit has been used to determine the live and dead numbers of bacterial cells. C. taeanense was inoculated in a 10 ml R2A medium for 8 to 10 h and a 1 ml culture was filtered through 0.22 μm membrane filter (Millipore Inc.). After that, the bacteria captured on the filter were stained with a BacLight bacterial viability kit (Molecular Probes Inc.) in the dark for 20 min. The filter was fixed on the glass slide and sealed well and was analyzed by using a BX 51 microscope (Olympus, Japan) with Cy3 and a FITC filters to capture the micrographs. Finally, the numbers of the live and dead cells were calculated from the micrographs using ImageJ. 2.2.5 Phage-to-host ratio analysis Phage-to-host ratio (PHR) is a critical parameter which is an index of infectivity of the bacteriophage towards the host bacterium. To determine the optimal PHR, 1:1, 10:1, 24 102:1, 103:1, and 104:1 ratios of bacteriophages to the host, C. taeanense K-2, were initiated. Following 9 h of infection, the cultures were tested for final O.D., Bacteria Live/Dead ratio, and VLPs enumerated. A negative control without bacteriophage was also tested along with the above samples. 2.2.6 One-step growth curve In order to determine and construct the one-step growth curve of the Chryseobacterium taeanense bacteriophage, 1 mL of Chryseobacterium taeanense culture, with a population of 105, was transfered into the sterile polypropylene tubes and each sample in triplicate. Subsequently, 3×106 plaque forming units (PFU) of the isolated bacteriophage solution was added into each tube and incubated at 37 ℃ for 1 min, after which, the pallet was resuspended in a fresh R2A medium after being centrifuged by 6000g for 1 min at 4 ℃. Next, the resuspended pallet was added to the 1 L Chryseobacterium culture which was growing in the log growth phase. The chryseobacterium culture contained bacteriophages that were incubated at 37 􀀀 with starring and 1 ml infected culture was taken to test the PFU each 10 min. Each subsample from the set of triplicate samples at each time interval was divided into two equal halves, of which, one half was used to calculate the total PFU (free phage plus any infectious intracellular phage particles) after adding the chloroform and the other half was used to calculate the free PFU (extracellular/nonadsorbed). The counts obtained from triplicate values were averaged and were plotted to obtain the one-step growth curve (Ellis and 25 Delbrűck 1939). The burst size was calculated by dividing the number of virus-like particles released from the cell with the number of virus particles initially added. 2.3 Biofilm quantification and analysis 2.3.1 Biofilms formation and phage therapy The biofilms were formed on the glass slide which was inoculated into 5-slidesmailer (Fish scientific Inc.). After inoculation of 16, 54, and 84 days, the glass slides were divided into two parts, one without phage infection, the other part infected by phages. The phage titer of 1013 of phage solution was used to infect the biofilm overnight. Live and dead bacterial cell analysis was done using a Baclight bacterial viability kit and (Molecular Probes Inc.). In addition, the EPS of the biofilm was stained with Calcoflour White (ENG Scientific, Inc). 2.3.2 Biofilms quantification using epifluorescence microscopy The biofilms with the dye mixture were incubated in the dark for 30 min and were analyzed using a BX 51 microscope (Olympus, Japan) using Cy3, Cy4, and FITC filters to capture Live/dead cells and EPS, respectively. At each parameter tested, multiple pictures were overlayed and captured to quantify the percentages of Live/dead cells and EPS using BioImageL. For biofilm image quantification, the cell accounting program of the BiomageL will enumerate the total bacterial biomass and calculate distribution of the green and red biomass. The biomass identified as other colors will be labeled as ‘Not Segmented' (NS). 26 2.3.3 Scanning electron microscope images analysis The scanning electron microscope (SEM) uses a high-energy electron beam to generate a signal at the surface of solid specimens. A scanning electron microscope can receive signals from the electron sample and collect data from selected areas of the surface of the sample. Then a 2-dimensional image is created that displays spatial variations in these properties. The range of areas from 1 cm to 5 microns in width can be imaged in a scanning mode using conventional SEM techniques (magnification ranging from 20X to approximately 30,000X, spatial resolution of 50 to 100 nm). Also, SEM can do the point locations analysis, such as chemical compositions, crystalline structure, or surface texture determination. Compared with traditional microscopes, scanning electron microscopes have many advantages. SEM has a larger depth of field and higher resolution, which allows the specimens to be magnified at higher levels. SEM use electromagnets instead of lenses. Therefore, it has a larger degree of magnification. All in all, based on these advantages, the scanning electron microscope becomes one of the most useful instruments in the image analysis researching field. First, the glass slides with incubated biofilms were divided into two parts; one is the control part without phage infection, the other part infected with phages overnight. Then, the slides were fixed using an EM solution (1% paraformadehyde and 2.5% glutaraldehyde). The biofilms on the slide were washed with 0.l M cacodylate buffer (Ted Pella Inc., USA) twice for 5 min. Then, the slides were treated for 45 min with 27 postfix solution (2% OsO4 in 0.1 M cacodylate buffer). Subsequently, the biofilm was dehydrated in a gradient of (50% - 70% - 90 % -95% - absolute) alcohol solution and dried using hexamethyl disilazane (3 treatments). The slides were left overnight and Gold coated for 15 -30 sec, using 108􀀀 Sputter Coater (Ted Pella Inc., USA). Finally, the slides were analyzed in a Hitachi S-2460N scanning electron microscope and micrographs were taken at 20kV. CHAPTER 3 RESULTS AND DISCUSSION 3.1 Characterization of Chryseobacterium taeanense k-2 3.1.1 Isolation of antibiotic-resistant bacterium, Chryseobacterium taeanense k-2 Antibiotic-resistant bacterium, Chryseobacterium taeanense K-2, was isolated from the mixed liquor of a local full-scale wastewater treatment plant in the presence of kanamycin (40μg/ml final concentration). Three distinct bacterial colonies based on differentiation of color appeared on kanamycin-spiked R2A agar plates. C. taeanense k-2 formed visible yellowish colonies 2-3 mm in diameter with clear and entire circular edges after incubation for 16-24 h at 37°C (Figure 5). Pure culture of C. taeanense k-2 was obtained following repeated subculture of isolated colonies using the streak-plate method. C. taeanense k-2 was found to grow in the presence of kanamycin both on agar plates and liquid R2A media. The other two isolated colonies of antibiotic-resistant bacteria were investigated in a parallel research (by graduate students). It is not surprising that antibiotic-resistant bacteria were found in water samples from many U.S. municipal wastewater treatment plants, because bacteria that are resistant to chemically modified and synthesized antibiotics are widespread in the environment, as a1 a2 Figure 5. Agar plates (a1) without (a2) with Kanamycin (40 μg/ml) depicting isolated colonies (b) purified streaked subculture of Chryseobacterium taeanense K-2. revealed in previous studies (Ash et al. 2002). Most studies, however, were focused on screening for established antibiotic-resistant pathogens with a clinical perspective. This study, on the contrary, focused on random and unconventional bacteria that were resistant to antibiotics. The intention was to investigate the prevalence of antibiotic resistance among uncommon bacteria and natural flora of wastewater treatment systems. Limitations of this study were: i) a culture-based method was used to screen antibiotic-resistant bacteria and it is necessary to note that most bacteria in nature cannot be cultivated; ii) kanamycin was the only antibiotic used for screening, however, wastewaters are expected to harbor much more diverse bacteria that could be resistant to much more potent antibiotics. 3.1.2 16S rRNA gene phylogenetic analysis Determination of evolutionary relationships between organisms can be inferred by comparing the nucleotide or amino acid sequence. Smaller subunit ribosomal RNA (rRNA)-based fingerprinting is a well-established molecular tool to discern taxonomic affiliation and evolutionary relationships in prokaryotes. Carl Woese was the first microbiologist to use rRNA sequence analysis as a measure of microbial phylogeny and revolutionize our understanding of cellular evolution. The divergences among the microorganisms are depicted in the form of a phylogenetic tree using a treeing algorithm. A complete sequence of 16S rRNA (1390bp long) of the isolated antibiotic-resistant bacterium obtained from the sequencing was analyzed for homology match on BLAST (NCBI). The BLAST revealed 99% maximum identity and 99% coverage with Chryseobacterium taeanense strain NBRC 100863 (NCBI Accession # AB681269) and strain PHA3-4 (NCBI Accession # NR_043254). To our knowledge, there is no pathogenecity or antibiotic resistance attributed to the strains Chryseobacterium taeanense; however, other species belonging to the genera Chryseobacterium have been reported which cause infections in humans. Since the bacterium isolated in the present study was the second of the three colonies isolated on the kanamycin plate, it was named Chryseobacterium taeanense strain K-2. C taeanense belongs to the family Flavobacteriaceae under the phylum Bacteroidetes as retrieved from the (NCBI) taxonomy database. The phylogenetic tree based on the 16S-rRNA gene sequences was populated using the maximum likelihood algorithm in MEGA version 5.0. Evolutionary distance matrices for the maximum likelihood method were generated according to the Tamura-Nei distance model. The resultant maximum likelihood tree topology was evaluated by bootstrap analysis based on 5000 resample datasets. The analysis substitution type used was "Nucleotide". Phylogenetic affiliation of Chryseobacterium taeanense strain K-2 towards other species in the genus of Chryseobacterium, and other Flavobacteria, was evident from the lineage that appeared in the phylogram (Figure 6). Among the eight Chryseobacterium species presented, Chryseobacterium taeanense strain PHA3-4 was the closest neighbor sharing 99% gene sequence similarity. 3.1.3 Growth analysis of Chryseobacterium taeanense K-2 For testing and describing the bacteria growth curve, it is necessary to determine the number of cells representative of each single sample in different individual periods, because it is not easy to measure the culture's accurate cell number. Therefore, optical density (Absorbance @600nm) profile and plate count method via enumeration of Colony Forming Units (CFUs) was employed to deduce the growth curve of the Figure 6. Phylogram depicting the phylogenetic relationship of Chryseobacterium taeanense K-2 bacterium (Figure 7a and b). There was no lag phase recorded in the growth profile as the inoculum used was actively dividing cells grown in the same nutrient media. The exponential (log) phage was around 10 h long, after which the stationary phase appeared. As deduced from the growth curve, the culture's cell population doubling time is 2.21 h and the maximum specific growth rate of Chryseobacterium taeanense K-2 was around 0.022/h (Figure 7c). Chryseobacteriumlineage Figure 7. Growth curve of Chryseobacterium taeanense K-2 obtained based on a) Optical density and b) Plate count method; c) depicts maximum specific growth rate a Chryseobacterium taeanense K-2 growth curve Time(h) 0 5 10 15 20 25 CFUs/ml 0 5e+7 1e+8 2e+8 2e+8 3e+8 3e+8 b y = 0.0221x + 0.0461 R² = 0.9351 0 0.05 0.1 0.15 0.2 0.25 0.3 0 2 4 6 8 10 O.D.(A600) Time(h) Time(h) 0 5 10 15 20 25 O.D.(A600) 0.00 0.05 0.10 0.15 0.20 0.25 0.30 c a 3.1.4 DAPI (4',6-diamidino-2-phenylindole) staining analysis DAPI binds strongly to the A-T rich region on the DNA and is therefore used extensively in fluorescence microscopy. DAPI can pass through an intact cell membrane; therefore, it can be used to stain both live and fixed cells, though it passes through the membrane less efficiently in live cells and therefore, the effectiveness of the stain is lower. DAPI staining reveals that the morphology of the Chryseobacterium taeanense K-2 is coccoid 0.5-1.0 μm cell-length (Figure 8) Myung et al. (2006) reported similar morphology of Chryseobacterium taeanense. 3.1.5 Gram staining During the staining process, the Crystal Violet (hexamethyl-para-rosaniline chloride) interacts with aqueous KI-I2 via a simple anion exchange to produce a chemical precipitate (Brock et al. 2004). The Gram-positive bacterial cell wall has more lipids than Gram-negative bacterial cell walls. Thus, during the distaining process, alcohol will dissolve the lipids presence in the membrane and cell wall. While the Gram-negative bacteria cell has thinner peptidoglycan, it does not have enough ability to retain dye-iodine. Hence, Gram-negative bacterial cells readily get decolorized. Moreover, the Gram-positive cells will dehydrate and the cell wall will shrink by ethanol treatment. Therefore, the dye iodine complex will stay inside the thick peptidoglycan layer and does not get decolorized. It is easy to narrow the scope of the bacteria to identify the isolated Figure 8. Micrographs obtained from DAPI analysis of Chryseobacterium taeanense K-2 from the log-phage culture. bacteria by Gram staining. Moreover, Gram reactivity can be used as a reference for the use of antibiotics in the medical field. In this study, the Gram staining analysis inferred that the Chryseobacterium taeanense K-2 was Gram-negative. Phase-contrast micrographs (Figure 9) confirmed the morphology (0.1-0.5 μm coccoid) revealed through DAPI staining of Chryseobacterium taeanense. Figure 9. Phase-contrast micrographs of Gram-negatively stained Chryseobacterium taeanense k-2 3.2 Characterization of bacteriophage of the host of Chryseobacterium taeanense K-2 3.2.1 Isolation of bacteriophage Lytic bacteriophages specific to Chryseobacterium taeanense K-2 were successfully isolated from the mixed liquor sample obtained from the full-scale wastewater treatment plant (Central Valley Water Reclamation Facility-CVWRF). The repeated plaque assay technique produced clear plaques that were visible on 1% top agar when concentration phage-stock was used. This finding clearly implies that the possibility of finding a lytic phage infecting a host bacterium from the same habitat is significantly higher. The agar plate (Figure 10) contained several clear plaques instead of cloudy or bull-eyed plaques (Ellis, and Delbru¨ ck, 1939; Uchiyama et al. 2009) obtained after serial infection of Chryseobacterium taeanense K-2, suggesting the plaques were formed by lytic but NOT lysogenic phages. Three distinct plaques based on size of plaques viz. CTP-1 (0.5-1mm diameter), CTP-2 (0.1-0.2 mm diameter), and CTP-3 (0.1-0.2 mm diameter) were chosen for subsequent screen. Repeated plaque assay found that CTP-3 can have consistent infectivity and a significantly higher rate of infection. Therefore, CTP-3 was chosen for further study. 3.2.2 Viral enumeration using epifluorescence microscopy (EFM) Figure 11 is the epifluorescent micrograph showing the distribution of Virus-Like-Particles (VLPs) present in the phage-stock of the lytic bacteriophage. The b Figure 10. Agar plate containing clear plaques. Figure 11. Virus-like particles (green dots) pointed by yellow arrows in the epifluorescent micrographs. yellow arrow in the figure points to the VLPs (distinct green dots), suspended in the phage-stock stained with SYBR Gold. The titer of the Chryseobacterium taeanense K-2 phage-stock obtained from progressive infection was calculated to be 1.11E+14 PFU/mL. EFM micrographs can provide a synoptic picture of Virus-like particles for their quantification, distribution, and abundance in a given samples. Due to the existence of single-stranded DNA or RNA viruses (which do not bind nucleic acid stains), it is not always accurate to estimate the abundance of virus using EFM. Furthermore, environmental samples may contain particles which can nonspecifically bind these nucleic-acid stains, resulting in false-positive enumeration. Therefore, direct plating plaque assay in associated with TEM-based and EFM-based VLP enumeration is recommended for the quantification of the virus. Despite these limitations, EFM-based technique can provide a rapid snapshot of VLP number in a sample. 3.2.3 Purification of lytic bacteriophage CsCl density gradient centrifugation of the phage-stock obtained from a smaller infection volume did not result in a visible band of purified virus at the interface of 1.4g/l and 1.6g/l CsCl. Therefore, a progressive infection of Chryseobacterium taeanense K-2 with the phage-stock using a final volume of 1L 10 h old culture was used. Following overnight centrifugation, the viral pellet resuspended in the SMG buffer was used for CsCl density gradient centrifugation which revealed a distinct band (Figure 12). Purified phage-stock obtained after the dialysis of this band was used for TEM analysis. CsCl purified phages band 1.4 g/l CsCl (interface) 1.6 g/l CsCl Figure 12. The bacteriophages' band after Cesium chloride density gradient purification 3.2.4 Transmission electron microscopy (TEM) analysis TEM analysis shows that the isolated lytic bacteriophages infecting Chryseobacterium taeanense K-2 have icosahedral geometry (Figure 13). The lytic bacteriophage had a very short noncontractile tail and a hexagonal head. The total length of the phage ranged from 90 to 100 nm. The diameter of the hexagonal head was 85 nm. The tail is noncontractile without subterminal fibers. The maximum length is ~23 nm. Table 1 enlists the multiple dimensions of the phages recorded in the TEM microgrpahs. Based on the morphology revealed by the TEM-micrographs (Figure 13 a and b), the isolated bacteriophages belong to the Podoviridae family. These viruses have been reported to be lytic phages rather than lysogenic. Some TEM micrographs also showed ellipsoid morphology of the bacteriophage. The possible reason for this morphology could be ‘‘empty'' or "ghost" virus, where the major axis of the ellipsoid can be 40% Figure 13. TEM micrographs of the lytic bacteriophage infecting Chryseobacterium taeanense K-2 200 nm Hexagonal Head Tail b a Head Tail 200 nm a Table 1: The bacteriophage's size obtained from TEM images longer than the minor axis (Pease et al. 2009). The possible reason for the deformation of the phage could be a result of fixing and staining preparation steps. Similar defects in the viral morphology were reported following the VP2-directed packaging of green fluorescent protein into MPV VLPs (Boura et al. 2005). 3.2.5 One-step growth curve The replication of bacteriophages during the infection of the host bacterium, Chryseobacterium taeanense k-2, was investigated following the method used for Haliscomenobacter hydrossis lytic bacteriophages (Kotay et al. 2011). The One-step growth curve of the lytic bacteriophage is depicted in Figure 14. The latent period of C. taeanense k-2 bacteriophage was found to be around 10 min and the eclipse period and latent period of the virus were synchronized. Latent and eclipse periods are very important parameters for the process of biocontrol. The latent time period spans from the point of phage adsorption to the point at which host lysis occurs and the eclipse time Name image number Head(nm) Tail(nm) Length(nm) CTP1 123401 77 0 77 CTP1 123402 99 8 107 CTP1 123403 84 6 92 CTP1 123404 93 0 93(89) CTP2 123393 77 15 92 CTP2 123394 77 15 92 CTP2 123396 77 0 77 CTP3 123397 69 23 92 CTP3 123398 77 0 77 CTP3 123399 84 0 84 CTP3 123400 77 0 77 Figure 14. One-step growth curve of lytic bacteriophage infecting Chryseobacterium taeanense K-2 period spans from the point of phage adsorption to the point at which the first phage progeny have matured within an infected cell. Following the curve obtained from chloroform treated samples (Figure 14), the trend was not typical as that of previously known phages like T4. It had no initial stable stage which was atypical for any infective virus previously demonstrated (Furness and Fraser 1962). When the bacteriophages infected the cells, they increased immediately until a peak of 1100 PFUs/ml was reached at 40 h. On the other hand, the chloroform untreated curve shows that the initial free bacteriophages was 420 PFUs/ml. In the first 10 min, the bacteriophage numbers were Time/mins 0 20 40 60 80 100 120 PFU/ml 0 200 400 600 800 1000 1200 Chloroform Treated Untreated PFUs/ml Time(minute) decreasing due to attachment of the host cells. After 10 min, the bacteriophage numbers were increasing, because of the first lytic cycle completion. The burst size was calculated to be 18±2 PFU/infected cell which was comparable to that of previously reported lytic phages (Kotay et al. 2011). 3.2.6 Phage-to-host ratio analysis Phage-to-host ratio is critical information necessary for engineering biocontrol applications. Some studies show that a higher phage-to-host ratio (PHR) can cause more than 90% mortality of H. hydrossis and VBR lower than 1:1000, resulting in mortality of less than 10% (Kotay et al. 2011). On the other hand, higher PHR does not always represent optimum lytic effect of the bacteriophage on the targeted hosts. For example, over death of filamentous bacteria could cause negative effects on the biomass settleability. Activated sludge bioreactor consists of a complex community of a variety of organisms (Kotay et al. 2011). In general, during the infecting and multiplying processing, bacteriophages have a critical threshold ratio target to the host populations. According to the curve of VLPs versus time (Figure 15) and Live/Dead cells ratio versus time (Figure 16), PHR of 100:1 was found to be most effective, where 98.7% of the host bacterium, Chryseobacterium taeanense K-2, were dead. In addition, the final viral count was 7.71E+7 VLPs/ ml after 9 h infection (Table 2.). However, the daughter bacteriophage numbers were found to be decreasing from 4.42E+08 and 1.62E+10 VLPs/ml to 2.65E+8 and 7.09E+8 VLPs/ml, in case of PHRs 1000:1 and 10000:1, Figure 15. Profile of VLPs at different PHR Time/h 0 2 4 6 8 10 Live/Dead cells ratio 0 1 2 3 4 5 6 PHR 1:1 PHR 10:1 PHR 100:1 PHR 1000:1 PHR 10000:1 Time(h) Figure 16. Profile of Live/Dead Cells ratios at different PHR Time/h 0 2 4 6 8 10 VLPs/ml 1e+5 1e+6 1e+7 1e+8 1e+9 1e+10 1e+11 PHR 1:1 PHR 10:1 PHR 100:1 PHR 1000:1 PHR 10000:1 Time(h) Table 2: Effect of initial phage/host ratio on bacteriolysis and VLP count respectively. The reduced efficacy at higher phage to host ratio may be because of competitive interference between bacteriophages (Delbrück, 1940); repeated initiation of replication can lyse the bacterium. This study, however, was limited to plantonic population of Chryseobacterium taeanense K-2. Higher PHR might be necessary in case of complex populations of the host like sludge flocs which contain a mixed community of bacteria and biofilms which could have multiple layers of host bacteria. Nevertheless, bacteriophage-based biocontrol is a viable strategy to regulate Chryseobacterium taeanense K-2 through an application of optimal PHR. Bacteriophages can also be used in an integrated approach in combination with other biocidal chemicals; however, it is not a panacea to solve all biofilm problems or biofouling in wastewater engineering. Bacteriophage-based biocontrol needs a lot of Initial Phage/ Host ratio Final O.D. Final bacteria Live/Dead Ratio Final VLPs/ml 1:1 0.172 0.1164 3.95E+07 10:1 0.183 0.0581 4.54E+07 100:1 0.193 0.013 7.71E+07 1000:1 0.19 0.709 2.65E+08 10000:1 0.171 0.0299 7.09E+08 Phage/Host Ratio and Final VLPs after 9 hrs infection preparatory research to determine the most efficacious dosing strategies and the most effective combinations of phages targeting bacterial hosts. 3.3 Biofilms disintegration using phage therapy 3.3.1 Phage therapy for biofilm Figures 17 a1 and a2 show the 16-day-old biofilm grown on the glass slides stained by Live/Dead BacLight™ and Calcoflour White. The micrographs indicated the morphology of biofilm prior to phage application. The morphologies present in the cells are growing and contacting as "microcolonies" in which the diameter is ranging from 1 to 5 μm. Very faint volumes of extra-polymeric substance (blue fluorescent) were observed. Microcolonies have been known as one of the steps in the development of a biofilm. On the other hand, no "microcolonies" were observed in the biofilm treated with 1013 PFUs/ml phages overnight (Figure 17 b1 and b2). This phenomena may attributed to the disruption of the microcolonies following infection by the lytic phage. Biofilms of the age 54 days were stained in a similar way. Figure 18 indicated that the morphology of biofilm has more cells and complex structure. Micrographs in Figure 18 a1 and a2 show the distributions of live (green) / dead (red) cells and a relatively higher volume/network of EPS than compared to 16-day-old biofilm. Following infection, integrity and uniformity of the biofilm was no longer observed (Figure 18 b1 and b2). Also, the distinct cell structure of the Chryseobacterium taeanense K-2 was lost, and cell lysis was evident from the red colored lumps (Figure 18 b1 and b2). This finding further Figure 17. Epifluorescent micrographs of 16-day biofilm (a1 & a2) before and (b1 & b2) after infection with phage. strengthens the earlier speculation of the feasibility of bacteriophages-based biocontrol of biofilms. On the contrary, the micrographs obtained from 84-day-old biofilm showed different cellular morphology and higher biovolume of EPS (Figure 19 a1 and a2). Differential cellular morphology of bacteria within a biofilm has been reported earlier. Following application of phage, there were no distinct cells lysis or EPS disintegration observed. It may be speculated that phage was not able to penetrate relatively older biofilm. Biofilms are known to gain complexity, thickness, and impermeability with age a1 b1 b2 a2 Figure 18. Epifluorescent micrographs of 54-day biofilm (a1 & a2) before and (b1 & b2) after infection with phage. 5.0 μ m 5.0 μm a1 a2 b1 b2 5.0 a1 a2 b1 b2 Figure 19. Epifluorescent micrographs of 84-day biofilm (a1& a2) before and (b1 & b2) after infection with phage. and therefore could resist phage infection. Further investigation will be necessary to understand this phenomenon. 3.3.2 Biofilms quantification using epifluorescence micrographs and BioImageL BioImageL is an image analysis software widely used for investigation of microbial biofilms. The basic principle is based on in situ color segmentation. BioImageL has the capability to identify and distinguish green (Live) and red (Dead) subareas and calculate different architectural parameters (Figure 20, Figure 21, and Figure 22). Figure 20. Effect of phage therapy on % Live cells in the biofilm of different ages Figure 21. Effect of phage therapy on % Dead cells in the biofilm of different ages Biofilm age(day) % of Dead cells 0 20 40 60 80 100 120 Before phage therapy After phage therapy 16 54 84 Biofilm age (day) % of Live cells 0 10 20 30 40 50 Before phage therapy After phage therapy 16 54 84 Figure 22. Effect of phage therapy on % EPS in the biofilm of different ages The percentage of live cells in 16-day-old biofilm decreased from 39.12% to 2.04% following infection with the lytic phage. The percentage of dead cells increased from 57.55% to 96.54% and the percentage of EPS decreased from 3.32% to 1.23% after overnight infection of 16-day biofilm with 1013 PFUs/ml. After overnight phages infection, the percentage of live cells in 54-day-old biofilm decreased from 15.67% to 13.79%, the percentage of dead cells decreased from 63.05% to 53.85%, and the percentage of EPS decreased from 21.28% to 32. 36%. After overnight phages infection, the percentage of live cells of 84-day biofilm decreased from 25.97% to 7.58%, the percentage of dead cells increased from 71.17% to 90.59%, and the percentage of EPS decreased from 2.84% to 1.82%. Based on the T-Test analysis on biofilm quantification of 16-day-old biofilm before and after phage infection, the P value of the percentages of Live/Dead cells and EPS Biofilm age(day) % of EPS 0 10 20 30 40 50 Before phage therapy After phage therapy 16 54 84 were found to be 0.002, 0.002, and 0.0493, and for 54-day biofilm were found to be 0.463, 0.213, and 0.0095. For 84-day biofilm, they were found to be 1.6857E-08, 1.4486E-08, and 0.063. The possible reason that P value was higher for 16-day and 54-day biofilm (0.463 and 0.213) could be attributed to the fact that there was no adjustment available in the software for the system error even though the noise factors were reduced in the biofilms micrograph during processing. Likewise, further studies are needed for Confocal laser scanning microscopy images analysis to determine the EPS distribution and quantify the thickness of biofilms. 3.3.3 Scanning electron microscope images analysis Scanning electron microscope (SEM) analysis of biofilm before and after the infection was performed to understand the extent and mechanism of infection at a high resolution. Figure 23 a1 and a2 show the surface structure of the 16-day-old biofilms prior to the phage infection. The structure reveals bacterial cells impregnated within a smooth EPS matrix. Figure 23 b1 and b2 show the surface structure of the 16-day-old biofilms after the phage infection. Hole-like structures were observed throughout the surface of the biofilm. These holes were consistent across the surface of the biofilm which may have emerged due to phage infection. Similar findings were observed in the SEM micrographs of 54-day and 84-day-old biofilms (Figures 24 and 25). Figure 23. SEM micrographs of 16-day-old biofilm before (a1 & a2) and after (b1 & B2) phage infection Figure 24. SEM micrographs of 54-day-old biofilm before (a1 & a2) and after (b1 & B2) phage infection a1 b1 b2 a2 a1 b2 a2 b1 Figure 25. SEM micrographs of 84-day-old biofilm before (a1 & a2) and after (b1 & B2) phage infection a1 b1 b2 a2 CHAPTER 4 CONCLUSIONS 4.1 Conclusions from study The goal of this research was to demonstrate bacteriophage-mediated biocontrol of biofilm formed by antibiotic resistant bacterium isolated from a full-scale wastewater treatment plant. Based on the results obtained, the following conclusions can be made: 1. The antibiotic resistant bacterium, Chryseobacterium taeanense K-2, was isolated from a wastewater treatment plant. This is the first report on the finding of an antibiotic-resistant strain of Chryseobacterium taeanense. The study is also one among the few to have isolated an unconventional antibiotic-resistant bacterium that has not previously been associated with pathogenesis. 2. A lytic bacteriophage specific to Chryseobacterium taeanense K-2 existed in the wastewater treatment system and was successfully isolated. The burst size was calculated to be 18±2 PFU/infected cells. Suitable Phage-to-Host ratio for infection was found to be 100:1 which caused the 98.7% death of host planktonic cells. 3. The results of biofilm quantification show that the lytic bacteriophage was able to disintegrate 16 and 54-day-old biofilms of Chryseobacterium taeanense K-2, but 84-day-old biofilm did not show a significant difference. APPENDIX Table 3: Live/dead cells and EPS distribution of the biofilm of the age of 16 days without phages infected Image Number Noise Reducin g Factor Field Size (um2) Cells Area (um2) Cells Area covered (%) Green Cells Area (um2) Green Cells (%) Red Cells Area (um2) Red Cells (%) EPS (um2) EPS (%) 1 0.15 45029 5176 11.5 253 4.9 4683 90.5 240 4.6 2 0.09 45029 2926 6.5 214 7.3 2693 92 19 0.6 3 0.1 45029 3634 8.1 156 4.3 3388 93.2 90 2.5 4 0.23 45029 5432 12.1 2988 55 2331 42.9 113 2.1 5 0.26 45029 18844 41.8 15188 80.2 3414 18.1 313 1.7 6 0.29 45029 2767 6.1 1333 48.2 1224 44.2 209 7.6 7 0.2 45029 4617 10.3 590 12.8 3024 65.5 1003 21.7 8 0.04 45029 1897 4.4 308 15.5 1655 83.7 15 0.8 9 0.13 45029 19174 42.6 15801 82.4 2400 12.5 974 5.1 10 0.01 45029 3158 7 1623 51.4 1533 48.5 2 0.1 11 0.31 45029 746 1.7 214 28.7 490 66.9 33 4.4 12 0.39 45029 4333 9.6 3462 79.9 793 18.3 78 1.8 13 0.16 45029 822 1.8 566 68.8 251 30.5 5 0.6 14 0.09 45029 194 0.4 93 47.9 100 51.5 1 0.6 15 0.09 45029 162 0.4 56 34.7 105 64.8 1 0.4 16 0.05 45029 419 0.9 91 21.7 328 78.1 1 0.2 17 0.05 45029 826 1.8 41 5 782 94.7 3 0.3 18 0.05 45029 398 0.9 124 31.1 273 68.5 2 0.4 19 0.03 45029 1980 4.4 1131 57.1 843 42.6 6 0.3 20 0.29 45029 163 0.4 68 41.8 82 50.5 12 7.6 21 0.15 45029 992 2.2 28 2.8 963 97.1 1 0.1 22 0.05 45029 1353 3 509 37.7 831 61.4 12 0.9 23 0.09 45029 644 1.4 37 5.8 606 94.2 0 0.1 24 0.06 45029 938 2.1 558 60.1 341 36.8 29 3.1 25 0.03 45029 525 1.2 420 80 86 16.3 19 3.7 26 0.03 45029 2000 4.4 152 7.6 1844 92.2 4 0.2 27 0.03 45029 656 1.5 344 52.4 301 45.9 11 1.7 28 0.06 45029 989 2.2 590 59.7 386 39 13 1.3 29 0.01 45029 462 1 299 64.6 147 31.7 17 3.6 30 0.32 45029 2371 5.3 1490 62.8 296 12.5 585 24.7 31 0.07 45029 1251 2.8 6 0.5 1244 99.5 0 0 Average 2898.355 6.445161 1572.032 39.11935 1207.645 57.55161 122.9355 3.316129 Biofilm of the Age of 16 days (control) Table 4: Live/dead cells and EPS distribution of the biofilm of the age of 16 days with phages infected Image Number Noise Reducin g Factor Field Size (um2) Cells Area (um2) Cells Area covered (%) Green Cells Area (um2) Green Cells (%) Red Cells Area (um2) Red Cells (%) EPS (um2) EPS (%) 1 0.07 45029 7407 16.4 27 0.4 6982 94.3 398 0.4 2 0.07 45029 8381 18.6 8 0.1 8287 98.9 86 1 3 0.11 45029 869 1.9 33 3.8 821 94.6 14 1.6 4 0.11 45029 171 0.4 7 4.1 156 91.6 7 4.3 5 0.01 45029 435 1 15 3.5 420 96.4 0 0 6 0.27 45029 3357 7.5 203 6.1 3054 91 100 3 7 0.21 45029 10249 22.8 2187 21.3 7547 73.6 515 5 8 0.24 45029 1483 3.3 55 3.7 1353 91.3 75 5 9 0.35 45029 2831 6.3 53 1.9 2683 94.8 95 3.3 10 0.34 45029 6694 14.9 476 7.1 6065 90.6 152 2.3 11 0.1 45029 5245 11.6 90 1.7 5147 98.1 9 0.2 12 0.07 45029 5576 12.4 26 0.5 5538 99.3 12 0.2 13 0.06 45029 3894 8.6 30 0.8 3863 99.2 1 0 14 0.07 45029 3227 7.2 21 0.6 3200 99.1 7 0.2 15 0.07 45029 5385 12 62 1.2 5307 98 15 0.3 16 0.05 45029 5424 12 4 0.1 5415 99.8 5 0.1 17 0.05 45029 5765 12.8 22 0.4 5724 99.3 19 0.3 18 0.05 45029 1612 3.6 5 0.3 1606 99.7 0 0 19 0.05 45029 4421 9.8 12 0.3 4409 99.7 0 0 20 0.05 45029 3212 7.1 10 0.3 3197 99.5 5 0.2 21 0.09 45029 1416 3.1 22 1.6 1392 98.3 1 0.1 22 0.05 45029 11346 25.2 21 0.2 10425 91.9 900 7.9 23 0.05 45029 3813 8.5 8 0.2 3804 99.8 1 0 24 0.05 45029 6074 13.5 24 0.4 6044 99.5 6 0.1 25 0.05 45029 6256 13.9 6 0.1 6247 99.8 4 0.1 26 0.05 45029 7520 16.7 33 0.4 7481 99.5 6 0.1 27 0.05 45029 3470 7.7 37 1.1 3425 98.7 8 0.2 28 0.05 45029 2884 6.4 0 0 2879 99.8 5 0.2 29 0.05 45029 6133 13.6 9 0.1 6122 99.8 2 0 30 0.05 45029 1714 3.8 5 0.3 1708 99.6 1 0 31 0.05 45029 3381 7.5 21 0.6 3292 97.4 67 2 Average 4504.677 10.00323 113.9355 2.03871 4309.452 96.54516 81.16129 1.229032 Biofilm of the Age of 16 days (infected) Table 5: Live/dead cells and EPS distribution of the biofilm of the age of 54 days without phages infected Image Number Noise Reducing Factor Field Size (um2) Cells Area (um2) Cells Area covered (%) Green Cells Area (um2) Green Cells (%) Red Cells Area (um2) Red Cells (%) EPS (um2) EPS (%) 1 0.09 45029 2358 5.2 1348 57.1 630 26.7 380 16.1 2 0.55 45029 2793 6.2 380 13.6 1921 68.8 491 17.6 3 0.2 45029 4950 11 1982 40 1620 32.7 1348 27.2 4 0.16 45029 4213 9.4 312 7.4 3295 78.2 605 14.4 5 0.14 45029 10614 23.6 1984 18.7 4053 38.2 4578 43.1 6 0.23 45029 5181 11.5 243 4.7 4557 88 381 7.3 7 0..1 45029 785 1.7 106 13.5 435 55.5 244 31 8 0.22 45029 8983 19.9 884 9.8 5999 66.8 2100 23.4 9 0.2 45029 45029 100 8990 19.7 29318 64.1 7407 16.2 10 0.17 45029 8107 18 5045 62.2 2265 27.9 797 9.8 11 0.13 45029 14230 31.6 904 6.4 10971 77.1 2355 16.6 12 0.21 45029 31322 69.6 12489 39.9 12331 39.4 6502 20.8 13 0.11 45029 45044 100 0 0 44392 98.6 647 1.4 14 0.13 45029 7918 17.6 408 5.2 7435 93.9 75 0.9 15 0.17 45029 2833 6.3 382 13.5 1129 39.9 1322 46.7 16 0.16 45029 4387 9.7 343 7.8 3007 68.6 1036 23.6 17 0.19 45029 10525 23.4 164 1.6 9424 89.5 937 8.9 18 0.23 45029 12010 26.7 4483 37.3 5849 48.7 1678 14 19 0.24 45029 14137 31.4 6274 44.4 5311 37.6 2551 18 20 0.39 45029 4818 10.7 1051 21.8 3042 63.1 725 15 21 0.34 45029 2611 5.8 533 20.4 583 22.3 1494 57.2 22 0.47 45029 1466 3.3 279 19 795 54.3 392 26.7 23 0.29 45029 11534 25.6 6 0.1 11082 96.2 435 3.8 24 0.12 45029 7549 16.8 1211 16 3330 44.1 3008 39.9 25 0.11 45029 31062 69 3330 10.7 15303 49.3 12429 40 26 0.19 45029 8959 19.9 54 0.6 8709 97.2 196 2.2 27 0.29 45029 14916 33.1 6 0 13483 90.4 1428 9.6 28 0.18 45029 5572 12.4 438 7.9 3384 60.7 1750 31.4 29 0.27 45029 15994 35.5 4111 25.7 3871 24.2 8013 50.1 30 0.29 45029 45413 100 0 0 35905 79.1 9507 20.9 31 0.25 45029 26779 59.5 553 2.1 26007 97.1 220 0.8 Average 13657.8 30.30667 1898.167 15.66667 9293.533 63.05 2488.367 21.28333 Biofilm of theAge of 54 days (control) Table 6: Live/dead cells and EPS distribution of the biofilm of the age of 54 days with phages infected Image Number Noise Reducing Factor Field Size (um2) Cells Area (um2) Cells Area covered (%) Green Cells Area (um2) Green Cells (%) Red Cells Area (um2) Red Cells (%) EPS (um2) EPS (%) 1 0.2 45029 12898 28.6 332 2.6 9763 75.7 2803 21.7 2 0.05 45029 22441 49.8 3913 17.4 11384 50.7 7145 31.8 3 0.2 45029 2248 5 176 7.9 1639 72.9 432 19.2 4 0.07 45029 13970 31 5426 38.8 3894 27.9 4649 33.3 5 0.11 45029 8158 18.1 5988 73.4 502 6.2 1668 20.4 6 0.16 45029 3414 7.6 75 2.2 2820 82.6 519 15.2 7 0.23 45029 5313 11.8 69 1.3 4290 80.7 954 18 8 0.15 45029 6187 13.7 621 10 1383 22.4 4182 67.6 9 0.29 45029 4433 9.8 28 0.6 1889 42.6 2516 56.8 10 0.47 45029 6928 15.4 2743 39.6 2574 37.2 1611 23.2 11 0.19 45029 8340 18.5 75 0.9 7354 88.2 910 10.9 12 0.28 45029 29845 66.3 2 0 28046 94 1797 6 13 0.11 45029 1971 4.4 80 4.1 1421 72.1 469 23.8 14 0.19 45029 8248 18.3 651 7.9 6640 80.5 957 11.6 15 0.07 45029 8371 18.6 2276 27.2 1892 22.6 4204 50.2 16 0.11 45029 14042 31.2 3665 26.1 5545 39.5 4832 34.4 17 0.18 45029 7110 15.8 117 1.6 5842 82.2 1151 16.2 18 0.18 45029 10085 22.4 570 5.7 6801 67.4 2714 26.9 19 0.2 45029 6694 14.9 629 9.4 3909 58.4 2156 32.2 20 0.2 45029 3799 8.4 7 0.2 1756 46.2 2037 53.6 21 0.25 45029 4632 10.3 1911 41.3 513 11.1 2208 47.7 22 0.11 45029 670 1.5 168 25.1 136 20.4 365 54.5 23 0.19 45029 15305 34 64 0.4 12030 78.6 3211 21 24 0.18 45029 5255 11.7 110 2.1 2562 48.8 2583 49.1 25 0.09 45029 8817 19.6 1282 14.5 3935 44.6 3600 40.8 26 0.14 45029 13323 29.6 117 0.9 10964 82.3 2242 16.8 27 0.14 45029 14787 32.8 2779 18.8 7472 50.5 4536 30.7 28 0.16 45029 5668 12.6 294 5.2 2820 49.8 2554 45.1 29 0.11 45029 5042 11.2 266 5.3 1227 24.3 3549 70.4 30 0.15 45029 9083 20.2 2115 23.3 5001 55.1 1967 21.7 31 45029 Average 8902.567 19.77 1218.3 13.79333 5200.133 53.85 2484.033 32.36 Biofilm of theAge of 54 days (infected) Table 7: Live/dead cells and EPS distribution of the biofilm of the age of the 84 days without phages infected Image Number Noise Reducing Factor Field Size (um2) Cells Area (um2) Cells Area covered (%) Green Cells Area (um2) Green Cells (%) Red Cells Area (um2) Red Cells (%) EPS (um2) EPS (%) 1 0.36 45029 26331 58.5 405 1.5 25239 95.9 687 2.6 2 0.11 45029 12944 28.7 1857 14.3 10982 84.8 105 0.8 3 0.25 45029 27933 62 13039 46.7 14298 51.2 596 2.1 4 0.13 45029 26207 58.2 2870 11 23193 88.5 144 0.5 5 0.34 45029 23386 51.9 6059 25.9 16709 71.4 618 2.6 6 0.23 45029 33829 75.1 615 1.8 32719 96.7 496 1.5 7 0.41 45029 23716 52.7 12271 51.7 8946 37.7 2498 10.5 8 0.08 45029 40311 89.5 28619 71 11171 27.7 522 1.3 9 0.42 45029 37527 83.3 19321 51.5 16281 43.4 1925 5.1 10 0.11 45029 8478 18.8 355 4.2 7871 92.8 252 3 11 0.15 45029 27932 62 851 3 25679 91.9 1402 5 12 0.03 45029 21223 47.1 271 1.3 20744 97.7 208 1 13 0.12 45029 31103 69.1 900 2.9 29793 95.8 410 1.3 14 0.14 45029 5467 12.1 50 0.9 5417 99.1 0 0 15 0.44 45029 34903 77.5 19500 55.9 14494 41.5 909 2.6 16 0.12 45029 45029 100 327 0.7 42766 94.5 2164 4.8 17 0.28 45029 42687 94.8 4553 10.7 35184 82.4 2950 6.9 18 0.03 45029 39305 87.3 20223 51.5 18789 47.8 293 0.7 19 0.18 45029 41011 91.1 18047 44 22215 54.2 748 1.8 20 0.23 45029 41280 91.7 18823 45.6 21521 52.1 936 2.3 21 0.02 45029 35687 79.3 13281 37.2 21893 61.3 514 1.4 22 0.13 45029 35246 78.3 5751 16.3 29086 82.5 408 1.2 23 0.06 45029 30751 68.3 7546 24.5 22298 72.5 907 2.9 24 0.13 45029 10872 24.1 2964 27.3 7820 71.9 88 0.8 25 0.17 45029 43561 96.7 12454 28.6 29809 68.4 1298 3 26 0.01 45029 35730 79.3 19251 53.9 13599 38.1 2880 8.1 27 0.09 45029 42202 93.7 2792 6.6 39138 92.7 272 0.6 28 0.1 45029 11460 25.5 1928 16.8 9171 80 362 3.2 29 0.14 45029 29044 64.5 11901 41 15904 54.8 1238 4.3 30 0.23 45029 23357 51.9 15306 65.5 6988 29.9 1063 4.6 31 0.24 45029 25446 56.5 636 2.5 24369 95.8 440 1.7 32 0.42 45029 13827 30.7 4753 34.4 8359 60.5 715 5.2 33 0.25 45029 26785 59.5 1654 6.2 24970 93.2 161 0.6 Average 28926.36 64.23333 8156.758 25.96667 19921.67 71.17273 854.8182 2.848485 Biofilm of the Age of 84 days (control) Table 8: Live/dead cells and EPS distribution of the biofilm of the age of the 84 days with phages infected Image Number Noise Reducing Factor Field Size (um2) Cells Area (um2) Cells Area covered (%) Green Cells Area (um2) Green Cells (%) Red Cells Area (um2) Red Cells (%) EPS (um2) EPS (%) 1 0.17 45029 44860 99.6 4 0 43963 98 893 2 2 0.05 45029 22272 49.5 2898 13 18086 81.2 1288 5.8 3 0.35 45029 29591 66.7 2515 8.5 26274 88.8 802 2.7 4 0.32 45029 14722 32.7 189 1.3 14154 96.1 380 2.6 5 0.18 45029 7500 16.7 351 4.7 7135 95.1 15 0.2 6 0.23 45029 9894 22 74 0.8 9105 92 715 7.2 7 0.32 45029 31050 69 49 0.2 29869 96.2 1133 3.6 8 0.1 45029 45088 100 142 0.3 43275 96 1671 3.7 9 0.1 45029 29385 65.3 94 0.3 29256 99.6 35 0.1 10 0.17 45029 42430 94.2 2871 6.8 38678 91.2 881 2.1 11 0.12 45029 42241 93.8 2750 6.5 39316 93.1 175 0.4 12 0.17 45029 22817 50.7 99 0.4 22616 99.1 103 0.5 13 0.14 45029 23575 52.4 186 0.8 22510 95.5 879 3.7 14 0.21 45029 18967 42.1 154 0.8 18721 98.7 92 0.5 15 0.07 45029 14021 31.1 295 2.1 13547 96.6 179 1.3 16 0.08 45029 12712 28.2 643 5.1 12051 94.8 18 0.1 17 0.22 45029 32247 71.6 11 0 31099 96.4 1138 3.5 18 0.14 45029 35014 77.8 1838 5.3 32768 93.6 407 1.2 19 0.13 45029 20479 45.5 1060 5.2 19401 94.7 18 0.1 20 0.11 45029 39109 86.9 148 0.4 38756 99.1 204 0.5 21 0.09 45029 34818 77.3 937 2.7 33433 96 449 1.3 22 0.17 45029 38211 84.9 3331 8.7 33950 88.8 939 2.5 23 0.12 45029 45365 100 25031 55.2 19893 43.9 440 1 24 0.15 45029 35636 79.1 776 2.2 34187 95.9 673 1.9 25 0.13 45029 29577 65.7 369 1.2 29146 98.5 62 0.2 26 0.18 45029 44678 99.2 3972 8.9 39580 88.6 1125 2.5 27 0.12 45029 35442 78.7 987 2.8 34319 96.8 136 0.4 28 0.11 45029 35146 78.1 24746 70.4 9727 27.7 673 1.9 29 0.09 45029 9146 20.3 259 2.8 8743 95.5 153 1.7 30 0.17 17193 38.2 4897 28.5 11756 68.4 540 3.1 31 0.16 33503 74.4 71 0.2 33263 99.3 169 0.5 32 0.05 16613 36.9 367 2.2 16091 96.9 155 0.9 33 0.15 37587 83.5 735 2 36642 97.5 209 0.6 Average 28814.82 64.00303 2510.576 7.584848 25797.27 90.59394 507.5455 1.827273 Biofilm of the Age of 84 days (infected) REFERENCES Amini, S., and Tavazoie, S. 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