INTERACTION BETWEEN LEGIONELLA PNEUMOPHILA …

INTERACTION BETWEEN LEGIONELLA PNEUMOPHILA

AND BIOFILM FORMING ORGANISM

PSEUDOMONAS AERUGINOSA

WON CHOONG YUN

(B.Sc. (Hons.), NUS)

A THESIS SUBMITTED

FOR THE DEGREE OF MASTER OF SCIENCE

DEPARTMENT OF MICROBIOLOGY

NATIONAL UNIVERSITY OF SINGAPORE

2006

Acknowledgements

Department of Microbiology, NUS i

Acknowledgements

I would like to express my heartfelt gratitude to the following people who have

made a difference in my life during the course of this study:

A/Prof Lee Yuan Kun for his invaluable guidance, constant encouragement and

patience throughout the course of this study.

Dr Gamini Kumarasinghe from the Department of Laboratory Medicine, National

University Hospital, A/Prof Zhang Lian Hui from Institute of Molecular and Cell

Biology, and A/Prof Tim Tolker-Nielsen from BioCentrum-DTU, The Technical

University of Denmark, for kindly providing bacterial strains for this study.

Mr Ma Xi from Nalco Company for his invaluable advice, generous assistance

and constant concern. Dr Chen Hui and Mr Tim Lim, also from Nalco Company,

for their generous sharing of experiences and gracious assistance.

Mr Low Chin Seng for his precious technical assistance and for being a fatherly-

figure in a laboratory setting. Mdm Chew Lai Meng for her encouragement and

warm friendship.

Ho Phui San, Lee Hui Cheng, Wang Shugui and especially Chow Wai Ling and

Janice Yong Jing Ying for their generous help, precious friendship and incredible

understanding when absentmindedness get the better of me. Post-graduate life has

never been better without them!

Acknowledgements

Department of Microbiology, NUS ii

Toh Yi Er and Lee Kong Heng from Confocal Microscopy Unit, and Toh Kok Tee

from Flow Cytometry Unit for their invaluable technical assistance.

My family and husband, Clement Choo, for their generous love, unwavering

support and relentless encouragement through difficult time of my life. Especially

my father, for his thought-provoking discussions and tremendous help in software

improvements for this study. My son for sharing his precious life with me.

Table of Contents

Department of Microbiology, NUS iii

Table of Contents

Acknowledgements i

Table of Contents iii

List of Tables x

List of Figures xi

List of Abbreviations xv

Summary xvii

Chapter 1: Introduction 1

Chapter 2: Literature Review 5

2.1 Legionella 5

2.1.1 Introduction to Legionella 5

2.1.2 General characteristics of Legionella 5

2.1.3 Taxonomy of Legionella 7

2.1.4 Legionella and Diseases 8

2.1.4.1 Clinical presentation 8

2.1.4.2 Diagnosis 9

2.1.4.3 Epidemiology 10

2.1.4.4 Epidemiology in Singapore 13

2.1.4.5 Treatment 15

2.1.5 Ecology of Legionella 16

2.1.5.1 Natural and man-made habitats 16

2.1.5.2 Distribution of Legionella in Singapore 18

2.1.5.3 Association of Legionella with protozoa 19

2.1.5.4 Association of Legionella with biofilm 21

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Department of Microbiology, NUS iv

2.1.5.5 Interaction of Legionella with Pseudomonas spp. 24

2.2 Biofilm 24

2.2.1 Introduction to biofilm 24

2.2.2 General characteristics of biofilm 25

2.2.3 Biofilm development 26

2.2.4 Stages of biofilm development 27

2.2.4.1 Stage 1: Reversible attachment 27

2.2.4.2 Stage 2: Irreversible attachment 28

2.2.4.3 Stage 3: Maturation-1 29

2.2.4.4 Stage 4: Maturation-2 29

2.2.4.5 Stage 5: Dispersion 30

2.2.5 Determinants of biofilm structure 31

2.2.6 Microbial diversity of biofilms 33

2.2.7 Microbial positioning in biofilm 34

2.3 Prevention of legionellosis 35

2.3.1 Control of legionellosis 35

2.3.2 Detection of Legionella 36

2.3.3 Risk assessment of cooling tower for Legionnaires’ disease

outbreaks 37

2.3.4 Water treatment in cooling towers 38

Chapter 3: Materials and Methods 41

3.1 Bacterial strains and culture 41

3.1.1 Bacterial Strains 41

3.1.2 Culture Media 41

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Department of Microbiology, NUS v

3.1.3 Maintenance of stock cultures 42

3.2 Growth kinetic studies 42

3.2.1 Growth kinetics of L. pneumophila 42

3.2.2 Growth kinetics of P. aeruginosa PAO1 43

3.2.3 Growth kinetics of P. aeruginosa PAO1-CFP 43

3.3 Determination of the influent flow rate (Q) for continuous culture in

CDC Biofilm Reactor (CBR) 43

3.4 Optimization of labelling processes 44

3.4.1 Optimization of L. pneumophila labelling with CFDA-SE 44

3.4.2 Optimization of planktonic P. aeruginosa PAO1-CFP labelling

with PI 44

3.4.3 Flow cytometry 45

3.4.4 Optimization of P. aeruginosa PAO1-CFP biofilm labelling with PI 45

3.5 P. aeruginosa PAO1-CFP biofilm formation in CDC Biofilm Reactor

(CBR) 46

3.5.1 CDC Biofilm Reactor 46

3.5.2 Setup of CDC Biofilm Reactor assembly 47

3.5.3 P. aeruginosa PAO1-CFP biofilm formation 48

3.6 Introduction of L. pneumophila into P. aeruginosa PAO1-CFP biofilms 50

3.7 Introduction of NALCO 7320 into developing and mature

P. aeruginosa PAO1-CFP biofilms containing L. pneumophila 51

3.8 Monitoring of each organism in CBR continuous flow system 52

3.8.1 Preparation for sampling 52

3.8.2 Taking samples 52

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Department of Microbiology, NUS vi

3.8.2.1 Sampling bulk fluid 52

3.8.2.2 Sampling biofilm 53

3.8.3 Preparation of coupons 53

3.8.3.1 Preparation of coupons intended for enumeration 53

3.8.3.2 Preparation of coupons intended for visualization by CLSM 53

3.8.4 Disaggregation by homogenization 54

3.8.5 Enumeration of each organism 55

3.8.5.1 Enumeration of P. aeruginosa PAO1-CFP by culture 55

3.8.5.2 Enumeration of L. pneumophila by immunofluorescence 56

3.8.6 Detection of exogenous contaminants 58

3.8.7 Visualization and image acquisition by CLSM 59

3.8.8 Application of COMSTAT image analysis software package 60

3.8.8.1 Preparation of image stacks 60

3.8.8.2 Thresholding of images 61

3.8.8.3 COMSTAT image analysis for P. aeruginosa PAO1-CFP

biofilm structure 61

3.8.8.4 COMSTAT image analysis for porosity of P. aeruginosa

PAO1-CFP biofilm 63

3.8.8.5 COMSTAT image analysis for L. pneumophila distribution 64

3.8.9 Statistical analysis 65

3.9 Screening for effective P. aeruginosa PAO1 biofilm-removing agent 65

3.9.1 Kinetics of P. aeruginosa PAO1 biofilm formation in microtiter

plate 65

3.9.2 Quantification of biofilm 66

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Department of Microbiology, NUS vii

3.9.3 Biofilm-removing agents used 67

3.9.4 P. aeruginosa PAO1 biofilm removal screening 68

3.10 Antimicrobial susceptibility testing of NALCO 7320 69

Chapter 4: Results 70

4.1 Growth kinetics 70

4.2 Determination of the influent flow rate (Q) for continuous culture in CDC

Biofilm Reactor (CBR) 72

4.3 Optimization of labelling processes 74

4.3.1 Optimization of L. pneumophila labelling with CFDA-SE 74

4.3.2 Optimization of planktonic P. aeruginosa PAO1-CFP labelling

with PI 75

4.3.3 Optimization of P. aeruginosa PAO1-CFP biofilm labelling with PI 77

4.4 Kinetics of P. aeruginosa PAO1-CFP biofilm formation in CDC

Biofilm Reactor (CBR) 80

4.4.1 Kinetics of biofilm formation 80

4.4.2 Structure of biofilm by image analysis 81

4.4.3 Detachment of biofilm 85

4.5 Introduction of L. pneumophila to developing and mature P. aeruginosa

PAO1-CFP biofilms 87

4.5.1 Adhesion and persistence of L. pneumophila in developing and

mature biofilms 87

4.5.2 Distributions of L. pneumophila cells in developing and mature

biofilms 90

4.5.3 Bio-volume distributions of developing and mature biofilms 95

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Department of Microbiology, NUS viii

4.5.4 Surface-to-biovolume ratio distributions of developing and

mature biofilms 97

4.5.5 Porosity distributions of developing and mature biofilms 100

4.5.6 Correlation between SBR and porosity 103

4.5.7 Correlation between legionellae adhesion and parameters of

P. aeruginosa PAO1-CFP biofilm 104

4.5.8 Localization of L. pneumophila in P. aeruginosa PAO1-CFP

biofilms 105

4.6 Screening for effective P. aeruginosa PAO1 biofilm removing agent 108

4.6.1 Kinetics of P. aeruginosa PAO1 biofilm formation in microtiter

plate 108

4.6.2 P. aeruginosa PAO1 biofilm removal screening 109

4.7 Characterization of NALCO 7320 111

4.7.1 Kinetics of P. aeruginosa PAO1 biofilm removal 111

4.7.2 Antimicrobial susceptibility testing 112

4.8 Introduction of NALCO 7320 into developing and mature P. aeruginosa

PAO1-CFP biofilms containing L. pneumophila 114

4.8.1 Persistence of P. aeruginosa PAO1-CFP in CBR 114

4.8.2 Structure of P. aeruginosa PAO1-CFP biofilms treated by NALCO

7320 115

4.8.3 Persistence of L. pneumophila in P. aeruginosa PAO1-CFP biofilms

treated with NALCO 7320 120

4.8.4 Distribution of L. pneumophila in P. aeruginosa PAO1-CFP biofilms

treated with NALCO 7320 123

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Department of Microbiology, NUS ix

4.8.5 Bio-volume distributions of developing and mature biofilms treated

with NALCO 7320 125

4.8.6 Porosity distributions of P. aeruginosa PAO1-CFP biofilms treated

with NALCO 7320 127

Chapter 5: Discussion 130

References 147

Appendix 175

List of Tables

Department of Microbiology, NUS x

List of Tables

Table 3.1. Table 3.2. Table 4.1. Table 4.2. Table 4.3. Table 4.4. Table 4.5. Table 4.6. Table 4.7. Table 4.8. Table 4.9.

Sampling points of 6 independent experiments for the study of P. aeruginosa PAO1-CFP biofilm formation. List of biofilm-removing agents used. Effect of treatment duration on staining and viability of L. pneumophila cells. Effect of treatment duration on staining of P. aeruginosa PAO1-CFP cells. Table showing Pearson’s correlation between Log (Number of L. pneumophila cells) and Log (Number of CFDA pixels per µm3). The ratio of SBR at the bottom 20% versus the top 20% of developing and mature biofilm. Comparing means of porosity over time. Table showing Pearson’s correlation between porosity and SBR. Table showing Pearson’s correlation between legionellae adhesion to P. aeruginosa PAO1-CFP biofilm (representing the number of legionellae per coupon per 106 legionellae inoculated into CBR) and parameters of the biofilm. Efficacy of biofilm removing agents. Table showing Pearson’s correlation between bio-volume and legionellae loss.

49 68 74 76 92 100 101 103 104 110 122

List of Figures

Department of Microbiology, NUS xi

List of Figures Figure 3.1. Figure 4.1. Figure 4.2. Figure 4.3. Figure 4.4. Figure 4.5. Figure 4.6. Figure 4.7. Figure 4.8. Figure 4.9.

Schematic diagram of the CDC Biofilm Reactor assembly. Growth curve of L. pneumophila cultured in BCYE broth at 37°C with shaking at 120rpm. Growth curve of P. aeruginosa PAO1 cultured in MM liquid media at 30°C with shaking at 120rpm. Growth curve of P. aeruginosa PAO1-CFP cultured in MM liquid media at 30°C with shaking at 120rpm. Graph of Ln(OD600nm) against time (hr) plotted for the exponential growth phase of P. aeruginosa PAO1-CFP. Histograms illustrating the number of events (cells) plotted against FL1-H (representing green fluorescence of CFDA-stained cells) for L. pneumophila cells that were (A) mock treated, or treated with CFDA-SE for (B) 20mins, (C) 30mins, or (D) 40mins. Histograms illustrating the number of events (cells) plotted against PMT4 Log (representing red fluorescence of PI-stained cells) for P. aeruginosa PAO1-CFP cells that were (A) mock treated, or treated with 1.0mg/ml PI for (B) 5mins, (C) 10mins, or (D) 15mins. Histograms illustrating the number of events (cells) plotted against PMT4 Log (representing red fluorescence of PI-stained cells) for P. aeruginosa PAO1-CFP cells that were (A) mock treated, or treated with 0.1mg/ml PI for (B) 5mins, (C) 10mins, (D) 15mins, or (E) 30mins. CLSM images of a 7 days old P. aeruginosa PAO1-CFP biofilm and adhered L. pneumophila, stained with 0.1mg/ml PI for 5mins: (A) P. aeruginosa PAO1-CFP biofilm (blue fluorescence), (B) CFDA-stained L. pneumophila (green fluorescence), (C) PI-stained P. aeruginosa PAO1-CFP biofilm, and (D) overlapping display of the above 3 images. CLSM images of a 7 days old P. aeruginosa PAO1-CFP biofilm and adhered L. pneumophila, stained with 0.1mg/ml PI for 15mins: (A) P. aeruginosa PAO1-CFP biofilm (blue fluorescence), (B) CFDA-stained L. pneumophila (green fluorescence), (C) PI-stained P. aeruginosa PAO1-CFP biofilm, and (D) overlapping

47 70 71 71 72 74 75 76 77 78

List of Figures

Department of Microbiology, NUS xii

Figure 4.10. Figure 4.11. Figure 4.12. Figure 4.13. Figure 4.14. Figure 4.15. Figure 4.16. Figure 4.17. Figure 4.18. Figure 4.19. Figure 4.20. Figure 4.21. Figure 4.22.

display of the above 3 images. CLSM images of a 7 days old P. aeruginosa PAO1-CFP biofilm and adhered L. pneumophila, stained with 0.1mg/ml PI for 30mins: (A) P. aeruginosa PAO1-CFP biofilm (blue fluorescence), (B) CFDA-stained L. pneumophila (green fluorescence), (C) PI-stained P. aeruginosa PAO1-CFP biofilm, and (D) overlapping display of the above 3 images. Viable cell counts of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. Bio-volume of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. Average thickness of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. Maximum thickness of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. Substratum coverage of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. Surface-to-biovolume ratio (SBR) of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. Roughness coefficient of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. Viable cell counts of planktonic P. aeruginosa PAO1-CFP in the bulk fluid of CBR at 30°C with stirring at 120rpm. CLSM image of a P. aeruginosa PAO1-CFP biofilm (blue) structure indicative of dispersion stage of biofilm development, with adhered L. pneumophila (green). Adhesion of L. pneumophila to different developmental stages of P. aeruginosa PAO1-CFP biofilm. Status of L. pneumophila in our continuous flow CBR system. Persistence of L. pneumophila in P. aeruginosa PAO1-CFP biofilm.

79 80 82 82 83 83 84 84 85 86 87 89 89

List of Figures

Department of Microbiology, NUS xiii

Figure 4.23. Figure 4.24. Figure 4.25. Figure 4.26. Figure 4.27. Figure 4.28. Figure 4.29. Figure 4.30. Figure 4.31. Figure 4.32. Figure 4.33. Figure 4.34. Figure 4.35. Figure 4.36. Figure 4.37.

Distribution of L. pneumophila in (A) developing, and (B) mature P. aeruginosa PAO1-CFP biofilms. Percentage loss of L. pneumophila in developing P. aeruginosa PAO1-CFP biofilm. Percentage loss of L. pneumophila in mature P. aeruginosa PAO1-CFP biofilm. Bio-volume distribution of (A) developing, and (B) mature P. aeruginosa PAO1-CFP biofilms. Surface-to-biovolume ratio (SBR) distribution of (A) developing, and (B) mature P. aeruginosa PAO1-CFP biofilms. Porosity of P. aeruginosa PAO1-CFP biofilm. Porosity distribution of (A) developing, and (B) mature P. aeruginosa PAO1-CFP biofilms. Scatterplot of porosity and SBR both obtained from all data of 6 independent experiments. CLSM images of P. aeruginosa PAO1-CFP biofilm (blue) with adhered L. pneumophila (green) taken on different occasions: (A) 3hrs after legionellae introduction to developing biofilm (3-days-old), (B) 4 days after legionellae introduction to developing biofilm, (C) 3hrs after legionellae introduction to mature biofilm (7-days-old), and (D) 4 days after legionellae introduction to mature biofilm. Kinetics of P. aeruginosa PAO1 biofilm formation in microtitre plate at 30°C. Highest percentage biofilm removal of various biofilm-removing agents. Kinetics of biofilm removal by NALCO 7320. Visual determination of minimum inhibitory concentration (MIC). Determination of minimum bactericidal concentration (MBC) of NALCO 7320. Viable cell counts of P. aeruginosa PAO1-CFP biofilms

93 94 94 96 99 101 102 103 107 108 109 111 113 113 114

List of Figures

Department of Microbiology, NUS xiv

Figure 4.38. Figure 4.39. Figure 4.40. Figure 4.41. Figure 4.42. Figure 4.43. Figure 4.44. Figure 4.45. Figure 4.46. Figure 4.47. Figure 4.48. Figure 4.49. Figure 4.50. Figure 4.51.

treated with NALCO 7320. Viable cell counts of planktonic P. aeruginosa PAO1-CFP in CBR treated with NALCO 7320. Bio-volume of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. Average thickness of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. Maximum thickness of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. Substratum coverage of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. Surface-to-biovolume ratio of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. Roughness coefficient of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. Persistence of L. pneumophila in P. aeruginosa PAO1-CFP biofilms treated with NALCO 7320. Cell counts of planktonic L. pneumophila in CBR treated with NALCO 7320. Scatterplot of bio-volume and legionellae loss, obtained from 4 independent experiments. Effect of NALCO 7320 on the distribution of L. pneumophila in (A) developing, and (B) mature P. aeruginosa PAO1-CFP biofilms. Effect of NALCO 7320 on the distribution of bio-volume in (A) developing, and (B) mature P. aeruginosa PAO1-CFP biofilms. Porosity of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. Effect of NALCO 7320 on porosity distribution of (A) developing, and (B) mature P. aeruginosa PAO1-CFP biofilms.

115 117 117 118 118 119 119 121 121 122 124 126 128 129

List of Abbreviations

Department of Microbiology, NUS xv


3OC12-HSL

µmax

θ

BCYE

CBR

CFDA-SE

CFU

CLSM

DBNPA

DFA

dH2O

EPS

H. vermiformis

LB

LB+gen

LD

LLAP

L. pneumophila

MBC

MIC

MM

MM+gen

N-(3-oxododecanoyl)-L-homoserine lactone

Maximum specific growth rate

Hydraulic residence time

Buffered charcoal yeast extract

CDC biofilm reactor

5-(and-6)-carboxyfluorescein diacetate, succinimidyl ester Colony forming unit

Confocal laser scanning microscope

2,2-Dibromo-3-nitrilopropionamide

Direct florescent antibody

Deionized water

Extracellular polysaccharides

Hartmannella vermiformis

Luria Bertani

Luria Bertani + 60µg/ml gentamicin

Legionnaires’ disease

Legionella-like amoebal pathogens

Legionella pneumophila

Minimum bactericidal concentration

Minimum inhibitory concentration

Minimal media

Minimal media + 60µg/ml gentamicin


Department of Microbiology, NUS xvi

OD

P. aeruginosa

Optical density

Pseudomonas aeruginosa

P. aeruginosa PAO1-CFP

PBS

PCR

PF

PFA

PI

ppm

Q

r

rpm

SBR

SDS

td

V

VBNC

Pseudomonas aeruginosa PAO1 tagged with cyan-fluorescent-protein (CFP) Phosphate buffer solution

Polymerase chain reaction

Pontiac fever

Paraformaldehyde

Propidium iodide

Parts per million

Nutrient influent flow rate

Correlation coefficient

Revolutions per minute

Surface to bio-volume ratio

Sodium dodecyl sulphate

Doubling time

Maximum volume of bulk fluid in CBR during continuous flow Viable but non-culturable

Summary

Department of Microbiology, NUS xvii

Summary

In present study, a reproducible model Pseudomonas aeruginosa PAO1-CFP

biofilm, with distinct stages of biofilm development, was established in CDC

Biofilm Reactor continuous flow system using defined minimal media at 30°C.

Splitting certain data, such as bio-volume and surface-to-biovolume ratio (SBR),

into 5 sections along biofilm thickness and applying a novel method of biofilm

porosity quantification in a 3-dimensional context provided greater insights of

biofilm structures and properties. Consequently, biofilm structures and

development were better described, and the first physical evidence of porous

channels within biofilm cell cluster was observed.

Legionella pneumophila adhesion study revealed that legionellae adhesion to

biofilms was independent of developmental stage of the latter. Instead, biofilm

structure and porosity were found to determine the amount and even localization

of legionellae adhesion to biofilm. Additionally, L. pneumophila persistence study

revealed that legionellae was least likely to get desorbed at bottom 60% of the

biofilms, especially at bottom 20%, and unbalanced advective transport of

legionellae towards biofilm surface commenced upon biofilm maturation, most

probably due to unbalanced cell growth.

Eight commercially available biofilm-removing agents were screened using

microtitre plate assay for one with the highest efficacy. Subsequently, application

of the selected biocide, NALCO 7320, (at bactericidal concentration to planktonic

P. aeruginosa PAO1) to P. aeruginosa PAO1-CFP biofilms yielded complete

Summary

Department of Microbiology, NUS xviii

disinfection of developing biofilm while a resistant subpopulation was found in

the remains of mature biofilm.

Porosity distribution and biofilm structural analysis suggested that NALCO 7320

caused biofilm detachment by affecting the nature of extracellular polysaccharides

(EPS) matrix that bound the microbial cells together as a microcolony, while

applying biocidal effect on P. aeruginosa PAO1-CFP cells within the biofilm.

Legionellae persistence in biocide-treated biofilms was found to be independent

on the stage of biofilm development and loss of biomass, but regions of the

biofilms in which legionellae best persist were detected. Since EPS is a major

component in biofilm matrix, it was hypothesized to play an important role in

legionellae persistence in biocide-treated biofilms.

Introduction

Department of Microbiology, NUS 1

Chapter 1: Introduction

L. pneumophila, the main species of the genus Legionella, was first recognized as

a pathogen after an outbreak of acute pneumonia, called Legionnaires’ disease that

occurred at the convention of the American Legion in Philadelphia, USA, during

1976 (Fraser et al., 1977). To date, forty eight species of Legionella have been

described, including 70 distinct serogroups (Borella et al., 2005). Approximately

half of the 48 species of legionellae have been associated with legionellosis, but it

is likely that most legionellae can cause human disease under appropriate

conditions (Fields, 1996). L. pneumophila is responsible of approximately 91% of

all reported community cases of legionellosis and among the 15 serogroups of this

species, L. pneumophila serogroup 1 accounts for the 84% of confirmed cases (Yu

et al., 2002).

The real number of cases of Legionnaires’ disease is unknown, although in the

USA, it is estimated that the incidence is 20 cases per million population (Borella

et al., 2005). In Europe, during the period 2003-2004, a total of 10,322 cases of

Legionnaires’ disease was reported, with national infection rates ranging from 0 to

28.7 cases per million population (Ricketts and Joseph, 2005). The mean annual

incidence rates were 0.9 (Heng et al., 1997) and 1.7 (Goh et al., 2005) per 100,000

population in Singapore, during the period 1986-1996 and 1998-2002

respectively. Because of the difficulty in distinguishing Legionella associated

diseases from other forms of pneumonia and influenza, many cases are

unreported. Nevertheless, the overall case-fatality rate is high especially among

Introduction


seriously immunosuppressed individuals, at 24% for the adequately treated and

80% for those without treatment (Fliermans, 1995).

Legionella associated diseases have emerged in the last half of the 20th century

because of human alteration of the environment. Legionella spp. is part of the

natural aquatic environment and the bacterium is capable of surviving extreme

ranges of environmental conditions (Fliermans et al., 1981). However, when

allowed to remain in their natural habitat, legionellae are rarely the causative

agents of human disease since natural freshwater environments have not been

implicated as reservoirs of legionellosis. Main sources of L. pneumophila are

waters from hot distribution systems and cooling towers. Numerous cases of

legionellosis have been found to occur after exposure to contaminated waters in

offices, hotels, hospitals and cruise ships, among other locations (Borella et al.,

2005).

Factors leading to outbreaks or sporadic cases are not completely understood, but

certain events are considered prerequisites for infection. These include the

presence of the bacterium in aquatic environment, amplification to an unknown

infectious dose and transmission via aerosol to a human host that is susceptible to

infection (Fliermans, 1995). Although amoebae are key factors in Legionella

amplification process (Fields, 1996), this pathogen is able to survive as free

organism for long periods within biofilms which are widespread in man-made

water systems. Its persistence has been attributed to survival within biofilms

(Rogers and Keevil, 1992; Rogers et al., 1994). Additionally, association of

Introduction


Legionella to biofilm may explain, at least in part, why legionellae are relatively

hard to eradicate in water systems, as biofilms exhibit a marked resistance to

biocidal compounds and chlorination (LeChevallier et al., 1988). Therefore a

more extensive knowledge on biofilm-associated legionellae may lead to the most

effective control measures to prevent legionellosis.

Majority of Legionella-biofilm studies employed naturally occurring microbial

biofilm communities, and failed to identify all the organisms present and their

contribution to the survival and multiplication of legionellae. Additionally,

Pseudomonas aeruginosa PAO1, a wound isolate (Holloway, 1955), is generally

found in the same aquatic environments as L. pneumophila (Murga et al., 2001), is

the most widely used P. aeruginosa laboratory strain (Stover et al., 2000) and its

biofilm development has been well documented (Sauer et al., 2002). Therefore in

the present study, a reproducible model P. aeruginosa PAO1-CFP biofilm was

established in a CDC Biofilm Reactor continuous flow system using defined

minimal media at 30°C. Since P. aeruginosa PAO1 biofilms are structurally and

dynamically complex biological systems with regulated developmental stages

(Sauer et al., 2002), it was hypothesized that legionellae interacts differently with

biofilms at different developmental stages and responds differently to biocidal

treatments while residing in biofilms at different developmental stages.

To allow further insights into biofilm development, current method of quantifying

biofilm structures was improved by splitting up certain descriptive data into 5

sections along the thickness of the biofilm and a novel method of quantifying

Introduction


biofilm porosity in a 3-dimensional context was developed. Using the model and

better descriptive methods of biofilm structure and porosity, it was determined if

there is any difference (in numbers and distribution pattern) in accumulation and

persistence of L. pneumophila in developing and mature biofilm, and if the

structure or porosity of biofilm plays a role in the accumulation and persistence of

L. pneumophila.

In a bid to deepen the knowledge on the effect of biocide on legionellae-

associated to biofilms, a biocide was first selected by screening through eight

commercially available biofilm-removing agents for one with the highest efficacy

using microtitre plate assay. Subsequently, the effects of the selected biocide,

NALCO 7320 (at bactericidal concentration to planktonic P. aeruginosa PAO1)

on the persistence and structure of P. aeruginosa PAO1-CFP biofilm, and the

persistence of biofilm-associated legionellae were characterized.

Literature Review


Chapter 2: Literature Review

2.1 Legionella

2.1.1 Introduction to Legionella

The terror of the unknown is seldom better displayed than by the response of a

population to the appearance of an epidemic, particularly when the epidemic

strikes without apparent cause. Between July 22 and August 3, 1976, there was a

remarkable incidence of febrile respiratory disease among persons who had

attended the American Legion Convention in Philadelphia from July 21 to 24.

“Legionnaires’ disease” (LD) is the term used to describe the illness that occurred

among persons attending the convention (Fraser et al., 1977).

The etiologic agent of LD was first isolated in guinea pigs from lung specimens

collected on autopsy and subsequently, serologic evidence for the etiological role

of the bacterium, designated L. pneumophila subsp. pneumophila, was obtained by

indirect fluorescent antibody staining (McDade et al., 1977). In fact, the first

strains of Legionella were already isolated in guinea pigs by using procedures for

the isolation of Rickettsia by Tatlock in 1943 (McDade et al., 1979).

2.1.2 General characteristics of Legionella

Members of the genus Legionella are faintly staining Gram-negative, aerobic rods

or filaments (usually found after growth in enriched laboratory media), 0.3-0.9µm

in width and 2-20µm or more in length (Brenner et al., 1985). They are neither

encapsulated nor acid-fast; they do not form endospores or microcysts (Brenner et

al., 1985). They are chemoorganotrophic, where amino acids are utilized as

Literature Review


carbon and energy sources, while carbohydrates are neither fermented nor

oxidized (Tesh and Miller, 1981). Furthermore, they are nutritiously fastidious

where L-cysteine-HCL is absolutely necessary for their growth and iron salts in

the medium enhance their growth (Feeley et al., 1978).

The cell wall is made up of two three-layered unit membranes (Brenner et al.,

1985) and is predominated by branched chain fatty acids (Fisher-Hoch et al.,

1979). The fatty acid composition of the cell wall varies among the different

species belonging to the genus Legionella, thus fatty acid analysis is useful for the

differentiation of Legionella species (Diogo et al., 1999). In addition, the cellular

fatty acid composition of the bacteria is found to be similar to that of known

thermophilic bacteria (Fliermans, 1995). Therefore, it is not surprising to see

Legionella associated with thermally elevated habitats (Verissimo et al., 1991).

Various L. pneumophila strains and isolates of species other than L. pneumophila

are able to produce flagella (Heuner et al., 1995), which are later shown to be a

positive predictor for virulence in Legionella (Bosshardt et al., 1997). Ott et al.

(1991) demonstrated that the expression of the gene flaA, encoding the flagella

subunit, is temperature-dependent. Further studies in the same laboratory revealed

that the expression of flaA is also influenced by the growth phase, the viscosity

and the osmolarity of the medium, and by amino acids (Heuner et al., 1999).

Similar to a number of other Gram-negative bacteria, Legionella is able to enter a

viable but non-culturable (VBNC) state under low-nutrient conditions (Hussong et

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al., 1987). Also, the loss of culturability appeared to be accelerated at higher

temperature of 37°C, as compared to 4°C (Hussong et al., 1987). Many

procedures to reactivate VBNC legionellae have failed, with the exception of the

passage through Acanthamoeba castellani (Steinert et al., 1997). Both amoeba-

reactivated cells and plate-grown L. pneumophila cells had the same capacity for

intracellular survival in human monocytes and intraperitoneally infected guinea

pigs, which is considered a parameter for virulence. However, reactivation of

VBNC cells was not observed in the animal model. Although there is a correlation

of Legionella infection of amoeba, human cell lines and animal models, it cannot

be excluded that VBNC forms are virulent for human.

2.1.3 Taxonomy of Legionella

The family Legionellaceae consists of the single genus Legionella (Fields et al.,

2002). At least 48 species comprising 70 serogroups have been distinguished

(Fields et al., 2002; Borella et al., 2005). Legionella pneumophila consists of 15

serogroups, of which serogroup 1 is the most common, followed by serogroups 4

and 6 (Den Boer and Yzerman, 2004). The number of species and serogroups of

legionellae continues to increase. Phylogenetically, the nearest relative to the

Legionellaceae is Coxiella burnetti, the etiologic agent of Q fever (Adeleke et al.,

1996 and Swanson and Hammer, 2000). These organisms have similar

intracellular lifestyles and may utilize common genes to infect their host.

Some legionellae cannot be grown on routine Legionella media and has been

termed Legionella-like amoebal pathogens (LLAPs; Adeleke et al., 1996). LLAP

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was first recovered and isolated from the sputum of a patient with pneumonia by

cocultivating with its host amoebae (Fry et al., 1991). Additional LLAP strains

may be human pathogens as well, but proving this is difficult because they cannot

be detected by conventional techniques used for legionellae (Fields et al., 2002).

2.1.4 Legionella and Diseases

2.1.4.1 Clinical presentation

Diseases caused by Legionella are collectively termed legionellosis. Legionellosis

classically presents as two distinct clinical entities, Legionnaires’ disease (LD;

Fraser et al., 1977), a severe multisystem disease involving pneumonia, and

Pontiac fever (PF; Glick et al., 1978), a self-limited flu-like illness.

Features of LD include fever, non-productive cough, headache, myalgias, rigors,

dyspnea, diarrohea and delirium (Tsai et al., 1979). Histological reports describe

intra- and extracellular bacteria in phagocytes, fibroblasts and epithelial cells

(Fields, 1996). Chest X-rays often show evidence of pneumonia, but it is

impossible to distinguish LD from other types of pneumonia on the basis of

symptoms alone (Edelstein, 1993). As a result, many cases go probably

unreported. This assumption is supported by serologic surveys which show that

many persons in an apparently healthy population have antibodies against

legionellae (Paszko-Kolva et al., 1993).

The clinically distinct self-limited and non-pneumonic PF is a milder, influenza-

like form of disease (Fields et al., 1990). It usually appears on an epidemic mode

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(Tossa et al., 2006) but many persons who seroconvert to Legionella will be

entirely asymptomatic (Boshuizen et al., 2001). Because of its benignity and lack

of specificity, the occurrence of PF is often undiagnosed and is therefore even less

reported than LD.

To date, there is no consensus on the duration of the incubation period, on its

clinical symptoms, nor on the causal species of Legionella. Since the microbe has

never been isolated from PF patients, it has been speculated that PF is caused by

VBNC forms of Legionella (Steinert et al., 1997). Other hypotheses to explain PF

include changes in virulence factors, toxic or hypersensitivity reactions to bacteria

(Kaufmann et al., 1981) or their products; high levels of endotoxin in aerosolized

water may be responsible for clinical symptoms (Fields et al., 2001).

2.1.4.2 Diagnosis

Although Legionella species are gram-negative bacilli, they are rarely visualized

on Gram stains of clinical material (Stout et al., 2003). A Gram stain of a sputum

specimen showing polymorphonuclear leukocytes without bacteria can be a

valuable clue to Legionella infection (Muder and Yu, 2002).

Clincal specimens used for culture of Legionella species include sputum or

bronchoalveolar lavage specimens, bronchial aspirates, lung biopsy specimens and

blood (Den Boer and Yzerman, 2004). Isolation of Legionella species from a

clinical specimen on selective media provides a definitive diagnosis. Buffered

charcoal yeast extract agar that contains antibiotics to suppress commensal flora is

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commercially available. However, certain media formulations that are selective

for L. pneumophila may inhibit growth of other Legionella species (Muder and

Yu, 2002). Preheating steps and acid washing procedures were developed to

reduce overgrowth by other microorganisms, thus serve as additional means of

increasing the sensitivity of sputum culture (Den Boer and Yzerman, 2004).

Species-specific DFA testing is more often applied directly to clinical specimens

(Stout et al., 2003). Since the sensitivity of the DFA stain is much lower than for

culture (range, 25 to 75%), it was suggested that this test should not be performed

routinely (Edelstein, 1993). It should be noted that the sensitivity and specificity

of DFA testing of clinical specimens is not precisely known for species other than

L. pneumophila (Muder and Yu, 2002).

The commercially available Legionella urinary antigen test reliably detects only

infection due to L. pneumophila serogroup 1. Urinary antigen test results are

occasionally positive in cases of disease due to other Legionella species, but the

sensitivity is low; consequently, a negative test result is of little value in excluding

Legionella infection (Muder and Yu, 2002). Potentially, PCR could detect all

known Legionella species. However, so far the sensitivity of the test varies from

11 to 100% and many publications report specificities of lower than 99% (Den

Boer and Yzerman, 2004).

At present, optimal sensitivity for diagnosis of LD will be achieved by using a

combination of culture, serological investigation and urinary antigen detection

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(Den Boer and Yzerman, 2004). After reviewing the diagnostic methods of LD,

Den Boer and Yzerman (2004) postulated that easy-to-perform PCR test with high

sensitivity and a specificity of above 99% may become accepted as new gold

standard for diagnosis of LD in the future. On the contrary, the sensitivity and

specificity of detecting seroconversion to Legionella species other than L.

pneumophila is still uncertain. While seroconversion alone can be used for the

diagnosis of infection due to other species, such diagnoses should be regarded as

presumptive unless there are supporting microbiologic or epidemiologic data

(Muder and Yu, 2002).

2.1.4.3 Epidemiology

Studies have estimated that between 8,000 and 18,000 persons are hospitalized

with legionellosis annually in the United States (Marston et al., 1997). Failure to

utilize available diagnostic tools may result in the mistaken impression that

Legionella infections are not occurring in a hospital or a community. For

Legionella infections in particular, national extrapolations are potentially

misleading because of the critical importance of local microenvironment.

As summarized by Fliermans (1995), the overall case-fatality rate is high. Among

previously healthy individuals, 7-9% die when treated with erythromycin, while

25% die when hospitalized but not treated with appropriate antibiotics. Among

seriously immunosuppressed individuals, the mortality rate is 24% for the

adequately treated and 80% for those without treatment.

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In the same study carried out by Marston et al. (1997) involving patients with

community-acquired pneumonia requiring hospitalization, Legionella spp. was

found to be responsible for 2–5% of the cases studied. In the USA, 91% of

isolates from Legionnaires’ disease patients are typed as Legionella pneumophila

serogroup 1 (Marston et al., 1997). This is in contrast to the situation in Australia

and New Zealand, where 30% of the cases of Legionnaires’ disease are caused by

Legionella longbeachae (Yu et al., 2002).

In an international collaborative study conducted by Yu et al. (2002), community

acquired LD is dominated by L. pneumophila serogroup 1 (84.2% of all isolates).

Species other than L. pneumophila were rare: L. longbeachae (3.9%) and L.

bozemanii (2.4%) accounted for most of the nonpneumophila cases. L. micdadei,

L. feeleii, L. dumoffii, L. wadsworthii and L. anisa combined accounted for 2.2%

of the remaining cases. Hospital-acquired pneumonia have also involved

serogroups other than L. pneumophila serogroup 1 (especially serogroups 4 and 6)

and Legionella species other than L. pneumophila, especially L. micdadei, L.

dumoffii and L. bozemanii (Fang et al., 1989). Nevertheless, L. pneumophila

serogroup 1 is still the dominant cause of legionellosis.

Epidemiological studies indicate that Legionella is an opportunistic pathogen,

with elderly and immuno-compromised patients being most susceptible

(Fliermans, 1996). Other risk factors for the disease include smoking, male sex,

chronic lung disease, hematologic malignancies, end-stage renal disease, lung

cancer, immunosuppresion and diabetes (Marston et al., 1994). Differences in host

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susceptibility and bacterial virulence make it difficult to clearly define an

infectious dose (Steinert et al., 2002).

The best-documented route for transmission of infection is the generation of an

infective aerosol from a Legionella contaminated water source (Winn, 1999).

Aspiration of contaminated potable water is another probable mechanism for

infection of the lower respiratory tract (Blatt et al., 1993). Entry through the

gastrointestinal tract has been suggested to explain abdominal infections, although

this portal of entry has not been proved. Direct entry of bacteria into flesh wound

may also cause nosocomial Legionella infection (Winn, 1999). There has been no

evidence of human-to-human transmission or documented laboratory infections.

2.1.4.4 Epidemiology in Singapore

To find out if the disease occurs in Singapore, legionellosis was made

administratively notifiable in 1985 and legally notifiable in 2000, to the

Quarantine and Epidemiology Department, Ministry of the Environment. The first

local case of LD, a 27 year old Chinese male plumber, was admitted to Toa Payoh

Hospital on 4th February, 1986 (Lim et al., 1986). Clinical suspicion of LD was

confirmed by the presence of serum antibody to L. pneumophila (titre 1:512) by

an indirect fluorescent antibody test and it is not known where the patient acquired

the illness from.

In an attempt to determine the level of antibodies to L. pneumophila serogroups 1

to 4 in 150 young normal adults who are blood bank donors, Nadarajah et al.

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(1987) discovered a 20% overall prevalence of antibodies to L. pneumophila in

the normal population studied. In addition, a national serologic survey of the

general population conducted in 1993 showed a prevalence of 10.3% in those

below 20 years of age and 21.9% in those 20 years of age and above (Heng et al.,

1997). These results suggest that L. pneumophila is wide spread in the

environment in Singapore, which is confirmed by a surveillance of Legionella

bacteria in various artificial water systems (Heng et al., 1997). Despite the

ubiquitous distribution of Legionella in artificial water systems in Singapore and

high prevalence of sero-converted individuals, there has been no clustering of

cases by person and place and no common source outbreak linked to any artificial

water system since the disease was made notifiable in 1985. Goh et al. (2005)

suggested that the absence of outbreak was due to the low prevalence of the highly

pathogenic Pontiac subtype of L. pneumophila locally or low Legionella counts in

the cooling towers and other water systems here (only one fifth with Legionella

colony count above 10 colony-forming units (CFU) /ml).

During the period 1986 to 1996, a total of 258 sporadic cases of community-

acquired legionellosis was reported, giving a mean annual morbidity rate of 0.9

per 100,000 population (Heng et al., 1997). However, a total of 273 cases,

including 37 imported cases, were reported during the period 1998-2002, giving a

mean annual incidence rate of 1.7 per 100,000 population (Goh et al., 2005).

These are lower than that of the USA (20 per 100,000 population per year; Borella

et al., 2005) and Scotland (5.1 per 100,000 population), and comparable to that of

Denmark (1.8 per 100,000 population), Germany (1.6 per 100,000 population) and

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England and Wales (0.2 per 100,000 population; Heng et al., 1997). In both

studies conducted in Singapore, cases were reported predominantly among males,

ethnic Indians, the elderly and those with concurrent medical conditions. The

overall case-fatality rate was 14.7% for the period 1986-1996 (Heng et al., 1997)

and 5.5% for 1998-2002 (Goh et al., 2005).

Legionella pneumonia accounted for 2% to 7% of the community-acquired

pneumonia among hospitalized patients in Singapore (Ong and Eng, 1995) The

incidence of community-acquired pneumonia due to legionellosis in USA is 2-5%

(Marston et al., 1997), in Italy is 5.9% (Montagna et al., 2006) and in Brazil is

5.1% (Chedid et al., 2005). Thus in most countries, it is less than 10%.

2.1.4.5 Treatment

In order to administer accurate treatment to patients with Legionnaires’ disease,

correct diagnosis is critical. Unfortunately, it is not possible to clinically

distinguish patients with Legionnaires’ disease from patients with other types of

pneumonia (Edelstein, 1993). Furthermore, delay in starting appropriate therapy

has been associated with increased mortality (Heath et al., 1996). Thus, Bartlett et

al. (2000) proposed that empirical therapy for persons hospitalized with

community-acquired pneumonia should include coverage for Legionnaires’

disease.

Historically, erythromycin has been the drug of choice for Legionnaires’ disease

(Fields et al., 2002). In vitro data suggest that azithromycin and many

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fluoroquinolone agents have superior activity against Legionella spp.

Additionally, these agents have fewer side effects than erythromycin (Edelstein,

1998). Since azithromycin and levofloxacin has been licensed by the Food and

Drug Admininstration for the treatment of Legionnaires’ disease, thus they are

preferred over erythromycin (Fields et al., 2002).

2.1.5 Ecology of Legionella

2.1.5.1 Natural and man-made habitats

Water is the major reservoir for legionellae and the bacteria are found in

freshwater environments worldwide (Fliermans et al., 1981). Legionellae have

been detected in as many as 40% of freshwater environments by culture and in up

to 80% of freshwater sites by PCR (Fields, 2002). Furthermore, Legionella has

been shown to survive in marine waters (Heller et al., 1998) and even ocean

waters receiving treated sewage have been found to contain Legionella species

(Palmer et al., 1993). In contrast with the aquatic environment, L. longbeachae is

a frequent isolate from potting soil (Steele et al., 1990). This species is the leading

cause of legionellosis in Australia and occurs in gardeners and those exposed to

commercial potting soil (Ruehlemann and Crawford, 1996).

L. pneumophila multiplies at temperatures between 25ºC and 42ºC, with an

optimal growth temperature of 35ºC (Katz and Hammel, 1987). However, in

nature or in association with algae, the optimum growth temperature of Legionella

spp. may be 45°C or higher (Fliermans et al., 1981). Most cases of legionellosis

can be traced to human-made aquatic environments where the water temperature

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is higher than ambient temperature (Fields et al., 2002). Thermally altered aquatic

environments can shift the balance between protozoa and bacteria, resulting in

rapid multiplication of legionellae, which can translate into human disease.

Legionellosis is a disease that has emerged in the last half of the 20th century

because of human alteration of the environment. Left in their natural state,

legionellae would be an extremely rare cause of human disease, as natural

freshwater environments have not been implicated as reservoirs of outbreaks of

legionellosis. Furthermore, the population densities of Legionella spp. in

freshwater are extremely low and at the highest densities measured Legionella

spp. account for less than 1% of the total bacterial population (Fliermans et al.,

1981).

Human infection occurs exclusively by inhalation of contaminated aerosols which

can be produced by air conditioning systems, cooling towers, whirlpools, spas,

fountains, ice machines, vegetable misters, dental devices and even shower heads

(Atlas, 1999). In addition, the presence of dead-end loops, stagnation in plumbing

systems and periods of non-use or construction have been shown to be technical

risk factors (Ciesielski et al., 1984; Atlas, 1999). Also, the material of the piping

system has been shown to influence the occurrence of high bacterial

concentrations. In this respect, the use of copper as plumbing material may help to

minimize the risk of legionellosis whereas plastic materials support high numbers

of L. pneumophila (Rogers et al., 1994).

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2.1.5.2 Distribution of Legionella in Singapore

The first L. pneumophila was isolated from hospital cooling towers in Singapore

(Nadarajah and Goh, 1986). Subsequently, Meers et al. (1989) took 87 water

samples from 48 cooling-towers on 15 sites and isolated 19 strains of Legionella

from 7 of the sites. All of the isolates are known to cause legionellosis (Yu et al.,

2002). 53% of the isolates belong to L. pneumophila serogroup 1, which is the

dominant causal agent for both community- and hospital-acquired legionellosis

(Fang et al., 1989; Yu et al., 2002).

For the period of 1991 to 1996, the overall isolation rate of Legionella bacteria

was 36% (1107 positive samples / 3095 samples taken) for cooling towers, 33%

(2/6) for public showers, 29% (30/103) for indoor decorative fountains, 15%

(10/68) for outdoor decorative fountains, 15% (4/26) for outdoor man-made

decorative waterfalls and 2% (1/48) for spa pools (Heng et al., 1997). The

isolation rate was not correlated with rainfall. The majority of the isolates (85.6%)

belonged to L. pneumophila while 46.9% belonged to serogroup 1.

Based on the samples collected during epidemiologic investigations in the period

of 1998 to 2002, Legionella bacteria were isolated from 550 (59.6%) of 923

cooling towers, 41 (38.3%) of 107 water fountains, 6 (16.2%) of 37 mist fans and

23 (23.7%) of 97 water taps/shower heads (Goh et al., 2005). Of 188 Legionella

bacteria isolated from cooling towers, L. pneumophila was found to be the

predominant species (65.4%) while 50.4% belonged to serogroup 1. The non-

pneumophila isolates are known to cause legionellosis (Yu et al., 2002) and they

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were L. bozemanii (14.9%), L. anisa (6.4%) and L. dumoffii (1.6%). Legionella

bacteria count was also found to be generally low, with 39 (20.7%) cooling towers

exceeding 10 CFU/ml.

2.1.5.3 Association of Legionella with protozoa

A universal trait of legionellae and LLAP organisms is their intracellular

existence. These bacteria are capable of infecting and multiplying within a variety

of mammalian and protozoan cell lines (Fields, 1996). Most of these studies were

conducted with L. pneumophila, primarily because this species is responsible for

the majority of legionellosis (Marston, 1994). It appears that L. pneumophila may

also have the most extensive host range of legionellae (Fields, 1996).

Protozoa do not only provide nutrients for the intracellular legionellae, but also

represent a shelter when environmental conditions become unfavorable (Thomas

et al., 2004). Compared to in vitro grown L. pneumophila, amoeba-grown bacteria

have been shown to be highly resistance to chemical disinfectants and to treatment

with biocides (Barker et al., 1992). Particularly inside Acanthamoeba cysts, the

bacteria are able to survive high temperatures, disinfection procedures and drying

(Rowbotham, 1986; Kilvington and Price, 1990; Winiecka-Krusnell and Linder,

1999). Furthermore, cooling tower amoebae containing legionellae may adapt to

biocides and may even be stimulated by biocides (Srikanth and Berk, 1993).

Legionella may also use protozoa to colonize new habitats where inhaled protozoa

represent a vehicle for effective transmission to humans (Cirillo et al., 1994). In

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addition, these vehicles of respirable size of 1-5µm containing L. pneumophila are

highly resistant to biocides (Berk et al., 1998). Interestingly, the same study

showed that more intense vesicle formation has been noticed before encystations

and when the amoeba are exposed to a mixed bacterial population, corresponding

to the conditions occurring in their natural environment. Interaction of Legionella

and protozoa also contributes and enhances the infection process itself (Cirillo et

al., 1994; Cirillo et al., 1999). However, the underlying mechanisms of this

phenomenon are not well elucidated yet (Steinert et al., 2002). In addition,

Brieland et al. (1997) demonstrated that L. pneumophila-infected amoeba were

more pathogenic than an equivalent number of bacteria or co-inoculum of the

bacteria and amoeba. A passage through Acanthamoeba castellanii was found to

reactivate viable but non-culturable (VBNC) Legionella into culturable state

(Steinert et al., 1997).

While protozoa are the natural hosts of legionellae, the infection of human

phagocytic cells is opportunistic (Fields et al., 2002). Much of our understanding

of the pathogenesis of legionellae has come from an analysis of the infection

process in both protozoa and human host cells. Studies contrasting the role that

virulent factors play in these two host populations allow speculation on the

bacteria’s transition from their obligatory relationship with protozoa to their

opportunistic relationship with humans.

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2.1.5.4 Association of Legionella with biofilm

Biofilms are the primary site of Legionella growth and persistence. In natural

hydrothermal areas, Legionella spp. were isolated in higher numbers from

biofilms than from water (Marrão et al., 1993). Similarly, in a man-made model

potable water system, the bacteria are more easily detected from swab samples of

biofilm than from flowing water (Rogers et al., 1994). Thus, suggesting that the

majority of the legionellae are biofilm associated. Furthermore, the association of

Legionella to biofilm may explain, at least in part, why legionellae are relatively

hard to eradicate in water systems, as biofilms exhibit a marked resistance to

biocidal compounds and chlorination (LeChevallier et al., 1988).

Only a limited number of studies attempted to characterize the bacteria’s

association within these complex ecosystems (Rogers and Keevil, 1992; Walker et

al., 1993; Rogers et al., 1994; Rogers et al., 1995). Rogers and Keevil (1992)

demonstrated that legionellae occurred in microcolonies within aquatic biofilm in

the absence of amoebae, thus providing the first evidence that legionellae is able

to grow extracellularly within the biofilm. Walker et al. (1993) evaluated the

effect of surface materials on growth of L. pneumophila using gas

chromatography-mass spectrometry analysis of genus-specific hydroxy fatty

acids, while Rogers et al. (1994) evaluated the effect of temperature and surface

materials on the growth of L. pneumophila. Rogers et al. (1995) used biofilm

models to evaluate silver efficacy against L. pneumophila and this study

represents a vast improvement over previous studies, which primarily evaluated

the susceptibility of agar-grown bacteria in sterile water.

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Majority of Legionella-biofilm studies that have been conducted employed

naturally occurring microbial communities. Such studies have the advantage of

representing a true and natural microbial community, but not all the organisms

present have been identified and their contribution to the survival and

multiplication of legionellae remains unknown. Nevertheless, biofilm matrices are

known to provide shelter and a gradient of nutrients. Complex nutrients available

in the biofilm may result in the multiplication of legionellae. Commensal bacteria

such as Flavobacterium breve, an environmental Pseudomonas, Alcaligenes and

Acinetobacter and blue-green algae (Cyanobacterium spp.) can stimulate the

growth of Legionella in the aquatic environment (Tison et al., 1980; Stout et al.,

1985; Wadowsky and Yee, 1985). Understanding the conditions under which L.

pneumophila can multiply extracellularly could have tremendous impact on

control strategies for the prevention of legionellosis.

Murga et al. (2001) attempted to detect extracellular growth of L. pneumophila by

using a biofilm reactor and a defined bacterial biofilm grown on non-

supplemented potable water. The base biofilm was composed of Pseudomonas

aeruginosa, Klebsiella pneumoniae and the Flavobacterium-like organism

isolated from a water sample containing legionellae. L. pneumophila was found to

associate with and persist in these biofilms with and without Hartmanella

vermiformis. However, L. pneumophila cells did not appear to develop

microcolonies and growth measurement studies indicate that L. pneumophila did

not multiply within this biofilm in the absence of amoebae. Nonetheless, L.

pneumophila did multiply in the biofilm and planktonic phase in the presence of

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H. vermiformis and the majority of these bacteria appeared to be shed into the

planktonic phase. These studies suggest that L. pneumophila may persist in

biofilms in the absence of amoebae, but in the model, the amoebae were required

for multiplication of the bacteria. Nevertheless, additional studies are needed to

determine if legionellae possess a means to multiply independent of a host cell

within biofilms.

Langmark et al. (2005) studied the accumulation and fate of microorganisms and

microspheres in biofilms formed in a novel pilot-scale water distribution system.

They demonstrated that the accumulation of L. pneumophila is independent of the

indigenous biofilm cell density; instead it is dependent on particle surface

properties, where hydrophilic spheres accumulated to a larger extent than

hydrophobic ones. Although combined chlorine concentration exceeding 0.2mg/L

inhibited the establishment of culturable L. pneumophila within system, the loss of

fluorescence in situ hybridization-positive cells closely resembled that of inert

fluorescent microspheres instead. Thus implying that the fate of culturable

legionellae within the system is best described in terms of loss of culturability

rather than physical desorption. Nevertheless, it is unknown if those persisting

legionellae were still viable despite the loss of culturability. Last but not least, this

study demonstrated that desorption is one of the primary mechanisms affecting the

fate of microspheres and legionellae in biofilms, followed by disinfection and

biological grazing.

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2.1.5.5 Interaction of Legionella with Pseudomonas spp.

Inhibitive effect of P. aeruginosa on Legionella has been reported (Hussong et al.,

1987; Gomez-Lus, 1993). In addition, Leoni and Legnani (2001) sampled hot

water supplies in public buildings and found that there is an inverse correlation (r

= -0.26; p < 0.05) between the concentration of Legionella spp. and that of P.

aeruginosa. Interestingly, Kimura et al. (2005) reported that the quorum-sensing

signal, N-(3-oxododecanoyl)-L-homoserine lactone (3OC12-HSL) of P.

aeruginosa suppressed growth of L. pneumophila in a dose-dependent manner. In

addition, significant suppressions of virulence factor genes (dotA, rtxA and lvh)

were demonstrated in L. pneumophila exposed to 3OC12-HSL.

Most recently, in an attempt to determine the fate of L. pneumophila when

introduced to single species or mixed cultures of defined heterotrophic bacteria,

Mampel et al. (2006) reported no attachment of L. pneumophila to a 2-day old

biofilm formed by the Pseudomonas spp. in a flow chamber system.

2.2 Biofilm

2.2.1 Introduction to biofilm

Putative biofilm microcolonies have been identified by morphology in the 3.3-3.4

billion year old South African Kornberg formation (Westall et al., 2001). In the

context of evolution and adaptation, biofilms appeared to provide homeostasis in

the face of the fluctuating and harsh conditions of the primitive earth, thereby

facilitating the development of complex interactions between individual cells

(Hall-Stoodley et al., 2004).

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2.2.2 General characteristics of biofilm

Biofilms develop on surfaces in diverse environments, both natural and man-

made, and they may contaminate industrial pipelines, cooling water towers, dental

unit water lines, catheters, ventilators and medical implants, causing biofouling

and significant financial losses (Characklis, 1990a; Flemming, 2002). Occurrence

in such wide range of environments, where a surface is exposed to adequate

moisture; therefore it is difficult to generalize about their structure and

physiological activities (Sutherland, 2001). Furthermore, biofilms can be

composed of a population that developed from a single species or a community

derived from multiple microbial species (Sutherland, 2001).

Nevertheless, Davey and O'Toole (2000) noted that biofilms formed from single

species in vitro and those produced in nature by mixed species consortia exhibit

similar overall structural features. Most biofilms have been found to exhibit some

level of heterogeneity in that patches of cells in biofilm microcolonies are held

together by an extracellular polysaccharides (EPS) matrix that varies in density,

creating open areas where water channels are formed. The ability of these

channels to facilitate efficient nutrient uptake by infusing fluid from the bulk

phase into the biofilm (Stoodley et al., 1994), thereby optimizing nutrient and

waste-product exchange, provided the first link between form and function.

EPS composition is complex, presumably varies from organism to organism and

includes polysaccharides, nucleic acids and proteins (Sutherland, 2001). The

nature of the matrix is dependent on both intrinsic and extrinsic factors, where the

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former arises in accordance with genetic profile of component microbial cells and

the latter includes the physico-chemical environment in which the biofilm and its

matrix are located (Sutherland, 2001).

Now, many laboratory biofilm studies have concentrated on laboratory models in

which Pseudomonas aeruginosa has been the microbial species of interest

(Sutherland, 2001), because it is a ubiquitous Gram-negative environmental

bacterium that forms biofilms on wet surfaces (Stover et al., 2000).

2.2.3 Biofilm development

Today, the combination of high resolution three-dimensional imaging techniques,

specific molecular fluorescent stains, molecular-reporter technology and biofilm-

culturing apparatus has shown that biofilms are not simply passive assemblages of

cells that are stuck to surfaces, but are structurally and dynamically complex

biological systems whose cells express genes in a pattern that differs profoundly

from that of their planktonic counterparts (Sauer et al., 2002).

Proteomic studies indicated that biofilm formation in Pseudomonas aeruginosa

proceeds as a regulated developmental sequence and five stages have been

proposed (Sauer et al., 2002, Stoodley et al., 2002). Briefly, stages one and two

are generally identified by a loose or transient association with the surface,

followed by robust adhesion. Stages three and four involve the aggregation of

cells into microcolonies and subsequent growth and maturation. Lastly, stage five

is characterized by a return to transient motility where biofilm cells are sloughed

Literature Review


or shed. The developmental life cycle comes full circle when dispersed biofilm

cells revert to the planktonic mode of growth. Microscopic observations

demonstrated that bacteria having physiologies from more than one stage of

biofilm could be present simultaneously within the biofilms (Sauer et al., 2002).

There is also evidence for developmental sequences in Escherichia coli (Reisner

et al., 2003) and Vibrio cholera biofilms (Watnick and Kolter, 1999). Until

recently, it was thought that highly regulated social behaviour in prokaryotes was

an unusual feature of myxobacteria only (Kaiser, 2003).

2.2.4 Stages of biofilm development

2.2.4.1 Stage 1: Reversible attachment

Initial event in biofilm development occurs when contact is made between the

surface of the cell and an interface. Using nonmotile mutants, a significant

decrease in attachment efficiency compared to flagellated cultures was observed

under continuous flow (Sauer et al., 2002). A similar observation was made

previously by O’Toole and Kolter (1998) for biofilms grown under static biofilm

conditions. Therefore, flagella-mediated motility appears to be required for a

planktonic bacterium to swim toward a surface and to initiate reversible (or

transient) attachment (O'Toole and Kolter, 1998) via the cell pole (Sauer et al.,

2002).

Individual adherent cells that initiate biofilm formation on a surface are

surrounded by only small amounts of EPS (Stoodley et al., 2002). Davies and

Literature Review


Geesey (1995) showed that within 15 minutes of P. aeruginosa cell initial contact

with substratum, the cluster of genes responsible for alginate production is

upregulated and that this genetic event initiates the process of biofilm formation.

2.2.4.2 Stage 2: Irreversible attachment

In continuous flow, irreversible attachment is indicated by cessation of motility of

the attached cells (Sauer et al., 2002) and the cells are associated to the substratum

via the long axis of the cell body (Caiazza and O’Toole, 2004). The earliest time

of onset of irreversible attachment was 2 hrs (Sauer et al., 2002). Caiazza and

O’Toole (2004) demonstrated that SadB locus is required for the transition from

reversible to irreversible attachment in P. aeruginosa. However, the exact

mechanism by which SadB promotes this transition is unknown.

Furthermore, the cell clusters formed during this stage were observed to remain

attached to the substratum through to the last stage of biofilm development (9 to

12 days of incubation) (Sauer et al., 2002). In the same study, the Las quorum-

sensing system became active upon irreversible attachment, as determined by

onset of reporter activity for the lasB gene, which has been shown to be

responsive to induction by PAI-1 autoinducer (Pearson et al., 1994). However,

proteomics study and in situ observation of lasB:lacZ reporter gene demonstrated

that cells of P. aeruginosa which are in the planktonic and early attachment stage

did not display substantially different physiologies (Sauer et al., 2002). Therefore,

this implies that quorum sensing does not have an influence on this transitional

stage of biofilm.

Literature Review


An earlier study by Davies et al. (1998) demonstrated that the Las quorum-

sensing system was involved in the development of P. aeruginosa biofilm. A lasI

mutant formed flat, undifferentiated biofilms that unlike wild-type biofilms are

sensitive to the biocide sodium dodecyl sulphate (SDS). In addition, mutant

biofilms appeared normal when grown in the presence of a synthetic signal

molecule. The differences in biofilm architecture imparted by las inactivation

were found to be related to changes in the structure of EPS (Sauer et al., 2002 and

Davies et al., 1998).

2.2.4.3 Stage 3: Maturation-1

After 3 days of biofilm development, cell clusters became progressively layered

and was defined as the point in time at which cell clusters are thicker than 10µm

(Sauer et al., 2002). In addition, this maturation-1 stage is accompanied by the

activation of the Rhl quorum-sensing system which was determined by the onset

of reporter activity for the rhlA gene (Sauer et al., 2002). The rhlA gene was

shown to be induced by the PAI-2 autoinducer (Pearson et al., 1995).

2.2.4.4 Stage 4: Maturation-2

As defined by Davies et al. (1998), the penultimate stage in biofilm development

is reached when cell clusters attain their maximum average thickness. In the

reactor study conducted by Sauer et al. (2002) reported that this stage of P.

aeruginosa PAO1 biofilm development was reached after 6 days of growth in

minimal media supplied with glutamic acid as sole carbon source. In the same

study, it was microscopically observed that cells within clusters were nonmotile,

Literature Review


the cell clusters reach their maximum dimensions, the majority of the cells are

segregated within cell clusters and clusters are displaced from the glass surface.

This is also the point at which biofilm bacteria are profoundly different from

planktonic bacteria with respect to the number of differentially expressed proteins.

More than 50% of all detectable proteins undergo changes in regulation between

planktonic growth and maturation-2 stage growth, with the majority being

upregulated. Many cells alter their physiological processes (e.g., grow

anaerobically) in response to conditions in their particular niches (Stoodley et al.,

2002)

2.2.4.5 Stage 5: Dispersion

After 9 days, Sauer et al. (2002) noted that cell clusters undergo alterations in

their structure due to the dispersion of bacteria from their interior portions. These

bacteria were motile and were observed to swim away from the inner portions of

the cell cluster through openings in the cluster and enter the bulk liquid, leaving

behind structures that appear shell-like, with a hollow center and walls of

nonmotile bacteria. Presumably, this dispersion allows cells to swim back into the

bulk liquid to gain better access to nutrients for the cells that remain in the

biofilm. Sauer et al. (2002) also noted that the protein patterns for this stage are

closer to that for planktonic bacteria than for maturation-2 stage cells and the

transition to dispersion phase is the only episode in biofilm development where

more proteins were downregulated than upregulated.

Literature Review


Recently, new levels of multicellular organization have been observed inside

mature microcolonies. These features included localized dissolution of the biofilm

matrix (Sauer et al., 2002 and Tolker-Nielsen et al., 2000) and dispersal of a

subpopulation of cells from internalized portions of the microcolony and death of

a subpopulation of cells inside the microcolony (Webb et al., 2003).

2.2.5 Determinants of biofilm structure

There is a continuing debate among biofilm researchers concerning the relative

contributions of genetics (active response) and environmental conditions (passive

response) to the development of biofilm structure and development (Kjelleberg

and Molin, 2002).

Davies et al. (1998) demonstrated that a quorum-sensing signal, N-(3-

oxododecanoyl)-L-homoserine lactone (3OC12-HSL), associated with the

production of virulence factors, is involved in the differentiation of individual

cells of P. aeruginosa PAO1 into complex multicellular structures, thus opening

the concept that biofilm structure was genetically regulated. A mutation that

blocks generation of the signal molecule hinders differentiation and the resulting

flat biofilm appears to be sensitive to detergent biocide SDS. Molecular

techniques, such as random transposon mutagenesis and knockout mutant studies,

have since been used extensively to identify ‘biofilm-specific’ genes (Heydorn et

al., 2002, Sauer et al., 2002 and Klausen et al., 2003a).

Literature Review


On the other hand, environmental factors such as hydrodynamics and nutrient load

have major impacts on biofilm structure. Stoodley et al. (1999) reported that under

conditions of low-shear laminar flow, the biofilm consisted of a monolayer of

cells with mound-shaped circular microcolonies but under high-shear, turbulent

flow conditions, the biofilm formed filamentous streamers. In addition, a study by

Liu and Tay (2001) found that biofilms grown at higher shear were smoother and

denser than those grown at lower shear. Klausen et al. (2003a) demonstrated that

when citrate was used as a carbon source, P. aeruginosa PAO1 formed a flat,

uniform biofilm, whereas when glucose was used, P. aeruginosa PAO1 formed a

heterogeneous biofilm containing mushroom-shaped multicellular structures

separated by water-filled channels.

van Loosdrecht et al. (2002) successfully modeled the structural and temporal

complexity of biofilms using simple rules that are based on localized growth

patterns determined by the distribution of nutrients and fluid shear. In another

words, biofilm differentiation into mature biofilms of organized communities with

functional heterogeneity does not necessarily require a genetic programme, but

may in fact constitute the sum of a large number of cellular adaptations and

growth cycles influenced by environmental factors.

Also, Purevdorj et al. (2002) demonstrated that environmental factors such as

hydrodynamics, can ‘override’ cell-cell communications as a principal

determinant of biofilm structure, illustrating that biofilm development is a

multifactorial process influenced by both environmental and genetic factors.

Literature Review


2.2.6 Microbial diversity of biofilms

Microorganisms are found in a wide range of diverse ecosystems as highly

structured, multi-species biofilms (Stoodley et al., 2002). Such bacterial

communities in nature play a key role in the production and degradation of

organic matter, the degradation of many environmental pollutants, and the cycling

of nitrogen, sulphur and many metals. Most of these processes require the

concerted effort of bacteria with different metabolic capabilities and it is likely

that bacteria residing within biofilm communities carry out many of these

complex processes (Davey and O’Toole, 2000). Particularly, microbial diversity is

a property that is important with regard to the potential of opportunistic pathogens

such as Legionella spp. to integrate into biofilms.

It has been shown that hydrodynamic shear rates affect biofilm diversity as well as

the relative proportions of aggregating bacteria (Rickard et al., 2004). Highest

proportion of autoaggregating bacteria was present at high shear rates, while

intermediate shear rate selected for the highest proportion of coaggregating

bacteria. Coaggregation (interactions between two genetically distinct planktonic

microorganisms) between freshwater bacteria is mediated by growth-phase-

dependent lectin-saccharide interactions, which are optimal in stationary phase

cultures (Rickard et al., 2000). Additionally, coaggregation often occurs between

bacteria that are taxonomically distant (intergeneric coaggregation) and

occasionally between strains belonging to the same species (intraspecies

coaggregation) (Rickard et al., 2002), and enhances the development of

freshwater multi-species biofilm (Rickard et al., 2003).

Literature Review


2.2.7 Microbial positioning in biofilm

The natural habitats prokaryotes are remarkably diverse (Pace, 1997). Their ability

to persist throughout the biosphere is due, in part, to their unequaled metabolic

versatility and phenotypic plasticity. One key element of their adaptability is their

ability to position themselves in a niche where they can propagate (Fenchel, 2002).

Numerous positioning mechanisms have been discovered. The most common

mechanism is flagellar motility and different methods of surface translocation,

including twitching, gliding, darting and sliding (Henrichsen, 1972).

Apart from active motility mechanisms, there are other mechanisms utilized by

bacteria to position themselves in response to their environment, such as

aggregation and attachment (Davey and O’Toole, 2000). Aggregation enhances

cell-cell interaction as well as the sedimentation rate of cells. Through attachment,

the bacteria not only position themselves on a surface, they can form communities

and obtain the additional benefit of the phenotypic versatility of their neighbours.

Cells of particular species are found consistently in certain locations, near the

colonized surface or at the apices of mushroom-structures of the biofilm. The

sessile cells that comprise single-species biofilms are located within the

microcolonies in species-specific distribution patterns, such that the

preponderance of the sessile cells may be found in the caps of mushrooms in a

highly organized pattern and the stalks of mushrooms formed by some species are

virtually devoid of cells (Stoodley et al., 2002).

Literature Review


Only a few reports with regards to cell movement within biofilm matrix have been

made. Okabe et al. (1996) demonstrated that trapped tracer beads were gradually

transferred from the depth of the biofilm to the surface. Klausen et al. (2003b)

demonstrated that the mushroom-shaped multicellular structures in P. aeruginosa

PAO1 biofilms are formed in a sequential process involving a non-motile bacterial

subpopulation and a migrating bacterial subpopulation. The non-motile bacteria

form the mushroom stalks by growth in certain foci of the biofilm while the

migrating bacteria form the mushroom caps by climbing the stalks and

aggregating on the tops in a process driven by type-IV pili. In addition, a biofilm

model, which is derived by combining individual description of microbial

particles with a continuum representation of the biofilm matrix, suggests cells in

biofilm matrix move due to pushing mechanism between cells in colonies and by

an advective mechanism supported by the EPS dynamics (Alpkvist et al., 2006).

In this model, the EPS matrix is described by a continuum representation as

incompressible viscous fluid, which can expand and retract due to generation and

consumption processes.

2.3 Prevention of legionellosis

2.3.1 Control of legionellosis

Maintaining a clean system is of critical importance in reducing the risk of

legionellosis and it is the goal of a maintenance program to provide efficient

operation of the system while minimizing the risk of legionellosis through

preventing the amplification of Legionella (Fliermans, 1995). Since cooling

towers have been identified as one of the reservoirs and amplifiers for Legionella

Literature Review


bacteria (Shelton et al., 1994; Bentham and Broadbent, 1993; Garbe et al., 1985),

routine maintenance service, including visual inspections, and mechanical and

physical cleaning programs designed to maintain year-round system cleanliness,

are an important part of an effective water treatment program.

Although good maintenance may reduce the likelihood of Legionella

amplification, there is very little data to indicate that cleaning alone is effective in

controlling Legionella (Fliermans, 1995). Therefore chemical biocidal treatment is

required. In addition, microbiological monitoring for Legionella must be included

as part of the quality assurance / quality control program to insure effectiveness of

any control measures.

Well maintained cooling towers with proper water treatment have generally not

been associated with outbreaks of legionellosis (Fliermans, 1995). In Singapore,

the Ministry of the Environment published a Code of Practice for the Control of

Legionella Bacteria in Cooling Towers (4th edition) in 2001 that provided

guidelines to cooling tower monitoring and maintenance to minimize the risk of

outbreaks of LD here.

2.3.2 Detection of Legionella

Unfortunately, measurements of water quality such as total bacterial counts, total

dissolved solids and pH have not proven to be good indicators of Legionella levels

in cooling towers (Boss and Day, 2003). Cultivation of Legionella remains the

standard method of detection (Steinert et al., 2002). The most widely used growth

Literature Review


medium is commercially available buffered charcoal yeast extract agar, which is

supplemented with cysteine, iron salts and α-ketoglutarate. However, a number of

factors, including other bacteria, can interfere with the growth of Legionella, even

on selective media (Feeley et al., 1979; Edelstein, 1982). Serology-based methods

are not regarded to be the gold standard anymore since the progressive

characterization of new species has shown that antigen cross-reactivity limits

specificity (Maiwald et al., 1998). Further routine methods rely on pulsed field gel

electrophoresis (PFGE), amplified fragment length polymorphism (AFLP),

arbitrarily primed and nested PCR (Benson and Fields, 1998). Additionally, gas

chromatographic mass spectrometry based on the unique 3-hydroxy and 2,3-

dihydroxy fatty acids of the Legionella lipopolysaccharide has been described for

complex microbial consortia (Walker et al., 1993).

2.3.3 Risk assessment of cooling tower for Legionnaires’ disease outbreaks

From historical data compiled from outbreaks of LD worldwide, Shelton et al.

(1994) suggested that high numbers of Legionella were unusual and could be

equated to an increased risk of disease outbreak. Thus, routine monitoring of

Legionella counts and total bacterial counts were used to assess the risk of cooling

towers for potential LD outbreaks.

Other risk factors for determining the likelihood that a cooling tower may be

associated with human illness are not well defined. However, some towers appear

to be more likely to be associated with an outbreak of LD than other towers.

Fliermans (1995) provided a general guideline to risk assessment of cooling

Literature Review


towers based on the location of host population and potential susceptibility of

host. Monthy monitoring was recommended for cooling towers with the highest

risk, while yearly monitoring was recommended for those with the least risk.

2.3.4 Water treatment in cooling towers

The method of choice for controlling bacterial populations within cooling towers

remains the use of industrial biocides. These may be oxidizing or non-oxidizing

(Characklis, 1990b). Traditional oxidizing agents such as chlorine and bromine

have been proven effective in controlling Legionella in cooling towers

(Characklis, 1990b; Fliermans, 1995).

While continuous chlorination at 0.2-0.3ppm is effective against a wide range of

bacteria (Kuchta et al., 1983), such levels are generally not effective in removing

Legionella from a highly contaminated cooling tower system (Fliermans et al.,

1982). Early field investigations demonstrated the effectiveness of 1.5ppm free

residual chlorine for a short duration in reducing the levels of Legionella in large

industrial cooling towers (Fliermans et al., 1979; Fliermans et al., 1982;

Fitzgeorge and Dennis, 1983). However, levels of free residual chlorine above

1ppm may be corrosive to the metallurgy of a system (Fliermans, 1995). In

addition, high levels of chlorine may also form toxic by-products with organic

substances present in water and may be of environmental and public health

concern.

Literature Review


Although Fliermans and Harvey (1984) reported the lack of effectiveness of

continuous bromo-chloro-dimethyl-hydantoin (BCD) treatment at 2.0ppm free

residual levels against Legionella, Australian studies (Broadbent, 1993) indicated

that a slow release of the bromocide at 300ppm were effective in controlling the

growth of Legionella. The latter study also indicated that quaternary ammonium

compounds which were frequently used for biofouling control in cooling towers

were not effective in controlling Legionella. Among non-oxidizing agents, 2,2-

dibromo-3-nitropropionamide appears to be the most effective in controlling

Legionella in water systems (Kim et al., 2002).

New biocidal actives are slow to emerge due to regulatory and environmental

concerns, but novel methods of delivery and new synergistic combinations of

existing biocides are continually being investigated for use in controlling

microorganisms in various process streams, including cooling towers (Wright,

2000). However, only few studies have been conducted on the effectiveness of

various biocides in controlling Legionella under field conditions (England et al.,

1982; Fliermans and Harvey, 1984; Elsmore, 1986; Yamamoto et al., 1991; Prince

et al., 2002). Since sensitivity testing of L. pneumophila suspended in tap water to

biocides cannot predict culture results from biocide-treated cooling towers

(England et al., 1982) and the application of biocides at concentrations

recommended by the manufacturer may not be able to reduce L. pneumophila in

cooling towers to source water concentration (Fliermans and Harvey, 1984), it is

necessary monitor Legionella counts to verify the effectiveness of new biocides

under field conditions.

Literature Review


Recurrence of Legionella in biocide-treated cooling towers has been reported

(Kurtz et al., 1982) and without biocide treatment, Legionella bacteria may reach

densities that present a health risk (Negron-Alvira et al., 1988). Furthermore,

Bentham (2000) found that the culture results from Legionella samples taken from

the same systems 2 weeks apart were not statistically related in 25 out of 28

cooling tower systems studied, suggesting that the determinations of health risks

from cooling towers cannot be reliably based upon single or infrequent Legionella

tests. Considering the ubiquity of Legionella, it is prudent to diligently execute

periodic application of industrial biocide to ensure control of Legionella growth

within these devices, while frequently monitoring the effectiveness of control

measures applied.

Materials and Methods


Chapter 3: Materials and Methods

3.1 Bacterial strains and culture

3.1.1 Bacterial Strains

Legionella pneumophila subsp. pneumophila, Philadelphia-1 (ATCC® 33152) was

provided by courtesy of Dr Gamini Kumarasinghe from the Department of

Laboratory Medicine, National University Hospital, Singapore. The wild type P.

aeruginosa PAO1 was a generous gift from Associate Professor Zhang Lian Hui

from Institute of Molecular and Cell Biology, Singapore while Pseudomonas

aeruginosa PAO1 tagged with cyan-fluorescent-protein (PAO1-CFP) was

generously granted by Associate Professor Tim Tolker-Nielsen from BioCentrum-

DTU, The Technical University of Denmark, Denmark (Klausen et al., 2003a).

3.1.2 Culture Media

In this study, L. pneumophila was cultured on Buffered Charcoal Yeast Extract

agar supplemented with ferric pyrophosphate, α-ketoglutarate and L-cysteine

(Edelstein BCYE agar, Oxoid Limited, U.K.) and in also Edelstein BCYE liquid

media (Appendix I). P. aeruginosa PAO1 was cultured on both Luria Bertani (LB;

Appendix II) media and minimal media (MM; Appendix III), where the latter is a

defined medium. Similarly, P. aeruginosa PAO1-CFP was cultured on LB and

MM, both media supplemented with 60µg/ml gentamicin (Sigma-Aldrich, U.S.A.;

LB+gen and MM+gen respectively) in order to select for the fluorescent tagged

cells.



3.1.3 Maintenance of stock cultures

L. pneumophila was grown on Edelstein BCYE agar at 37°C for at least 2 days. P.

aeruginosa PAO1-CFP and P. aeruginosa PAO1 were grown on LB+gen and LB

agar respectively, and were incubated at 37°C for 1 day. After incubation, all the

agar plates were stored at 4°C for 3 weeks before subculturing.

To prepare cryogenized stock strains, L. pneumophila and P. aeruginosa cells at

late-log phase were harvested at 24 and 28hrs of growth respectively. Cryogenized

stocks were then prepared by suspending the cells of each strain in a final

concentration of 25% glycerol in fresh media. Finally, the cell suspensions were

dispensed into NUNCTM CryoTube Vials (NUNC, Denmark) and stored at -70°C.

3.2 Growth kinetic studies

3.2.1 Growth kinetics of L. pneumophila

Overnight culture of L. pneumophila was inoculated into fresh Edelstein BCYE

liquid medium at a 1:50 ratio and incubated at 37°C with shaking at 120rpm.

Subsequently, optical density (OD) of the culture was taken every hourly, using a

spectrophotometer (Shimadzu, Japan) at λ = 600nm. Dilutions were performed

when OD exceeds 0.5 and this study was triplicated.



3.2.2 Growth kinetics of P. aeruginosa PAO1

Overnight culture of P. aeruginosa PAO1 was inoculated into fresh MM medium

at a ratio of 1:100 and incubated at 30°C with shaking at 120rpm. Subsequently,

OD600nm of the culture was taken every hourly, using spectrophotometer. Dilutions

were performed when OD exceeds 0.5 and this study was triplicated.

3.2.3 Growth kinetics of P. aeruginosa PAO1-CFP

Overnight culture of P. aeruginosa PAO1-CFP was inoculated into fresh

MM+gen medium at a 1:100 ratio and incubated at 30°C with shaking at 120rpm.

Subsequently, OD600nm of the culture was taken every hourly, using

spectrophotometer. Dilutions were performed when OD exceeds 0.5 and this study

was triplicated.

3.3 Determination of the influent flow rate (Q) for continuous

culture in CDC Biofilm Reactor (CBR)

Maximum specific growth rate (µmax) of P. aeruginosa PAO1-CFP was

determined by obtaining the gradient of exponential growth from the graph of

OD600nm against time. Doubling time (td) of P. aeruginosa PAO1-CFP was

calculated using the following formula: µmax = (ln2)/td. The determination of the

nutrient influent flow rate (Q) to be used for continuous system required that the

hydraulic residence time, θ is less than td and the following formula was applied: θ

= V/Q (where V is the maximum volume of bulk fluid in CBR during continuous

flow).



3.4 Optimization of labelling processes

3.4.1 Optimization of L. pneumophila labelling with CFDA-SE

L. pneumophila cells at late-log phase were harvested at 24 hrs of growth and

washed with PBS (Appendix IV). Using PBS, cell concentration was adjusted to

approximately 109 colony forming units (CFU)/ml (corresponding to 10×

concentrate of cell suspension at OD600nm = 1) before a final concentration of

10µM 5-(and-6)-carboxyfluorescein diacetate, succinimidyl ester (CFDA-SE;

Molecular Probes Inc., U.S.A.; Appendix V) was added. The mixture was mixed

well, dispensed into 3 tubes and incubated at 37°C in the dark with shaking at

120rpm for 20, 30 and 40 mins respectively. To terminate the labelling process,

the L. pneumophila cells were centrifuged at 5,000g for 10 mins and washed twice

with PBS to remove residual CFDA-SE. A portion of the labelled cells were

plated onto BCYE agar using Miles and Misra method (Harrigan, 1998) to ensure

that the cells remained viable after the labelling process, while the remaining

portion were fixed with 1% formaldehyde (Merck, Germany) at 4°C overnight,

before analysis with flow cytometry. A tube of L. pneumophila cells that were

mock-treated with PBS instead of CFDA-SE served as a negative control and

blank.

3.4.2 Optimization of planktonic P. aeruginosa PAO1-CFP labelling with PI

P. aeruginosa PAO1-CFP cells at late-log phase were harvested at 28 hrs of

growth, fixed with 1% formaldehyde at 4°C overnight and washed with PBS.

Using PBS, cell concentration was adjusted to approximately 107 CFU/ml

(corresponding to P. aeruginosa PAO1-CFP cell suspension at OD600nm = 0.5) and



separated into 2 portions. Final concentrations of freshly prepared 0.1 and

1.0mg/ml propidium iodide (PI; Sigma-Aldrich, U.S.A.) in PBS were then added

into each portion respectively. After vortexing, the mixtures were incubated at

room temperature in the dark and immediately analyzed using flow cytometry at 5

mins interval each. A tube of P. aeruginosa PAO1-CFP that were similarly

processed but mock-treated with PBS instead of PI served as a negative control

and blank.

3.4.3 Flow cytometry

A total of 10,000 cells were analyzed using flow cytometry, FACSVantageTM SE

(Becton Dickinson, U.S.A.) operated on CellQuest program. The WinMDI

Version 2.8 software was used to plot histograms with number of events against

green CFDA-SE fluorescence in 4 decade log (as FL1-H) or red PI fluorescence in

4 decade log (as PMT4 Log).

3.4.4 Optimization of P. aeruginosa PAO1-CFP biofilm labelling with PI

Approximately 107 CFU/ml CFDA-labelled L. pneumophila was inoculated into 7

days old P. aeruginosa PAO1-CFP biofilms (grown in CDC Biofilm Reactor

continous culture system) and allowed to adhere without continuous flow for 1hr.

After allowing for re-stabilization of the continuous system for 3 hours, a coupon

(on which the biofilm was formed) was harvested and soaked in 6ml 4%

paraformaldehyde (PFA) for 30mins in the dark. Then, freshly prepared 600µl of

1mg/ml PI was added into the 4% PFA, and mixed gentle (taking care not to

disturb the biofilm), and incubated at room temperature for 5, 15 and 30 mins in



the dark. The staining process was terminated by transferring the coupon into

60ml sterile PBS contained in a standard petri dish, with the biofilm surface facing

upwards. A biofilm mock-treated with PBS served as a negative control.

These coupons were then viewed using confocal laser scanning microscope

(CLSM). For PI stains, image scanning was carried out with 543nm laser line

from a HeNe-G laser. To reduce background, emission filter BA-610IF was used.

Similarly, for CFP and CFDA detection, image scanning was carried out with the

405nm laser line from a LD405 laser and 488nm laser line from an M-Ar laser,

respectively. Background was also reduced using BA430-460 and BA505-525

emission filters, respectively. Images of the biofilm in the x-y plane or sections

through the biofilm were generated using Olympus FLUOVIEW Ver.1.3 Viewer.

3.5 P. aeruginosa PAO1-CFP biofilm formation in CDC Biofilm

Reactor (CBR)

3.5.1 CDC Biofilm Reactor

The CBR (BioSurface Technologies Corp., U.S.A.) is a one litre glass vessel with

an effluent spout at approximately 400ml. Continuous mixing of the reactor’s bulk

fluid was provided by a Teflon baffled stir bar that was magnetically driven by a

CERAMAG Midi magnetic stirrer (Ika®, U.S.A.). An UHMW (ultra high

molecular weight) polyethylene lid supports 8 independent polypropylene coupon

holders. Each coupon holder houses 3 removable stainless steel coupons

(diameter: 12.7mm), which served as the biofilm growth surfaces, for a total of 24

sampling opportunities. The coupons experienced a consistent high shear from the



rotation of the baffled stir bar at 120rpm. The CBR operates as a continuous flow

stirred tank reactor (CFSTR), meaning that nutrients are continuously pumped

into and flow out of the reactor at the same rate.

3.5.2 Setup of CDC Biofilm Reactor assembly

Connector

Air vent

Baffled stir bar

Stainless steel

coupon

Coupon holder

Connector

Influent tap

Effluent tap

Air vent

Magnetic stirrer (120rpm)

Nutrient carboy

CBR

Air vent Peristaltic pump

Flow break

Incubator (30°C)

Waste carboy

Figure 3.1. Schematic diagram of the CDC Biofilm Reactor assembly used in this study. The arrow head indicated the direction of continuous flow when the peristaltic pump was switched on. Biofilm was formed on the surface of stainless steel coupons which was facing the baffled stir bar indicated by the striped coupons.



An autoclavable 10L carboy (Nalge Nunc International, U.S.A.) was used to

contain fresh media to feed into CBR. The carboy top was equipped with 2 barbed

fittings to accommodate the tubings for nutrient and air vent, HEPA-VENTTM

(Whatman, U.K.) attachment. Incorporation of an autoclavable connector before

peristaltic pump facilitated the changing of emptied nutrient carboy with another

that was filled with sterile 10L fresh media. Peristaltic pump, Masterflex® Digital

Console Pump (Cole-Parmer Instrument Company, U.S.A.), was only switched on

for pumping media into CBR during continuous flow phase and the Masterflex®

precision tubing (Cole-Parmer Instrument Company, U.S.A.) passing through the

pump head had an internal diameter of 14mm. Before the fresh media entered

CBR, a flow break (BioSurface Technologies Corp., U.S.A.) prevented backward

contamination of media carboy from CBR. The bulk fluid in CBR was well mixed

by the baffled magnetic stir bar and extra fluid in CBR was drained into the waste

carboy. Positioning of a connector after the spout of CBR allowed changing of a

filled waste carboy with an autoclaved empty carboy.

3.5.3 P. aeruginosa PAO1-CFP biofilm formation

An overnight culture of P. aeruginosa PAO1-CFP (grown in MM+gen at 37°C

with shaking at 120rpm) was inoculated into 400ml fresh MM medium in CBR at

a ratio of 1:100 under sterile conditions. The inoculated CBR was then operated as

a batch culture system at 30°C with 120rpm stirring for 24 hours, with closed

influent and effluent taps. After which, the CBR was switched to continuous

culture phase where both influent and effluent taps were released and a continuous

flow of fresh MM medium was pumped into the CBR at a constant influent flow



rate of 2.5ml/min (Refer to Chapter 4.2 for the determination of influent flow

rate). Kinetics of P. aeruginosa PAO1-CFP biofilm formation was monitored by:

• Taking planktonic and biofilm samples,

• Enumerating P. aeruginosa PAO1-CFP by plating onto LB+gen plates,

• Detecting contamination from exogenous source(s) by plating onto LB

plates,

• Visualizing biofilm structure using confocal laser scanning microscopy

(CLSM), and

• Image analysis using COMSTAT image analysis software package

(Heydorn A. et al., 2000)

For the study of P. aeruginosa PAO1-CFP biofilm formation, six independent

experiments were performed. For each experiment, there were 6 sampling points

as shown in table 3.1. For image data acquisition, at least 3 image stacks were

taken from 1 or 2 coupon samples at each time point.

Table 3.1. Sampling points of 6 independent experiments for the study of P. aeruginosa PAO1-CFP biofilm formation.

Days from start of continuous culture

Experiment 1 to 3 Experiment 4 to 6

2 3 4 5 6 7 8 9 10 11



3.6 Introduction of L. pneumophila into P. aeruginosa PAO1-CFP

biofilms

Approximately 109 CFU/ml L. pneumophila grown to late log phase, was stained

with 10µM CFDA-SE for 30 mins and washed with PBS twice, before a ratio of

1:100 was inoculated into the CBR containing developing (Day 3) or mature

biofilm (Day 7). A portion of the legionellae inoculum was serially diluted and

enumerated by immunofluorescence. The continuous flow was stopped for the

adhesion of L. pneumophila onto pre-formed P. aeruginosa PAO1-CFP biofilm

and restarted 1 hr later. Samples of bulk fluid and biofilm were taken immediately

before the addition of L. pneumophila, 3 hrs after the continuous flow was

restarted (to allow for re-stabilization of the continuous flow system) and

everyday for up to 5 days after inoculation. Adhesion and persistence of L.

pneumophila was monitored by:

• Enumerating L. pneumophila by immunofluorescence method,

• Visualizing legionellae distribution using confocal laser scanning

microscopy (CLSM), and

• Image analysis using COMSTATWCY image analysis software package.

At the same time, the surface-to-biovolume ratio and porosity distributions of the

biofilm were also monitored by applying COMSTATLAYER and

COMSTATWCY image analysis software package respectively. At least 3

independent experiments were carried out for each developing stage of biofilm

development. For image data acquisition, at least 3 image stacks were taken from

1 or 2 coupon samples at each time point.



3.7 Introduction of NALCO 7320 into developing and mature P.

aeruginosa PAO1-CFP biofilms containing L. pneumophila

According to the procedure mentioned in Chapter 3.6, L. pneumophila was

introduced into developing and mature biofilms on day 3 and day 7 respectively.

Twenty four hours later, the peristaltic pump was stopped and fresh culture media

with a final concentration of 100ppm of NALCO 7320 was connected to CBR

continuous flow system. At time zero, 315µl of 100,000ppm NALCO 7320 was

added into the CBR, yielding a final concentration of 100ppm NALCO 7320

within the CBR with the constant mixing by baffle. At the same time, the

peristaltic pump was switched on again, bringing in fresh supply of nutrients and

biofilm removing agent.

Samples of bulk fluid and biofilm were taken before and immediately after the

addition of NALCO 7320. Samples were also taken in the subsequent 4, 8, 12 and

24hrs. Persistence of P. aeruginosa PAO1-CFP and L. pneumophila was

monitored by:

• Enumerating P. aeruginosa PAO1-CFP and L. pneumophila by culture and

immunofluorescence methods respectively,

• Visualizing biofilm structure, biofilm porosity and legionellae distribution

using confocal laser scanning microscopy (CLSM), and

• Image analysis using COMSTAT and COMSTATWCY image analysis

software package respectively.



Two independent biofilm experiments were carried out for each developing stage

of biofilm development. However, at least 3 image stacks were taken from each of

the 2 coupon samples at each time point.

3.8 Monitoring of each organism in CBR continuous flow system

3.8.1 Preparation for sampling

In preparation of sampling under sterile conditions, 75% denatured ethanol was

sprayed on and around the CBR lid and allowed to air dry for approximately 2

mins. Disturbance to the air within the 30°C incubator was minimized to prevent

contamination of the continuous culture.

3.8.2 Taking samples

3.8.2.1 Sampling bulk fluid

For every sampling point, two 1ml planktonic samples were taken and processed

in parallel. To take planktonic samples, one of the 8 coupon holders was removed

and placed inside a 1L sterile glass bottle (for biofilm sampling) before inserting a

1ml pipette into the CBR to sample the bulk fluid repeatedly. Once removed, the

coupon holder with biofilm covered coupons cannot be replaced back into CBR

because air-water interval of the bulk fluid can detach the biofilm significantly.

For every coupon holders removed, a sterile rubber bung was used to stopper the

hole in the CBR lid to prevent the entry of exogenous contaminants.



3.8.2.2 Sampling biofilm

For every sampling point, one coupon holder accommodating 3 coupons was

removed. One out of the 3 coupons was used for enumeration by plating and the

rest were prepared for visualization by CLSM. The coupon holder was held

straight up and removed from the 1L bottle. Any drips were collected in a sterile

petri dish placed beneath the rod. All 3 set screws were loosen with sterile set

screw driver (BioSurface Technologies Corp., U.S.A.) to release the coupons,

which were then removed using sterile haemostat, being careful not to disturb the

biofilm on coupon surface that was facing the interior of CBR. Once the coupons

were removed, the coupon holder was re-inserted back to the CBR so as to

minimize any changes in the final volume of bulk fluid in the CBR.

3.8.3 Preparation of coupons

3.8.3.1 Preparation of coupons intended for enumeration

A coupon was held in place on a petri dish with a sterile haemostat and scraped on

the side that faced the baffle with a sterile toothpick. The loosened biofilm was

washed into an empty eppendorf tube using 1ml PBS and then the toothpick was

rinsed by swirling on the bottom of the tube.

3.8.3.2 Preparation of coupons intended for visualization by CLSM

A coupon was immersed in 6ml 4% paraformaldehyde (PFA; Appendix VI)

contained in small sterile tissue culture plates (35×10mm; NuncTM, Denmark) at

room temperature for 30 mins in the dark. Slow immersion of coupon into liquid

resulted in significant biofilm sloughing off, thus each coupon was held near to



the liquid surface using a haemostat with the biofilm surface facing upwards and

dropped directly into the liquid below. This was done as soon as the coupon was

removed from CBR so as to prevent drying up of the biofilm. Then, freshly

prepared 600µl of 1mg/ml PI was added into 6ml 4% PFA, pipetted up and down

for gentle mixing, and incubated at room temperature for 5mins in the dark. The

staining process was promptly terminated by dropping the coupon into 60ml

sterile PBS contained in a standard petri dish, with the biofilm surface facing

upwards.

Taking care not to touch the top surface of the coupon where the biofilm was to be

visualized, the bottom of coupon was dried using a tissue paper and then placed in

a humid chamber with the biofilm surface facing upwards. To preserve the

fluorescence in the samples, a 20×60mm coverslip with 20µl of FluorSaveTM

Reagent (Merck, Germany) dropped in the middle was inverted and mounted onto

the biofilm surface of the coupon. The mounted coupons were then stored at 4°C

in a humid chamber in the dark for not more than 1 week. These mounted coupons

were air dried at room temperature in the dark overnight before viewing under

CLSM.

3.8.4 Disaggregation by homogenization

For a more accurate enumeration of each organism in either planktonic or biofilm

samples, the samples were subjected to disaggregation by homogenization before

serial dilution. Firstly, 750µl of planktonic and biofilm samples were transferred

into sterilized disposable culture tubes (Asahi Techno Glass Corporation, Japan)



and homogenized at 20,500rpm for 30 secs using a sterile homogenizer probe, T

25 basic ULTRA-TURRAX® (Ika®, U.S.A.). The probe was cleaned between

samples by firstly rinsing for 30 secs at 20,500rpm in 10ml of sterile PBS,

followed by rinsing at 20,500rpm for 15 secs in 10ml of 75% ethanol. The probe

was then soaked in the ethanol for 1 min. Finally, the probe was rinsed two more

times with 10ml PBS at 20,500rpm for 30 secs each. Any excess liquid on the

probe was removed by gently tapping the tip of the probe against the inner wall of

the last tube containing PBS before inserting the probe into the sample.

3.8.5 Enumeration of each organism

3.8.5.1 Enumeration of P. aeruginosa PAO1-CFP by culture

The disaggregated planktonic and biofilm samples were serially diluted using PBS

as diluent and plated onto LB+gen plates respectively, using Miles and Misra

method (Harrigan, 1998) in duplicates. The plates were then incubated at 37°C for

24 hrs. Density of planktonic P. aeruginosa PAO1-CFP was expressed as Log10

[colony-forming units (CFU)/ml of bulk fluid] while that of biofilm P. aeruginosa

PAO1-CFP was expressed as Log10 (CFU/mm2). Respective calculations were

shown below:

Planktonic P. aeruginosa PAO1-CFP count

Log10 (CFU/ml) = Log10 [(Average CFU/drop of 25µl) × 40 × Dilution Factor]

Biofilm P. aeruginosa PAO1-CFP count

Log10 (CFU/mm2)

= Log10 {[(Average CFU/drop of 25µl) × 40 × Dilution Factor] / Area of coupon}



= Log10 {[(Average CFU/drop of 25µl) × 40 × Dilution Factor] / [π (12.7/2) 2]}

3.8.5.2 Enumeration of L. pneumophila by immunofluorescence

L. pneumophila counts were monitored by immunofluorescence method where L.

pneumophila Serogroup 1 Direct Fluorescent Antibody Kit (PRO-LAB

Diagnostics, Canada) was used. Briefly, 20µl of homogenized planktonic and

biofilm samples was placed on the 6mm diameter wells in the Bellco® Antibody

Slides (Bellco® Glass Inc., U.S.A.), air-dried and gently heat fixed. L.

pneumophila sergroup 1 DFA Reagent (FITC-monoclonal antibody conjugate)

was applied to each well and the slides were incubated in a moist chamber for 30

mins at 37°C in the dark. The slides were then rinsed with PBS to remove the

conjugates and rinsed with dH2O before air dried. Lastly the slides were mounted

and examined using a fluorescence microscope (Olympus, Japan) within 24 hrs.

FITC-labelled antibody-antigen complex was detected by exposing the slide to

ultraviolet light and L. pneumophila cells appeared as bright yellow-green bacilli

under a 40× objective. At least 3 fields were examined for legionellae count.

Wells containing only L. pneumophila cells and only P. aeruginosa PAO1-CFP

cells served as positive and negative control respectively.

Density of planktonic L. pneumophila was expressed as Log10 (cells/ml of bulk

fluid) while that of biofilm L. pneumophila was expressed as Log10 (cells/mm2).

Calculations were shown below:



Planktonic L. pneumophila count

Log10 (cells/ml)

= Log10 [(Average number of cells per field) × (Area of each well/Area of each

field*) × (50/concentration factor)]

= Log10 [(Average number of cells per field) × (π(3)2 / π(0.2)2) ×

(50/concentration factor)]

Biofilm L. pneumophila count

Log10 (cells/mm2)

= Log10 {[(Average number of cells per field) × (Area of each well/Area of each

field*) × (50/concentration factor)] / (Area of coupon)}

= Log10 {[(Average number of cells per field) × (π(3)2 / π(0.2)2) ×

(50/concentration factor)] / (π(12.7/2)2)}

* Diameter of each field of 40× objective was measured using a stage micrometer.

L. pneumophila adhesion to P. aeruginosa PAO1-CFP biofilm

Number of legionellae adhering to biofilm per coupon per 106 legionellae

inoculated into CBR

= (Number of legionellae per coupon/ Total number of legionellae inoculated into

CBR) × 106



L. pneumophila persistence in P. aeruginosa PAO1-CFP biofilm

Percentage legionellae remaining in biofilm

= (Biofilm legionellae count at day n/ Biofilm legionellae count on day of

inoculation) × 100%

where, n = number of days following legionellae inoculation

Loss of L. pneumophila from biofilm

Amount of legionellae loss from biofilm

= (Biofilm legionellae count before the addition of biocide - Biofilm legionellae

count at 24hrs of exposure to biocide)

Loss of L. pneumophila per unit biomass lost

Legionellae loss per unit biomass lost

= (Biofilm legionellae count before the addition of biocide - Biofilm legionellae

count at 24hrs of exposure to biocide)*1000 / (Bio-volume before addition of

biocide – Bio-volume at 24hrs of exposure to biocide)

= (cells/mm3)

3.8.6 Detection of exogenous contaminants

In addition to plating on LB+gen for P. aeruginosa PAO1-CFP counts, serially

diluted samples were also plated on LB for detection of exogenous contaminants

using Miles and Misra method. Similarly, these plates were incubated at 37°C for

24 hrs before observation.



3.8.7 Visualization and image acquisition by CLSM

At each sampling point, two coupons were processed for visualization by CLSM.

From each coupon, at least three image stacks were acquired from random

positions using Olympus FV500 confocal scanning laser microscope (Olympus

Corporation, Japan).

Images were acquired at 1.0µm intervals down through the biofilm, thus the

number of images in each stack varied according to the thickness of the biofilm.

The 512 pixels × 512 pixels images were obtained with a PlanApo 60× /1.40 oil

immersion objective. Together, each pixel was considered as a box (voxel) with

the dimensions 0.41432µm3 (x-axis) × 0.41432µm3 (y-axis) × 1.000µm3 (z-axis).

Since each image had a coverage of 45,000µm2, thus a total of 3 images per

coupon reflected the coverage of >100,000µm2 per coupon, sufficient to obtain a

representative data of the Pseudomonas biofilm (Korber et al., 1993). For CFP, PI

and CFDA, image scanning was carried out using the 405nm laser line from a

LD405 laser, 543nm laser line from a HeNe-G laser and 488nm laser line from an

M-Ar laser respectively. To reduce background, either emission filter BA430-460,

BA-610IF and BA505-525 was used respectively. Variables that may influence

the quality of the images for each fluorescence, like the photomultiplier tube

(PMT), confocal aperture (CA) and laser power, were kept constant for all

experiments.



3.8.8 Application of COMSTAT image analysis software package

COMSTAT (Heydorn et al., 2000) was written as a script in MATLAB 5.1 (The

MathWorks Inc., Natick, Massachusetts), equipped with Image Processing

Toolbox. The COMSTAT package contained 5 programs (COMSTAT,

CHECKALL, LOOK, LOOKTIF and CONVERT000) and a number of functions

used by the programs.

3.8.8.1 Preparation of image stacks

In order to store image data in a format that can be analyzed by COMSTAT, the

images were prepared as follows:

• Images of a stack, of each fluorescence type, acquired by CLSM were

extracted and saved as individual ‘.tif’ files in the MATLABR11/work

folder using Olympus FLUOVIEW Ver.1.3 Viewer.

• An ‘.info’ file was created using Notepad by writing a text file with the

extension ‘.info’ and the ‘.info’ file contained vital information of the

image stack arranged in the following order:

Range #1# #13# Pixelsize (x) #0.41432# Pixelsize (y) #0.41432# Pixelsize (z) #1.000# ‘Range’ reflected the number of images in this stack. The ‘pixelsize’

represented the distance between 2 neighboring pixels and was given in



micrometer. The name of the ‘.info’ file was also changed according to the

name of the images in the stack.

• The original MATBLAB script of the CONVERT000 program was

improved to allow renaming and reversing the order of the images of a

stack in a way that COMSTAT can analyze. The improved program was

named “CONVERTWKL”.

• CHECKALL program was run to check that all the stacks of images were

intact.

3.8.8.2 Thresholding of images

After the image preparations, LOOK program was run to allow manual

determination of the threshold value for each stack of images of different

fluorescence type. LOOKTIF program allowed closer view of individual images.

Such thresholding of a stack of images in COMSTAT resulted in the formation of

a 3-dimensional matrix with a value of ONE in positions where pixel values in the

original image were above or equal to the threshold value (representing biomass

or data point of biofilm) and ZERO where the pixel values were below the

threshold value (representing background).

3.8.8.3 COMSTAT image analysis for P. aeruginosa PAO1-CFP biofilm

structure

To study P. aeruginosa PAO1-CFP biofilm structure, stacks of CFP images were

analyzed by COMSTAT program running on ‘connected volume filtered images’.

Connected volume filtration of the stacks of images removed noise from images



by removing biomass that is not in some way connected to the substratum. The

following COMSTAT image analysis features were run:

• Bio-volume (µm3/µm2) - volume of the biofilm normalized by the surface

area of the field of view.

• Average thickness (µm) – average biofilm height taken over the entire

field of view.

• Maximum thickness (µm) – maximum distance from the substratum that

the biofilm reaches.

• Surface to bio-volume ratio (SBR) (µm2/µm3) – the area summation of all

biomass voxel surfaces exposed to the background per unit bio-volume,

thus reflects the fraction of biofilm apparently exposed to nutrient flow.

• Substratum coverage (%) – the area coverage in the first image of the

stack, i.e. at the substratum, thus reflects how efficiently the substratum is

colonized by bacteria of the population.

• Roughness coefficient – a measure of variability in biofilm thickness and

consequently, an indicator of biofilm heterogeneity.

To obtain the distribution of bio-volume, the image analysis program COMSTAT

was improved to report the number of CFP pixels in each layer of the biofilm. The

improved program was named “COMSTATWCY” and run with connected

volume filtration. For each stacks of images, the number of CFP pixels belonging

to each sections of the biofilm were summed up. Bio-volume of the biofilm was

then calculated using the following formula:



Bio-volume

Bio-volume (µm3µm-2) = [(Number of CFP pixels × voxel size) / (area of the field

of view)]

Loss of bio-volume

Amount of bio-volume loss

= (Bio-volume before the addition of biocide – Bio-volume at 24hrs of exposure

to biocide)

To obtain the distribution of SBR, the image analysis program COMSTAT was

improved to report the SBR in each layer of the biofilm. The improved program

was named “COMSTATLAYER” and run with connected volume filtration.

Surface area of each layer of the biofilm was then obtained by dividing SBR with

corresponding bio-volume. For each stack of images, both the surface area and

bio-volume belonging to each sections of the biofilm were summed up separately

before calculating the sectional SBR by dividing sectional surface area with

corresponding sectional bio-volume.

3.8.8.4 COMSTAT image analysis for porosity of P. aeruginosa PAO1-CFP

biofilm

To study the porosity of the biofilms, COMSTATWCY was used to analyze the

stacks of PI and CFP images, and run without connected volume filtration. The

report of the analysis detailed the number of PI and CFP pixels in each images of

a stack, that is, in each layer of the biofilm. For each stack of images, the numbers



of PI and CFP pixels belonging to each sections of the biofilm were summed up.

Porosity of the biofilm was then calculated using the following formula:

Porosity

Porosity = [Number of PI pixels / (Number of CFP pixels × voxel size)] = Number

of PI pixels per µm3 of biomass

3.8.8.5 COMSTAT image analysis for L. pneumophila distribution

COMSTATWCY was also used to analyze the stacks of CFDA images, without

connected volume filtration. The report stated the number of CFDA pixels in each

layer of the biofilm. For each stack of images, the numbers of CFDA pixels

belonging to each sections of the biofilm were summed up while the CFP data was

the same as that in the above section. Subsequently, the following calculations

were performed:

L. pneumophila concentration

Log (Legionellae concentration)

= Log [Number of CFDA pixels / (Number of CFP pixels × voxel size)]

= Log (Number of CFDA pixels per µm3 of biomass)

L. pneumophila loss from P. aeruginosa PAO1-CFP biofilm

% Loss of legionellae

= {[(legionellae concentration at day of inoculation) – (legionellae concentration

at day n)] / (legionellae concentration at day of inoculation)} × 100%

where, n = number of days following legionellae inoculation



3.8.9 Statistical analysis

All statistical analyses in this study were carried out using SPSS 13.0.

Independent samples t-test was used to compare the means for 2 groups of cases.

If the significance value for the Levene test was high (typically greater than 0.05),

equal variances for both groups were assumed. A low significance value for the t-

test (typically less than 0.05) indicated significant difference between the 2 group

means. In addition, if the confidence interval for the mean difference did not

contain zero, this indicated that the difference was significant.

Pearson correlation was used to determine if there is a linear association between

variables on the assumption that the data are normally distributed. The values of

the correlation coefficient ranged from -1 to 1. The sign of the correlation

coefficient indicated the direction of the relationship while the absolute value of

the correlation coefficient indicated the strength, with larger absolute values

indicating stronger relationships. The significance level (or p-value) was the

probability of obtaining results as extreme as the one observed.

3.9 Screening for effective P. aeruginosa PAO1 biofilm-removing

agent

3.9.1 Kinetics of P. aeruginosa PAO1 biofilm formation in microtiter plate

An overnight culture of P. aeruginosa PAO1 (grown in MM at 37°C with shaking

at 120rpm) was inoculated into fresh MM medium at a ratio of 1:100. The freshly

inoculated medium was dispensed into each of the 8 wells (100µl per well) of the

first column of non-tissue culture treated, flat bottom, 96-well polystyrene plates



(BD FalconTM, U.S.A.). The inoculated 96-well plate was then incubated at 30°C

with shaking at 120rpm, while the remaining inoculated medium was stored at

4°C to prevent any growth. At every 2 hr interval, the inoculated medium was

mixed well and dispensed in the same way, into the subsequent column of 8 wells.

One column of every 96-well plate contained only MM (blank and negative

control). After 40 hrs from the first inoculation, the biofilm formed on the walls of

each wells were quantified as described below. Three independent experiments

were conducted for this study.

3.9.2 Quantification of biofilm (O’Toole et al., 1999)

After biofilm was formed in 96-well plates, optical density reflecting total

bacterial growth (OD600nm) of the 96-well plates were first taken using ELISA

Touch Screen plate reader (Tecan, Austria) operated on Magellan2 software. The

spent culture medium, together with unattached bacteria, were then carefully

removed from each wells and replaced with 100µl of 1% (w/v) crystal violet in

deionized water (dH2O). After 10 mins of incubation at room temperature, excess

crystal violet was washed away by rinsing the plate in several basins of dH2O.

These washed plates were then tapped to remove excess water and air-dried.

Biofilm-bound crystal violet (reflecting the amount of biofilm formed) was

solubilized by adding 100µl of 95% ethanol to each well, incubated at room

temperature with shaking at 100rpm for 10 mins and then quantified by measuring

OD470nm of the 96-well plates.



3.9.3 Biofilm-removing agents used

To obtain the most effective biofilm-removing agent, commercially available

products of a variety of nature were screened. All 8 biofilm-removing agents used

in this study were listed in table 3.2.

Table 3.2. All biofilm-removing agents used. * represents active ingredients of the various biofilm-removing agents.

Biofilm-removing

agents Ingredients Proportion

%(w/w) Applications

- Sodium Hypochlorite* 5.0-10.0 - Sodium Hydroxide* 1.0-5.0

ACTI-PLUS 2818

- Inorganic salt (s) and water To 100

Stabilized sodium hypochlorite microorganism control chemical

- Dimethyl-Dioctyl-Ammonium Chloride*

10.0-30.0

- Ethanol 1.0-5.0

NALCO 90001

- Water To 100

Algaecide

- 2,2-Dibromo-3-nitrilopropionamide (DBNPA)*

10.0-30.0

- Dibromoacetonitrile 0.1-1.0

NALCO 7320

- Glycol and water To 100

Microorganism control chemical

- 5-chloro-2-methyl-4-isothiazolin-3-one*

1.1

- 2-methyl-4-isothiazolin-3-one*

0.1-1.0

NALCO 7330

- Water To 100

Biocide

- Glutaraldehyde* 30.0-60.0 NALCO 7338 - Water To 100

Biocide

NALSPERSE ® 7348

- Polyglycol* 100 Biodispersant

- C10-16 Polyglycoside* 10.0-30.0 - C8-10 Polyglycoside* 10.0-30.0

NALCO 73550

- Water 100

Biodetergent

- Enzyme protein Protease Subtilisin*

proprietary COOLING TOWER QUARTERLY CLEANER

- Enzyme protein Metalloprotease*

proprietary

Concentrated enzymatic formulation for digestion of



- Enzyme protein Lipase triacylglycerol*

proprietary

- Enzyme protein Cellulase* proprietary - Enzyme protein Cellulase* proprietary - Enzyme protein Amylase* proprietary - Enzyme protein Alpha-Amylase*

proprietary

- 5-Chloro-2-methyl-4-isothiazolin-3-one*

<0.09

- 2-Methyl-4-isothiazolin-3-one*

<0.09

- Linear alkyl pyrrolidone <5 - Polyethylene oxide derivative of synthetic alcohols

<20

- Polyoxyethylene, polyoxypropylene, polyoxybuthylene ether of a mixture of aliphatic alcohols

<10

- Glycerine <10 - Borax <10

biological debris, biofilm, protozoa and reducing bacterial resistance to common biocides

3.9.4 P. aeruginosa PAO1 biofilm removal screening

P. aeruginosa PAO1 biofilm was formed in 96-well plates for 12 hrs at 30°C with

shaking at 120rpm before biofilm-removing agents, diluted with dH2O to varying

concentrations (in terms of parts per million, ppm), were added into each column

of 8 wells and their efficiency of biofilm removal were monitored over the

subsequent 8 hrs (with 2 hrs interval). The concentrations of biofilm-removing

agents used were 10, 50, 100, 500 and 1000ppm. One column of every 96-well

plate was mock-treated with dH2O (negative control) and similarly, one column of

every 96-well plate contained only MM (blank). Three independent experiments

were conducted for this study.



3.10 Antimicrobial susceptibility testing of NALCO 7320

Standard macrodilution method (Ferraro, 2003) was used to ascertain the

minimum inhibitory concentration (MIC) and minimum bactericidal concentration

(MBC) of NALCO 7320. P. aeruginosa PAO1 and L. pneumophila cells grown to

late-log phase were harvested at 28 and 24 hrs of growth respectively, and re-

suspended in PBS. Cell concentrations were adjusted to OD600nm = 0.5 and

enumerated by Miles and Misra method on appropriate agar. 3ml of culture media,

MM for P. aeruginosa PAO1 and 20% BCYE broth for L. pneumophila,

containing different concentrations of biofilm-removing agents (1,000ppm,

500ppm, 100ppm, 50ppm and 10ppm) were inoculated with 100µl of the cell

suspension and incubated at 30°C for 24 hrs. One tube was mock-treated with

dH2O (negative control) and similarly, one tube contained culture media only

(blank).

The MIC was recorded as the lowest concentration of biofilm-removing agent that

completely inhibited visible growth. The MBC was determined by spread plating

100µl from the tubes with no visible growth onto appropriate agar. Three

independent experiments were conducted for this study.

Results


Chapter 4: Results

4.1 Growth kinetics

Figure 4.1, 4.2 and 4.3 illustrated the growth kinetics and time taken to reach late

log phase of L. pneumophila ATCC 33152, P. aeruginosa PAO1 and P.

aeruginosa PAO1-CFP respectively. All Legionella, P. aeruginosa PAO1 and P.

aeruginosa PAO1-CFP cells used in the subsequent experiments were harvested at

late log phase, unless otherwise stated.

Growth Curve of L. pneumophila ATCC 33152

0.0

1.0

2.0

3.0

4.0

5.0

6.0

0 5 10 15 20 25 30 35

Time (hr)

OD

(600

nm)

Late Log Phase (24hr)

Figure 4.1. Growth curve of L. pneumophila cultured in BCYE broth at 37°C with shaking at 120rpm. The error bars represent standard deviation of 3 independent experiments.

Results


Growth Curve of Pseudomonas aeruginosa PAO1

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

1.8

2.0

0 5 10 15 20 25 30 35

Time (hr)

OD

(600

nm)


Figure 4.2. Growth curve of P. aeruginosa PAO1 cultured in MM liquid media at 30°C with shaking at 120rpm. The error bars represent standard deviation of 3 independent experiments.

Growth Curve of Pseudomonas aeruginosa PAO1-CFP

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

1.8

2.0

0 5 10 15 20 25 30 35

Time (hr)

OD

(600

nm)


Exponential Phase

Figure 4.3. Growth curve of P. aeruginosa PAO1-CFP cultured in MM liquid media at 30°C with shaking at 120rpm. The error bars represent standard deviation of 3 independent experiments.

Results


4.2 Determination of the influent flow rate (Q) for continuous

culture in CDC Biofilm Reactor (CBR)

Exponential growth phase of Pseudomonas aeruginosa PAO1-CFP

y = 0.2505x - 6.5649R2 = 0.9951

-2.0

-1.5

-1.0

-0.5

0.0

0.5

1.0

18 20 22 24 26 28 30

Time (hr)

Ln [O

D (6

00nm

)]

Figure 4.4. Graph of Ln(OD600nm) against time (hr) was plotted for the exponential growth phase of P. aeruginosa PAO1-CFP, so as to obtain the maximum specific growth rate, which was reflected by the gradient of the best straight line plotted. The error bars represent standard deviation of 3 independent experiments.

From figure 4.4, the maximum specific growth rate (µmax) = 0.2505 hr-1

= 4.18 × 10-3 min-1.

Doubling time (td) = ln2 / µmax = ln2 / (4.18 × 10-3) = 165.8 mins = 2.76 hr.

In order to select for biofilm growth in the CBR, the hydraulic residence time (θ)

must be less than the doubling time for the suspended cells. This will result in the

suspended cells washing out of the reactor, leaving only biofilm.

Results


To determine the nutrient influent flow rate (Q) such that θ < td, the following

calculations were performed:

Since θ = V/Q,

V/Q < 165.8

Q > V/165.8

Q > 400 / 165.8

∴Q > 2.41 ml/min

Where, V is maximum volume of bulk fluid in CBR during continuous flow (with

all coupons and coupon holders removed) = 400ml.

In conclusion, continuous culture were conducted with Q = 2.5ml/min.

As such, highest possible θ = V/Q = 400 / 2.5 = 160mins = 2.6hr.

Results


4.3 Optimization of labelling processes

4.3.1 Optimization of L. pneumophila labelling with CFDA-SE

Together, figure 4.5 and table 4.1, show that 30mins was the longest treatment

duration (with 10µM CFDA-SE) that resulted in more than 95% of Legionella

cells being labeled with CFDA without compromising viability.

(A) (B) (C) (D)

FL1-H FL1-H FL1-H FL1-H

Figure 4.5. Histograms illustrating the number of events (cells) plotted against FL1-H (representing green fluorescence of CFDA-stained cells) for L. pneumophila cells that were (A) mock treated, or treated with CFDA-SE for (B) 20mins, (C) 30mins, or (D) 40mins. Table 4.1. Effect of treatment duration on staining and viability of L. pneumophila cells. * represents % Cells stained represented the area of graph (see above) under Marker 1 (M1). Duration of treatment with CFDA-SE Mock-treated 20mins 30mins 40mins % Cells stained*

1.90% 97.0% 97.0% 97.5%

% Viable cells after staining process

- 100% 100% 86.8%

Results


4.3.2 Optimization of planktonic P. aeruginosa PAO1-CFP labelling with PI

Figure 4.6 and table 4.2 below, show that within 5mins of 1.0mg/ml PI treatment,

majority (up to 88.1%) of formaldehyde fixed P. aeruginosa PAO1-CFP cells

picked up PI stain. Unsurprisingly, fewer cells attained the comparable level of

fluorescence for the same treatment duration when 0.1mg/ml PI was used instead

(figure 4.7 and table 4.2). In addition, even after 30mins of treatment with

0.1mg/ml PI, only 64.2% of the cells attained comparable high level of

fluorescence. These imply that the PI concentration of 0.1mg/ml was limited for

substantial staining of a 107 CFU/ml P. aeruginosa PAO1-CFP cell suspension.

igure 4.6. Histograms illustrating the number of events (cells) plotted against

(A) (B) (D) (C)

FPMT4 Log (representing red fluorescence of PI-stained cells) for P. aeruginosa PAO1-CFP cells that were (A) mock treated, or treated with 1.0mg/ml PI for (B) 5mins, (C) 10mins, or (D) 15mins.

Results


(A) (B) (C)

(D) (E)

Figure 4.7. Histograms illustrating the number of events (cells) plotted against PMT4 Log (representing red fluorescence of PI-stained cells) for P. aeruginosa PAO1-CFP cells that were (A) mock treated, or treated with 0.1mg/ml PI for (B) 5mins, (C) 10mins, (D) 15mins, or (E) 30mins. Table 4.2. Effect of treatment duration on staining of P. aeruginosa PAO1-CFP cells. * represents % Cells stained represented the area of graph (see above) under Marker 1 (M1).

Duration of treatment in 1.0mg/ml PI

Duration of treatment in 0.1mg/ml PI

Negative control

5mins 10mins 15mins 5mins 10mins 15mins 30mins

% Cells stained*

9.91% 88.7% 91.8% 92.8% 55.0% 53.3% 60.2% 64.2%

Results


4.3.3 Optimization of P. aeruginosa PAO1-CFP biofilm labelling with PI

When a low concentration of 0.1mg/ml PI was applied to formaldehyde-fixed 7

days old P. aeruginosa PAO1-CFP biofilm with L. pneumophila for merely

5mins, regions of the biofilm with the greatest access to the external dye was

observed to be stained with a higher intensity of redness (figure 4.8 (C)). Since

freshly introduced L. pneumophila co-localized with these regions as seen in

figure 4.8 (D), this further affirms the proposition that the more PI pixels per unit

biomass, the higher the porosity of the biofilm.

(A) (B)

(C) (D)

Figure 4.8. CLSM images of a 7 day old P. aeruginosa PAO1-CFP biofilm and adhered L. pneumophila, stained with 0.1mg/ml PI for 5mins: (A) P. aeruginosa PAO1-CFP biofilm (blue fluorescence), (B) CFDA-stained L. pneumophila (green fluorescence), (C) PI-stained P. aeruginosa PAO1-CFP biofilm, and (D) overlapping display of the above 3 images. The scale represents 30µm in each image. Arrow indicates the porous flow channel within cell clusters of biofilm.

Results


However, when the PI treatment duration was increased to 15mins (figure 4.9) and

30mins (figure 4.10), the biofilms were over-stained and regions of higher

porosity were not discernible. Henceforth, a concentration of 0.1mg/ml PI and

treatment duration of 5mins were applied to all biofilm samples intended for

CLSM examination.

(C) (D)

(A) (B)

Figure 4.9. CLSM images of a 7 day old P. aeruginosa PAO1-CFP biofilm and adhered L. pneumophila, stained with 0.1mg/ml PI for 15mins: (A) P. aeruginosa PAO1-CFP biofilm (blue fluorescence), (B) CFDA-stained L. pneumophila (green fluorescence), (C) PI-stained P. aeruginosa PAO1-CFP biofilm, and (D) overlapping display of the above 3 images. The scale represents 30µm in each image.

Results


(C) (D)

(A) (B)

Figure 4.10. CLSM images of a 7 day old P. aeruginosa PAO1-CFP biofilm and adhered L. pneumophila, stained with 0.1mg/ml PI for 30mins: (A) P. aeruginosa PAO1-CFP biofilm (blue fluorescence), (B) CFDA-stained L. pneumophila (green fluorescence), (C) PI-stained P. aeruginosa PAO1-CFP biofilm, and (D) overlapping display of the above 3 images. The scale represents 30µm in each image.

Results


4.4 Kinetics of P. aeruginosa PAO1-CFP biofilm formation in

CDC Biofilm Reactor (CBR)

4.4.1 Kinetics of biofilm formation

Figure 4.11 illustrated the steady increase in the average number of viable P.

aeruginosa PAO1-CFP cells in biofilm until it became relatively levelled off (at

7.04 ×104 CFU/mm2) after 6 days of growth in the continuous flow CBR system.

This developmental plateau was observed in at least 3 independent experiments,

thus was reproducible in this system. Henceforth, mature biofilm was demarcated

by the 6th day of development in this system, while developing biofilm

corresponded to the days before the plateau was reached. Even so, there was a

slight but insignificant increase (independent sample t-test, p > 0.1; assuming

equal variance) in the average viable cell counts on Day 10 and 11 of biofilm

Figure 4.11. Viable cell counts of P. aeruginosa PAO1-CFP biofilm formed in

development to 1.18 × 105 CFU/mm2 and 1.64 × 105 CFU/mm2 respectively.

CBR at 30°C with stirring at 120rpm. The error bars represent standard deviation of at least 3 independent experiments.

Pseudomonas aeruginosa PAO1-CFP counts in biofilm

0.0

1.0

2.0

3.0

4.0

5.0

6.0

2 3 4 5 6 7 8 9 10 11Days

Log

PAO

1-C

FP c

once

ntra

tion

[Log

(C

FU/m

m2 )]

Mature biofilmDeveloping biofilm

Results


4.4.2 Structure of biofilm by image analysis

Image analysis revealed that the profile of biofilm bio-volume (figure 4.12),

average thickness (figure 4.13) and maximum thickness (figure 4.14)

corresponded with that of P. aeruginosa PAO1-CFP viable counts up to day 9.

Despite the relatively constant maximum thickness, there was a noticeable but not

significant increase in bio-volume and average thickness on day 10 and 11. Figure

4.15 illustrated yet another perspective of the biofilm where average substratum

coverage peaked on day 3 at 74.5% ± 10.2% and dropped drastically to reach a

low of 13.1% ± 8.88% on day 6. On subsequent days, the average substratum

coverage remained low between 20.0%-30.0% as compared to >50.0% for early

developing biofilm.

urface-to-biovolume ratio (SBR) remained comparable (between 0.400 – 0.650

µm2/µm3) throughout biofilm development, as shown in figure 4.16. However,

figure 4.17 demonstrated that the roughness coefficient started off high at 0.483 ±

0.145 and decreased gradually to 0.319 ± 0.220 on day 8, before dropping

noticeably to between 0.110 – 0.130 on day 9 onwards.

S

Results


Bio-volume

0.0

5.0

10.0

15.0

20.0

25.0

30.0

35.0

40.0

2 3 4 5 6 7 8 9 10 11Days

Bio

-vol

ume

(µm

3 /µm

2 )Developing biofilm Mature biofilm

Figure 4.12. Bio-volume of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. The bio-volume of each experiment was obtained from 3-8 image stacks from 1 or 2 coupons. The error bars represent standard deviation of at least 3 independent experiments.

Average thickness

0.0

5.0

10.0

15.0

20.0

25.0

30.0

35.0

40.0

45.0

2 3 4 5 6 7 8 9 10 11Days

Ave

rage

thic

knes

s (µ

m)

Developing biofilm Mature biofilm

Figure 4.13. Average thickness of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. The average thickness of each experiment was obtained from 3-8 image stacks from 1 or 2 coupons. The error bars represent standard deviation of at least 3 independent experiments.

Results


Maximum thickness

0.0

10.0

20.0

30.0

40.0

50.0

60.0

2 3 4 5 6 7 8 9 10 11Days

Max

imum

thic

knes

s (µ

m)


Figure 4.14. Maximum thickness of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. The maximum thickness of each experiment was obtained from 3-8 image stacks from 1 or 2 coupons. The error bars represent standard deviation of at least 3 independent experiments.

Substratum coverage

0%

10%

20%

30%

40%

50%

60%

70%

80%

90%

100%

2 3 4 5 6 7 8 9 10 11Days

Subs

trat

um c

over

age

(%)


Figure 4.15. Substratum coverage of P. aeruginosa PAO1-CFP biofilm formed in CBR at 30°C with stirring at 120rpm. The substratum coverage of each experiment was obtained from 3-8 image stacks from 1 or 2 coupons. The error bars represent standard deviation of at least 3 independent experiments.

Results


Surface-to-biovolume ratio0.90

0.00

0.10

0.20

0.30

0.40

0.50

0.60

0.70

0.80

2 3 4 5 6 7 8 9 10 11Days

Surf

ace-

to-b

iovo

lum

e ra

tio (µ

m2 /µ

m3 )


Figure 4.16. Surface-to-biovolume ratio (SBR) of P. aeruginosa PAO1-CFP iofilm formed in CBR at 30°C with stirring at 120rpm. The SBR of each

CBR at 30°C with stirring at 120rpm. The roughness coefficient of each

bexperiment was obtained from 3-8 image stacks from 1 or 2 coupons. The error bars represent standard deviation of at least 3 independent experiments.

Roughness coefficient

0.000

0.100

0.200

0.300

0.400

0.500

0.600

0.700

2 3 4 5 6 7 8 9 10 11Days

Rou

ghne

ss c

oeffi

cien

t


Figure 4.17. Roughness coefficient of P. aeruginosa PAO1-CFP biofilm formed inexperiment was obtained from 3-8 image stacks from 1 or 2 coupons. The error bars represent standard deviation of at least 3 independent experiments

Results


4.4.3 Detachment of biofilm

Figure 4.18 illustrated a gradual increase in the average number of P. aeruginosa

om a low of 1.01 × 105 CFU/ml on day 2 to 2.26 ×

andard deviation of at least 3 independent experiments .

PAO1-CFP in the bulk fluid fr

105 CFU/ml on day 5, followed by a significant increase (independent sample t-

test, p = 0.006, assuming equal variance) to 6.58 × 105 CFU/ml on day 6. After

which, the average planktonic viable count remained relatively constant but

increased slightly from day 10 onwards to a high of 1.65 × 106 CFU/ml on day 11.

Such viable counts of P. aeruginosa PAO1-CFP in the bulk fluid reflected

instantaneous detachment of biofilm because the high nutrient influent flow rate

applied to the CBR would have resulted in the washout of planktonic P.

aeruginosa PAO1-CFP before these cells could replicate (Chapter 4.2).

Planktonic P. aeruginosa PAO1-CFP counts

0.0

1.0

2.0

3.0

4.0

5.0

6.0

7.0

2 3 4 5 6 7 8 9 10 11Days

Log

PAO

1-C

FP c

once

ntra

tion

[Log

(C

FU/m

l)]


Figure 4.18. Viable cell counts of planktonic P. aeruginosa PAO1-CFP in the bulk fluid of CBR at 30°C with stirring at 120rpm. The error bars represent st

Results


Biofilm structures indicative of the final stage in Pseudomonas biofilm

development, namely the dispersion stage (Tolker-Nielsen et al., 2000; Sauer et

(blue) structure L.

neumophila (green). Biofilm stained with PI (red) reflected porous regions. The -y view of the biofilm (main view) is flanked by y-z (right) and x-z (bottom)

al., 2002), were observed occasionally and earliest seen on day 7 of P. aeruginosa

PAO1-CFP biofilm development (Figure 4.19). The “wall” of P. aeruginosa

PAO1-CFP cells that encompassed the void space containing sparse amount of P.

aeruginosa PAO1-CFP cells were low in porosity, thereby preventing PI staining

of the P. aeruginosa PAO1-CFP cells within the void. It is also worth noting that

no legionellae has been found within such voids.

Void

Figure 4.19. CLSM image of a P. aeruginosa PAO1-CFP biofilmindicative of dispersion stage of biofilm development, with adhered pxsections of the biofilm, with red arrows pointed towards the top of biofilm. The scale represents 30µm in each image.

Results


4.5 Introduction of L. pneumophila to developing and mature P.

eruginosa PAO1-CFP biofilmsa

4.5.1 Adhesion and persistence of L. pneumophila in developing and mature

biofilms

per coupon per 106 inoculated legionellae and 38.7 ± 26.4 cells per

The amount of legionellae adhering to developing and mature biofilm were 10.8 ±

9.0 cells

coupon per 106 inoculated legionellae respectively, as shown in figure 4.20. Using

SPSS, independent samples t-test was calculated. It was found that p = 0.056

(assuming equal variances), thus we cannot conclude that there was significant

difference between the adhesion of L. pneumophila to each coupon of developing

and mature biofilms.

Adhesion of L. pneumophila to P. aeruginosa PAO1-CFP biofilm

0.000

10.000

20.000

30.000

40.000

50.000

60.000

Developing Mature

Num

ber o

f leg

ione

llae

per c

oupo

n pe

r 106

legi

onel

lae

inoc

ulat

ed in

to C

BR

70.000

Figure 4.20. Adhesion of L. pneumophila to different developmental stages of P. aeruginosa PAO1-CFP biofilm. The error bars represent standard deviation of 5 independent experiments.

Results


With regards to the persistence of L. pneumophila in P. aeruginosa PAO1-CFP

biofilm, figure 4.21 illustrated an approximately 1 log decrease in legionellae

counts over 5 and 4 days following its introduction to developing and mature

biofilm respectively. Interestingly, in developing biofilms, L. pneumophila cell

counts only started to decrease noticeably on day 6 of the biofilm development,

which corresponds to the maturation of the P. aeruginosa PAO1-CFP biofilm.

Figure 4.21 also showed that the average planktonic L. pneumophila cell counts

remained high at >1.00 × 10 cells/ml, even after 3 hrs of allowance for washout

following legionellae introduction into both types of biofilms, revealing the

instability of initial legionellae adhesion to the biofilms. Nevertheless, on

subsequent days after L. pneumophila introduction into both types of biofilm,

planktonic legionellae dropped drastically and remained low at <100 cells/ml.

When the persistence of L. pneumophila in P. aeruginosa PAO1-CFP biofilms

was examined more closely in figure 4.22, only 35.2% ± 16.8% of legionellae

remained in mature biofilm 1 day after its introduction into CBR while 98.8% ±

1.0% of legionellae remained in developing biofilm. It was 3 days after

legionellae introduction, which corresponded to the maturation of the developing

biofilm, when the biofilm legionellae started to decrease, leaving 47.9% ± 9.8% in

the biofilm. The release of legionellae from matured biofilms was fastest initially

and slowed down within the next 3 days. Finally, legionellae loss from mature

biofilm tended to stabilize with slightly >10% of legionellae remaining in the

mature biofilm 4 days after the introduction of exogenous legionellae into CBR.

4

Results


Status of L. pneumophila in continuous flow CBR system

0.0

1.0

2.0

3.0

4.0

5.0

6.0

1 2 3 4 5 6 7 8 9 10 11 12

Days

Log

Legi

onel

la c

once

ntra

tion

in

plan

kton

ic p

hase

[Log

(c

ells

/ml)]

0.0

1.0

2.0

3.0

4.0

5.0

6.0Log Legionella concentration in

biofilm [Log (cells/m

m2)]

Figure 4.21. Status of L. pneumophila in our continuous flow CBR system. The error bars represent standard deviation of 3 independent experiments.

Figure 4.22. Persistence of L. pneumophila in P. aeruginosa PAO1-CFP biofilm. The error bars represent standard deviation of 3 independent experiments.

Developing biofilm - planktonic Mature biofilm - planktonicDeveloping biofilm - biofilm Mature biofilm - biofilm

Legionella addedLegionella added

Persistence of L. pneumophila in P. aeruginosa PAO1-CFP biofilm

98.8100.0

0 1 2 3 4 5 6Days following inoculation of Legionella

% L

egio

nella

rem

aini

ng in

bio

film

18.7

29.2

47.9

91.9

35.2

20.715.1 14.2

0.0

20.0

40.0

60.0

80.0

100.0

120.0Developing biofilmMature biofilm

Results


4.5.2 Distributions of L. pneumophila cells in developing and mature biofilms

Table 4.3 demonstrated that there was a significant positive linear correlation (at

0.511) between Log (Number of Legionella cells) and Log (Number of CFDA

pixels per µm3 of biofilm) from 6 independent experiments. Henceforth, Log

(Number of CDFA pixels per µm3 of biofilm) can well represent the amount of

CFDA-labelled legionellae in the biofilm.

To study the distribution of L. pneumophila cells in P. aeruginosa PAO1-CFP

biofilm, the latter was divided into 5 sections along its height (z-axis). The x<20%

represented the bottom most section of the biofilm while x≥80% represented the

top most section.

Adhesion distribution of L. pneumophila in both developing and mature biofilm

appeared similar in figure 4.23(A) and (B) respectively, where most legionellae

adhered to the top of the biofilms. Referring to figure 4.23(A), there was a shift in

the peak of legionellae unimodal distribution from 40%-60% to 20%-40% and

finally bottom 20% of biofilm from day 1 to day 3. On day 4, legionellae

distribution became bimodal. Here, least amount of legionellae was found in the

middle section (40%-60%) of the biofilm (where Log (Number of CFDA pixels

per µm3) was -3.84µm-3), sandwiched between two peaks at 20%-40% and 60%-

80% of the biofilm (-3.40µm-3 and -3.41µm-3 respectively). Finally, on day 5,

distribution of legionellae remained bimodal, but with most of them found at the

bottom 20% of biofilm where Log (Number of CFDA pixels per µm3) was -

3.71µm-3, followed by 60%-80% of the biofilm where Log (Number of CFDA

Results


pixels per µm3) was -3.78µm-3. On the other hand, least legionellae were found

distributions (refer to figure 4.23(B)).

he distribution changed to and fro from a unimodal distribution on day 1 (peak

odal distribution on day 4 (peaks of -2.52µm-3 and -2.43µm-3 at

ottom 20% and 60%-80% of biofilm respectively).

both at 40%-60% and top 20% of the biofilm where Log (Number of CFDA pixels

per µm3) were both -3.97µm-3.

In contrast, legionellae distribution in mature biofilm from day 1 to day 3

fluctuated between unimodal and bimodal

T

of -1.88µm-3 at 40%-60% of biofilm) to a bimodal distribution on day 2 (peaks of

-2.22µm-3 and -2.21µm-3 at bottom 20% and 60%-80% of biofilm respectively), to

unimodal distribution on day 3 (peak of -2.40µm-3 at bottom 20% of biofilm) and

finally back to bim

b

The losses of legionellae from different regions of developing and mature P.

aeruginosa PAO1-CFP biofilms were illustrated in figure 4.24 and 4.25

respectively. Generally, highest legionellae losses were located at the top 40% of

both types of biofilm while lowest legionellae losses were found at bottom 60%,

especially at bottom 20%. Legionellae was lost faster from bottom 60% of

developing biofilm than that of mature biofilm, since 2-3 days were enough for

legionellae loss at bottom 60% of developing biofilm to reach >90% while 4 days

were required in mature biofilm. On the contrary, the rate of legionellae loss at top

40% of both developing and mature biofilms were comparable.

Results


Table 4.3. Table showing Pearson’s correlation between Log (Number of L. 3

CFDA pixels per µmpneumophila cells) and Log (Number of CFDA pixels per µm ). Each number of

each experiment. The correlation between the 2 variables was obtained from all

3 was obtained from 3-8 image stacks from 1 or 2 coupons of

data of 6 independent experiments.

Correlations

1 .511**. .002

33 33.511** 1.002 .

33 33

Pearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)N

Log (Number of CFDApixels per um3 of biofilm)

Log (Number oflegionellae)

Log

CFDA pixelsper um3 of

(Number of

biofilm)Log (Numberof legionellae)

Correlation is significant at the 0.01 level (2-tailed).**.

Results


Dis

trib

utio

n of

Leg

ione

lla in

de

velo

ping

bio

film

-4.5

-4.0

-3.5

-3.0

-2.5

-2.0

-1.5

-1.0

-0.50.00

x<20

% 000000000 (botto

m)

Sect

ions

of b

iofil

m

Log (Number of CFDA pixels per µm3) D

ay 0

Day

1D

ay 2

Day

3D

ay 4

Day

5

Dis

trib

utio

n of

-4.5

0

-4.0

0

-3.5

0

-3.0

0

-2.5

0

-2.0

0

-1.5

0

-1.0

0

-0.5

0

0.00

x<20

% (bott

om)

Legi

onel

la in

mat

ure

biof

ilm

Sect

i

Log (Number of CFDA pixels per µm3)

ons

of b

iofil

mD

ay 0

Day

1D

ay 2

Day

3D

ay 4

Figu

re 4

.23.

Dis

tribu

tion

in (

A)

deve

lopi

ng,

and

(B)

P.

aeru

gino

sa

PAO

1-C

FP b

iofil

ms.

Day

0

deno

ted

the

day

of

of

in

to

tive

biof

ilms.

The

num

ber

of C

FDA

pix

els

of

ea

ch

rs

repr

esen

t rr

or

of

3

of L

. pne

umop

hila

mat

ure

intro

duct

ion

le

gion

ella

ere

spec

per

µm3

expe

rimen

t w

as o

btai

ned

from

3-

8 im

age

stac

ks

from

1 o

r 2 c

oupo

ns. T

he

erro

r ba

stan

dard

e

inde

pend

ent e

xper

imen

ts.

B

)(

A

)(

Results


Figure 4.24. Percentage loss of L. pneumophila in developing P. aeruginosa

AO1-CFP biofilm. The error bars represent standard deviation of 3 independent ents. x represents the spacial location within biofilm.

igure 4.25. Percentage loss of L. pneumophila in mature P. aeruginosa PAO1-F iofilm. The error bars represent standard deviation of 3 independent

experiments. x represents the spacial location within biofilm.

Pexperim

FC P b

Loss of L. pneumophila from develo

Day 1

Day 2

Day 3

ping biofilm

50% 60% 70% 80% 90% 100% 110%

Day 4

Day 5

Day

s af

ter l

egio

nella

e ad

ditio

n


x≥80% (top)60%≤x<80%40%≤x<60%20%≤x<40%x<20% (bottom)

Loss of L. pneumophila from mature biofilm

50% 60% 70% 80% 90% 100% 110%

Day 1

Day 2

Day 3

Day 4

Day

s af

ter l

egio

nella

e ad

ditio

n


x≥80% (top)60%≤x<80%40%≤x<60%20%≤x<40%x<20% (bottom)

Results


4.5.3 Bio-volume distributions of developing and mature biofilms

Bio-volume distributions of both biofilm types, to which L. pneumophila was

introduced, were presented in figure 4.26(A) and (B) respectively. On day 0 and 1

(corresponding to day 3 and 4 of biofilm development in continuous flow system),

the peak of bio-volume distribution in developing biofilm was found at 20%-40%

region of the biofilm while least bio-volume was found at top 20% of the biofilm

(Figure 4.26(A)). After which, the peak was shifted to 40%-60% region of the

biofilm and the amount of bio-volume found at bottom 20% of the biofilm

decreased appreciably from within the range of 2.50µm3µm-2 - 3.00µm3µm-2 and

remained low within the range of 1.50µm3µm-2 - 2.50µm3µm-2.

Figure 4.26(B) illustrated another peak shift from 40%-60% to 60%-80% region

of the biofilm on day 2 (corresponding to day 9 of biofilm development in

continuous flow system). Generally, majority of the cell mass of mature biofilm

resided within 40%-80% region while least fraction of the cell mass was found at

the bottom 20% of mature biofilm.

Results


Bio

-vol

ume

dist

ribut

ion

in

deve

lopi

ng b

iofil

m

0.00

1.00

2.00

3.00

4.00

5.00

6.00

7.00

8.00

9.00

x<20

% (bott

om)

Sect

ions

of b

iofil

m

Bio-volume (µm3µm

-2) D

ay 0

Day

1D

ay 2

Day

3D

ay 4

Day

5

Bio

-vol

u

0.00

1.00

2.00

3.00

4.00

5.00

6.00

7.00

8.00

9.00 20

% (tto

m)

me

dibu

ia

bi

m

x<

bo

stri

tion

n m

ture

ofil

Sect

ions

of b

iofil

m

Bio-volume (µm3µm

-2)

Day

0D

ay 1

Day

2D

ay 3

Day

4

Figu

re 4

.26.

Bio

-vol

ume

dist

ribut

ion

of

(A)

deve

lopi

ng,

and

(B)

mat

ure

P.

aeru

gino

sa

PAO

1-C

FP b

iofil

ms.

Day

0

deno

ted

the

day

of

intro

duct

ion

ofle

gion

ella

resp

ectiv

e bi

ofilm

s. T

bio-

volu

me

of

each

ex

perim

ent

was

obt

aine

d fr

om

3-8

imag

e st

acks

fr

om 1

or 2

cou

pons

. The

stan

dard

er

ror

of

3 in

depe

nden

t exp

erim

ents

. e

erro

r ba

rs

repr

es

in

to

he

ent

(A)

(B)

Results


4.5.4 f developing and mature

iofi

urfa th biofilm types, to which L.

neu was introduced, were presented in figures 4.27(A) and (B)

spe troduction to biofilm), SBR

BR of 0.839µm2µm-3 at top 20% sector, followed by 0.583µm2µm-3 at bottom

0% sector and lowest SBR of 0.257µm2µm-3 at 40%-60% region of the biofilm.

contrast, SBR distribution in mature biofilm on day 0 tended to be a sharper

V” shape with highest SBR of 1.87µm2µm-3 at bottom 20% sector, followed by

.04µm2µm-3 at top 20% sector and lowest SBR of 0.321µm2µm-3 at 40%-60%

gion of the biofilm.

efer to figure 4.27(A), SBR at the top and bottom 20% of developing biofilm

creased at different rates as days pass. At the bottom, SBR increased drastically

iofilm development in continuous flow system) and then remained high within

e range of 1.50µm2µm-3-2.00µm2µm-3. In comparison, SBR at the top remained

latively comparable within the range of 0.800µm2µm-3– 1.20µm2µm-3. The

west SBR also remained comparable within the range of 0.200µm2µm-3-

.400µm2µm-3 at 40%-80% region of the biofilm (more prone towards the 40%-

0% region). As such, the “U” shape of SBR distribution in early developing

iofilm became a more distinct “V” shape on day 2 and remained so afterwards.

Surface-to-biovolume ratio distributions o

lms

ce-to-biovolume ratio (SBR) distributions of bo

mophila

ctively. On day 0 (the day of legionellae in

b

S

p

re

distribution in developing biofilm tended to be a wide “U” shape with highest

S

2

In

“

1

re

R ring

in

from 0.770µm2µm-3 on day 1 to 1.68µm2µm-3 on day 2 (corresponding to day 5 of

b

th

re

lo

0

6

b

Results


Referring to figure 4.27(B), SBR at the bottom 20% of mature biofilm remained

high within the range of 1.50µm2µm-3-2.00µm2µm-3 while that at the top 20%

e range of 2.00-2.50 on day 2 onwards in the developing

iofilm (corresponding to day 5 of biofilm development in continuous flow

decreased from >0.900µm2µm-3 on day 0 and 1, to within the range of

0.550µm2µm-3-0.600µm2µm-3 on day 2 (corresponding to day 9 of biofilm

development in continuous flow system) onwards. Lowest SBR remained

relatively stable within the range of 0.100µm2µm-3-0.400µm2µm-3 but was shifted

from 40%-60% to 60%-80% region of the mature biofilm on day 1.

The change in SBR ratio (SBR at bottom 20%: top 20% of biofilm) was shown in

table 4.4. At day 0, the SBR ratio was 0.795 ± 0.407 and 2.31 ± 1.60 in

developing and mature biofilm respectively. With time, the SBR ratio increased

and stabilized within th

b

system). However in mature biofilm, the SBR ratio increased further to >3.00 on

day 2 onwards (corresponding to day 9 of biofilm development in continuous flow

system).

Results


Figu

re 4

.27.

Sur

face

-to-

R)

dist

ribut

ion

of

(A)

deve

lopi

ng,

and

(B)

mat

ure

P.

aeru

gino

sa

PAO

1-C

FP b

iofil

ms.

Day

0

deno

ted

the

day

of

intro

duct

ion

ofle

gion

ella

in

to

resp

ectiv

e bi

ofilm

s. Th

e SB

Rof

eh

exer

imen

t w

as

obta

ined

fr

om

3-8

imag

e st

acks

fro

m 1

or

2 he

err

or b

ars

repr

esen

t st

rror

of

3

inde

pend

ent

expe

rimen

ts.

biov

olum

e ra

tio

(SB

e

ac

p

coup

ons.

Tan

dard

e

Surf

ace-

to-b

iovo

lum

e ra

tio

dist

ribut

ion

in d

evel

opin

g bi

ofilm

0.0

0.5

1.0

1.5

2.0

2.5

x<20

% (bott

om)

Sect

ions

of b

iofil

m

Surface-to-biovolume ratio (µm2µm

-3) D

ay 0

Day

1D

ay 2

Day

3D

ay 4

Day

5

S-to

-bvo

lum

e ra

tiodi

bu i

ae

biof

ilur

face

iost

ritio

nn

mtu

r

0.0

0.5

1.0

1.5

2.0

2.5

x<20

% (bott

om)

m

Seon

silm

cti

of b

iof

Surface-to-biovolume ratio (µm2µm

-3)

Day

0D

ay 1

Day

2D

aD

ay 4

y 3

(B

)

A)

(

Results


Tab sus the top 20% of developing and obtained from 3 independent xp

top 20% of biofilm)

le 4.4. The ratio of SBR at the bottom 20% ver mature biofilm. The standard deviations wereeriments.

SBR ratio (SBR at bottom 20% : SBR at

e

Days Developing biofilm Mature biofilm 0 0.795 ± 0.407 2.315 ± 1.604 1 1.070 ± 0.723 2.396 ± 1.026 2 2.195 ± 0.541 3.007 ± 0.789 3 2.437 ± 1.113 4.322 ± 3.067 4 2.087 ± 1.214 3.481 ± 2.327 5 2.230 ± 1.870 -

Figure 4.28 illustrated that as the P. aeruginosa PAO1-CFP biofilm developed,

s decreased steadily, reaching 5.68 ± 1.25µm-3 on

p < 0.05; Table 4.5) to

-3 and remained low, within the range of 2.50 – 3.50µm-3 till day

1

ted that the distribution profile of biofilm porosity

ained similar throughout the experimental period, with the highest porosity

e biofilm and least porosity at bottom 40% region.

4.5.5 Porosity distributions of developing and mature biofilms

the overall porosity of biofilm

day 7. By day 8, the overall porosity dropped significantly (

3.44 ± 1.05µm

1 .

In addition, figure 4.29 illustra

rem

located at top 40% of th

Results


Porosity

2.0

4.

6.

8.

10.0

12.0

3 4 5 6 7 8 9 10 11

Num

ber o

f PI p

ixel

s pe

r µm

3 )

experiment was obtained from 3-8 image stacks from 1 or 2 coupons. The error

Table 4.5. Comparing means of porosity over time, using one-way ANOVA.

Figure 4.28. Porosity of P. aeruginosa PAO1-CFP biofilm. The porosity of each

bars represent standard deviation of at least 3 independent experiments.

0.0

0

0

0

Days

Poro

sity

(

Developing biofilm re biofilmMatu

Multiple Comparisons

Dependent Variable: PorosityScheffe(I) Day: 3

1.509280 1.210937 .989 -3.74689 6.765452.041144 1.210937 .934 -3.21503 7.297311.849624 1.210937 .962 -3.40655 7.105792.882132 1.048702 .500 -1.66984 7.434115.124096* 1.048702 .018 .57212 9.676075.669294* 1.210937 .027 .41312 10.925465.901725* 1.210937 .018 .64556 11.157895.609074* 1.210937 .029 .35290 10.86524

(J) Day

MeanDifference

(I-J) Std. Error Sig. Lower Bound Upper Bound95% Confidence Interval

4567891011

The mean difference is significant at the .05 level.*.

Results


Figu

re

4.29

. Po

rosi

ty

dist

ribut

ion

of

(A)

deve

lopi

ng,

and

(B)

mat

ure

P.

aeru

gino

sa

PAO

1-C

FP b

iofil

ms.

Day

0

deno

ted

the

day

of

intro

duct

ion

of

legi

onel

lae

into

re

spec

tive

biof

ilms.

The

num

ber

of P

I pi

xels

of

each

ex

perim

ent

was

ob

tain

ed f

rom

3-8

im

age

stac

ks

from

1

or

2 co

upon

s. Th

e er

ror

bars

re

pres

ent

stan

dard

er

ror

of

3 in

depe

nden

t ex

perim

ents

.

Poro

sity

dis

trib

utio

n in

dev

elop

ing

biof

ilm

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

20% (b

ottom

)

x<

Sect

ions

of b

iofil

m

Porosity (Number of PI pixels per µm3) D

ay 0

Day

1D

ay 2

Day

3D

ay 4

Day

5

Poro

sity

dis

trib

utio

n in

mat

ure

biof

ilm

x<

(bott

om)

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

20%

Day

4

Sect

ions

of b

iofil

m

Porosity (Number of PI pixels per µm3)

Day

3D

ay 2

Day

1D

ay 0

B)

(

A)

(

Results


4.5

Sta significant linear correlation

bet lack of obvious relationship

bet e scatterplot below (Figure

4.3

Table 4.6. Table showing Pearson’s correlation between porosity and SBR. Each ontributing porosity and SBR data was obtained from 3-8 image stacks from 1 or

2 coupons of each experiment. The correlation between the 2 variables was btained from all data of 6 independent experiments.

Figure 4.30. Scatterplot of porosity and SBR both obtained from all data of 6 independent experiments.

.6 Correlation between SBR and porosity

tistical analysis revealed that there was no

ween SBR and porosity (Table 4.5), and the

ween SBR and porosity was further verified by th

0).

c

o

Correlations

1 -.012. .946

33 33-.012 1.946 .

33 33

Pearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)N

Porosity

SBR

Porosity SBR

0.200 0.400 0.600 0.800

SBR

2.000

4.000

6.000

8.000

10.000

Poro

sity

Results


4.5.7 Correlation between legionellae adh

Correlations

Legionellae adhesion to PAO1-CFP biofilm1.

10-.485.156

10.639*.047

10.030.934

10.303.394

10.309.385

10-.626.053

10-.178.623

10

Pearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)NPearson CorrelationSig. (2-tailed)N

Legionellae adhesionto PAO1-CFP biofilm

Porosity

SBR

Biomass

Average thickness

Maximum thickness

Substratum coverage


Correlation is significant at the 0.05 level (2-tailed).*.

esion and parameters of P.

amount of legionellae adhering to P. aeruginosa PAO1-CFP biofilm was

gionellae adhesion and the rest of the biofilm parameters.

Table 4.7. Table showing Pearson’s correlation between legionellae adhesion to . aeruginosa PAO1-CFP biofilm (representing the number of legionellae per

coupon per 106 legionellae inoculated into CBR) and parameters of the biofilm. or each experiments, 3-8 image stacks from 1 or 2 coupons were used. The

correlations between the variables were obtained from 10 independent xperiments.

aeruginosa PAO1-CFP biofilm

Pearson’s correlations between the number of legionellae per coupon per 106

legionellae inoculated into CBR and biofilm parameters are presented in Table

4.6. The

significantly positively correlated (at 0.05 level) to the overall SBR of biofilm.

Otherwise, there was no significant linear relationship between the amount of

le

P

F

e

Results


4.5.8 Localization of L. pneumophila in P. aeruginosa PAO1-CFP biofilms

Figure 4.31(A) and (C) revealed that L. pneumophila adhered to regions of high

porosity in both developing and mature P. aeruginosa PAO1-CFP biofilms, and

could also be found near the substratum. Four days later, majority of the

remaining legionellae were found “hidden” within the biofilms, away from

regions stained by PI (figure 4.31(B) and (D)).

(A)

Results


(C)

(B)

Results


(D)

Fi (blue) with

gionellae introduction to developing biofilm (3-days-old), (B) 4 days after legionellae introduction to developing biofilm, (C) 3hrs after legionellae

troduction to mature biofilm (7-days-old), and (D) 4 days after legionellae introduction to mature biofilm. Biofilm stained with PI (red) reflected porous

gions. The x-y view of the biofilm (main view) is flanked by y-z (right) and x-z (bottom) sections of the biofilm, with red arrows pointed towards the top of

iofilm. The scale represents 30µm in each image.

gure 4.31. CLSM images of P. aeruginosa PAO1-CFP biofilmadhered L. pneumophila (green) taken on different occasions: (A) 3hrs after le

in

re

b

Results


4

Kinetics of P. aeruginosa PAO1 Biofilm Formation at 30°C

0.000

0.010

0.020

0.030

0.040

0.050

30282624222018161412

OD

470nm

0.000

0.100

0.200

0.300

0.400

OD

600n

m

0.060

0.070

80

4038363432Time (hr)

0.500

0.600 0.0

Amount of biofilm (OD470nm) Amount of total bacteria (OD600nm)

.6 Screening for effective P. aeruginosa PAO1 biofilm removing

gent

P. aeruginosa PAO1 biofilm formation in microtiter plate

P. aeruginosa PAO1 biofilm formation reached a

ximum on the 18th hour, detached drastically after 20th hour and subsequently

ained low with OD470nm at approximately 0.010. The detachment corresponded

rise and high level of total bacteria in the well. Therefore biofilm removal

hour intervals starting from the 12th hour of

formation.

Figure 4.32. Kinetics of P. aeruginosa PAO1 biofilm formation in microtitre plate at 30°C. The error bars represent standard deviations of 3 independent experiments.

a

4.6.1 Kinetics of

As shown in figure 4.32,

ma

rem

to the

assays were conducted at every 2

biofilm

Results


4.6.2 P. aeruginosa PAO1 biofilm removal screening

Figure 4.33 revealed that NALCO 7330, NALCO 7320 and ACTI-PLUS 2818

had the highest comparable P. aeruginosa PAO1 biofilm removing efficiency. But

table 4.7 showed that NALCO 7320 had the highest efficacy since only 50ppm

was required to yield such a high percentage biofilm removal of 71.8% ± 16.8%.

Hence, NALCO 7320 was chosen for further characterization in P. aeruginosa

biofilm removal.

Figure 4.33. Highest percentage biofilm removal of various biofilm removing agents. The error bars represent standard deviations of 3 independent experiments.

Highest percentage biofilm removal of each biofilm removing agents

100.0%

(

-60.0%-40.0%-20.0%

0.0%20.0%40.0%60.0%80.0%

NALCO 73

30

NALCO 73

20

ACTI-PLU

S 2818

CT quart

erly c

leane

r

NALCO 90

001

NALSPERSE® 73

48

NALCO 73

38

NALCO 73

550Pe

rcen

tage

bio

film

rem

oval

%)

Results


Table 4.8. Efficacy of biofilm removing agents. The standard deviations were obtained from 3 independent experiments.

Biofilm removing agent Percentage biofilm Concentration Time

removal (%) (ppm) taken (hr) NALCO 7330 74.0 ± 4.2 1,000 6 NALCO 7320 71.8 ± 16.8 50 8

CT quarterly cleaner 51.9 ± 9.6 50,000 8

NALSPERSE® 7348 24.4 ± 7.8 50 0

NALCO 735

ACTI-PLUS 2818 67.6 ± 15.4 500 8

NALCO 90001 50.0 ± 22.9 500 6

NALCO 7338 17.5 ± 17.2 50 0 50 -7.9 ± 40.0 5 0

Results 4.7 Characterization of NALCO 7320

4.7.1 Kinetics of P. aeruginosa PAO1 biofilm removal

As shown in figure 4.34, biofil NALCO s time dependent

but no dependent ency of lowering removal efficacy

with increasing concentration abo Nevertheless, at 10ppm, no biofilm

rem

Figure 4.34. Kinetics of biofilm removal by NALCO 7320. The error bars represent standard deviations of 3 independent experiments.

m removal by 7320 wa

t concentration , with a tend

ve 50ppm.

oval was observed.

Kinetics of biofilm removal by NALCO 7320

-250.0%

-200.0%

-150.0%

-100.0%

-50.0%

0.0%

50.0%

100.0%

0 2 4 6 8Exposure time (hr)

Perc

enta

ge b

iofil

m re

mov

al

(%)

1000ppm500ppm100ppm50ppm10ppmMock treated


Results


4.7.2 Antimicrobial susceptibility testing

Minimum inhibitory concentration (MIC) is defined the lowest concentration of

spread plating.

Figure 4.35 demonstrated that MIC of NALCO 7320 on P. aeruginosa PAO1 and

L. pneumophila were 50ppm and 10ppm respectively. In addition, figure 4.36

revealed that MBC of NALCO 7320 on P. aeruginosa PAO1 and L. pneumophila

were 100ppm and 50ppm respectively. Hence, to ensure maximum biofilm

removal with bactericidal effect on planktonic P. aeruginosa PAO1, a final

concentration of 100ppm of NALCO 7320 was chosen for further characterization

in P. aeruginosa biofilm removal.

antimicrobial agent that completely inhibits the growth of the organism as

detected by the unaided eye while minimum bactericidal concentration (MBC) is

the lowest concentration of antimicrobial agent that completely eradicated the

organism as detected by

Results


NALCO 7320

Figure 4.35. Visual determination of minimum inhibitory concentration (MIC).

Figure 4.36. Determination of minimum bactericidal concentration (MBC) of NALCO 7320. The error bars represent standard deviations of 3 independent experiments. The dashed line denotes the detection limit of spread plating technique. ‘to’ represent the initial bacterial concentration before the addition of NALCO 7320.

L. pneumophila

P. aeruginosa

Bla

nk

PAO1

Moc

k tre

ated

500p

pm

100p

pm

50pp

m

10pp

m

1,00

0ppm

Determination of minimum bactericidal concentration (MBC)

9.0010.00

0.001.002.003.004.005.006.007.008.00

to

Mock t

reated

1k pp

m

500 p

pm

100 p

pm

50 pp

m

10 pp

m

Log

(CFU

/ml)

P. aeruginosa L. pneumophila

Results


Persistence of P. aeruginosa PAO1-CFP biofilms treated with NALCO 7320

0.0

1.0

2.0

3.0

4.0

5.0

6.0

Beforetreatment

0hr 4hr 8hr 12hr 24hr

Exposure time

Log

PAO

1-C

FP

conc

entr

atio

n [L

og

(CFU

/mm

2 )]

Developing biofilm - expt 1 Developing biofilm - expt 2Mature biofilm - expt 1 Mature biofilm - expt 2

4.8 Introduction of NALCO 7320 into developing and mature P.

aeruginosa PAO1-CFP biofilms containing L. pneumophila

4.8.1 Persistence of P. aeruginosa PAO1-CFP in CBR

Figure 4.37 showed that the concentration of viable P. aeruginosa PAO1-CFP

cells in mature biofilms decreased at a greater extent than that in developing

biofilms during the first 8hrs of exposure to NALCO 7320 and remained relatively

constant within the range of 1.00–2.00CFU/mm2 after 8 hours of exposure. The

concentration of viable P. aeruginosa PAO1-CFP cells in developing biofilms

decreased steadily for the first 12 hours of exposure and was not detected after

24hrs of exposure to NALCO 7320. Figure 4.38 showed that viable P. aeruginosa

of

treatment with NALCO 7320.

PAO1-CFP cell was not detected anymore in the bulk fluid of CBR after 8hrs

Figure 4.37. Viable cell counts of P. aeruginosa PAO1-CFP biofilms treated with NALCO 7320.

Results


P. aeruginosa PAO1-CFP in bulk fluid of CBR treated with NALCO 7320

7.0

0.0

1.0

3.0

4.0

treatment

Log

PAO

1-C

conc

entr

(CFU

/ml)]

2.0

Before 0hr 4hr 8hr 12hr 24hr

Exposure time

atio

n [L

og

5.0

6.0

FP


treated with NALCO 7320. The dashed line represented the detection limit of the

Figure 4.38. Viable cell counts of planktonic P. aeruginosa PAO1-CFP in CBR

plating technique used.

4.8.2 Structure of P. aeruginosa PAO1-CFP biofilms treated by NALCO 7320

Upon addition of NALCO 7320, bio-volume (Figure 4.39), average thickness

(Figure 4.40) and maximum thickness (Figure 4.41) of developing biofilm

immediately dropped and subsequently recovered by the 4th hr. Next, bio-volume

(Figure 4.39) and average thickness (Figure 4.40) dropped even lower than before

and remained low within the range of 6.00-10.0µm3µm-2 and 6.00-11.0µm

respectively, from the 8th hr of exposure to NALCO 7320 onwards. However, the

maximum thickness (Figure 4.41) dropped slightly and remain within the range of

15.0-22.0µm until 24th hr.

Results


In mature biofilm, by the 4th hr of exposure to NALCO 7320, bio-volume (Figure

ple t-test, p = 0.015, assuming equal variance) 4hrs

fter the addition of NALCO 7320 and remained at <65% thereafter. Substratum

ALCO 7320. Figure 4.43 demonstrated that the surface-to-biovolume ratio of

dependent sample t-test, p = 0.01, assuming equal

ariance) from <0.20 to 0.612 ± 0.220.

4.39) and average thickness (Figure 4.40) decreased significantly (independent

sample t-test, p = 0.002 and 0.007 respectively, assuming equal variance) and

remained within the range of 18.0-22.0µm3µm-2 and 21.0-26.0µm respectively, for

the subsequent 8hrs. However, maximum thickness (Figure 4.41) of mature

biofilm merely exhibited a decreasing trend. Nevertheless, at the end of 24hrs, the

bio-volume, average thickness and maximum thickness in both developing and

mature biofilm were comparable.

Figure 4.42 showed that substratum coverage of developing biofilm dropped

significantly (independent sam

a

coverage of mature biofilm remained comparable with or without exposure to

N

developing biofilm increased to >0.800µm2µm-3 after 24hrs of exposure. In

contrast, NALCO 7320 had no apparent effect on the surface-to-biovolume ratio

of mature biofilm.

Roughness coefficient of developing biofilm increased significantly (independent

sample t-test, p = 0.031, assuming equal variance) from <0.15 to >0.55 after 8hrs

of exposure to NALCO 7320 (figure 4.44). However, it was only after 24hrs of

exposure to NALCO 7320, when the roughness coefficient of mature biofilm

increased significantly (in

v

Results


with NALCO 7320. The bio-volume was obtained from at least 3 image stacks per coupon. The error bars represent standard deviation of 4 coupons from 2 independent experiments.

Figure 4.39. Bio-volume of P. aeruginosa PAO1-CFP biofilm in CBR treated

represent standard deviation of 4 coupons

Figure 4.40. Average thickness of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. The average thickness was obtained from at least 3 image stacks per coupon. The error barsfrom 2 independent experiments.

Bio-volume

25.0

30.0

35.0B

o-vo

lum

e (µ

m3 /µ

m2

0.0

5.0

10.0

15.0

20.0

Beforetreatment


Exposure time

i)

Developing biofilm (day 4) Mature biofilm (day 8)

Average thickness

20.0

25.0

35.0

40.0

ickn

ess

(µ

0.0

5.0

15.0

30.0

Exposure time

e th

m)

10.0

Beforetreatment


Ave

rag


Results


Figure 4.41. Maximum thickness of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. The maximum thickness was obtained from at least 3 image stacks per coupon. The error bars represent standard deviation of 4 coupons from 2 independent experiments.

Maximum thickness

0.0

5.0

10.0

15.0

20.0

25.0

30.0

35.0

40.0

45.0

Beforetreatment


Exposure time

Max

imum

thic

knes

s (µ

m)


Substratum coverage

0.0%

20.0%

40.0%

60.0%

80.0%

100.0%

120.0%

Beforetreatment


Exposure time

Subs

trat

um c

over

age

(%)


Figure 4.42. Substratum coverage of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. The substratum coverage was obtained from at least 3 image stacks per coupon. The error bars represent standard deviation of 4 coupons from 2 independent experiments.

Results


Figure 4.43. Surface-to-biovolume ratio of P. aeruginosa PAO1-CFP biofilm in CBR treated with NALCO 7320. The surface-to-biovolume ratio was obtained from at least 3 image stacks per coupon. The error bars represent standard deviation of 4 coupons from 2 ind

e error bars represent standard deviation of 4 coupons from 2 independent experiments.

igure 4.44. Roughness coefficient of P. aeruginosa PAO1-CFP biofilm in CBR

ependent experiments.

igure 4.44. Roughness coefficient of P. aeruginosa PAO1-CFP biofilm in CBR

Surface-to-biovolume ratio

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

1.8

Beforetreatment


Exposure time

Surf

ace-

to-b

iovo

lum

e ra

tio (µ

m2 /µ

m3 )



0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

Beforetreatment


Exposure time

Rou

ghne

ss c

oeffi

cien

t


FFtreated with NALCO 7320. The roughness coefficient was obtained from at least 3 image stacks per coupon. The error bars represent standard deviation of 4 coupons from 2 independent experiments.

treated with NALCO 7320. The roughness coefficient was obtained from at least 3 image stacks per coupon. The error bars represent standard deviation of 4 coupons from 2 independent experiments.

Results


4.8.3 Persistence of L. pneumophila in P. aeruginosa PAO1-CFP biofilms

o apparent linear correlation was found between legionella and bio-volume loss

treated with NALCO 7320

As shown in figure 4.45, there was a steady and gradual decrease in legionellae

viable cell counts in both developing and mature biofilms. Figure 4.46

demonstrated the presence of legionellae in the bulk fluid of CBR (at between 2.0-

2.5 cells/ml for every experiments) even after 24hrs of exposure to NALCO 7320.

Loss of legionellae per unit biomass lost from developing and mature biofilms

were calculated and no significant difference between them were found

(independent sample t-test, p = 0.414, equal variances not assumed). In addition,

n

(Table 4.8). Nevertheless, the scatterplot (Figure 4.47) revealed that 3 out of 4

data points have a linear relationship, thus implying the need for more work to

examine this relationship.

Results


Persistence of L. pneumophila in biofilms treated with NALCO 7320

1.0

3.5

Beforetreatment


Exposure time

Log

gion

ella

2

0.00.5

1.52.02.5

Leco

ncen

tr(c

ells

/mm

3.0

4.0at

ion

[Log

)]


treated with NALCO 7320.

Figure 4.46. Cell counts of planktonic L. pneumophila in CBR treated with NALCO 7320.

Figure 4.45. Persistence of L. pneumophila in P. aeruginosa PAO1-CFP biofilms

L. pneumophila in bulk fluid of CBR treated with NALCO 7320

3.5

4.5

atio

n [L

og

0.00.51.01.52.02.53.0

4.0

Beforetreatment


Exposure time

Log

Legi

onel

la

conc

entr

(cel

ls/m

l)]


Results


Correlations

1 .510. .4904 4

.510 1

.490 .4 4

Pearson CorrelationSig. (2-tailed)N

Bio-volumeloss

Legionellaeloss

Bio-volume loss

Pearson CorrelationSig. (2-tailed)

Legionellae loss

N

Table 4.9. Table showing Pearson’s correlation between bio-volume and legionellae loss. Each contributing bio-volume data was obtained from at least 3 image stacks from each of the 2 coupons used per experiment. The correlation between the 2 variables was obtained from 4 independent experiments.

dependent experiments.

Figure 4.47. Scatterplot of bio-volume and legionellae loss, obtained from 4

3000.000 3500.000 4000.000 4500.000

Legionellae loss

20.000

14.000

15.000

16.000

17.000

18.000

19.000

21.000

Bio

-vol

ume

loss

in

Results


4.8.4 Distribution of L. pneumophila in P. aeruginosa PAO1-CFP biofilms

treated with NALCO 7320

Despite increasing exposure time to NALCO 7320, the distribution of L.

ost L.

s while least

biofilm

rema L.

ature

biofilm

bottom

pneumophila in developing biofilm (figure 4.48(A)) remained similar, with the

exception of the fourth hour after NALCO 7320 addition. Generally, m

pneumophila resided in the 20%-60% region of developing biofilm

legionellae were found at the top 20% of the biofilm. But 4hrs after NALCO 7320

addition, the peak was temporarily shifted to 60%-80% of the biofilm.

Figure 4.48(B) showed that the distribution of L. pneumophila in mature

ined similar with increasing exposure time to NALCO 7320. Most

pneumophila resided in the 40%-80%, especially 60%-80% region of m

s while least legionellae were found at comparable levels at both top and

20% of mature biofilms.

Results


Ef

fect

of

L.

in

(A)

P.

aeru

gino

sa

obta

ied

e

ent

Figu

re

4.48

.N

ALC

O

7320

on

th

e di

strib

utio

n of

pn

eum

ophi

la

deve

lopi

ng,

and

(B)

mat

ure

PAO

1-C

FP b

iofil

ms.

The

num

ber

of C

FDA

pix

els

per

µm

3

nfr

om

at

leas

t 3

imag

erro

r ba

rs

repr

es

was

stac

ks p

er c

oupo

n. T

he

stan

dard

dev

iatio

n of

4

coup

ons

from

2

inde

pend

ent e

xper

imen

ts.

(A)

(B)

Effe

ct o

f NA

LCO

732

0 tr

eatm

ent o

n di

strib

utio

n of

L. p

neum

ophi

la i

n de

velo

ping

bio

film

-4.0

0

-3.5

0

-3.0

0

-2.5

0

-2.0

0

-1.5

0

-1.0

0

x<20

% (bott

om)

Sect

ions

of b

iofil

m

Log (Number of CFDA pixels per µm3) B

efor

e tre

atm

ent

0hr

4hr

8hr

12hr

24hr

Effe

ct o

f A

732

dist

ibut

n of

L.m

a b

i

-2.0

0

-1.5

0

-1.0

0

tom)

NLC

O

0 tr

eaen

tr

io

pne

umla

ture

ofilm

-4.0

0

-3.5

0

-3.0

0

-2.5

0

x<20

% (bot

tm o

n op

hi in

Sect

ins

of

o b

iofil

m

Log (Number of CFDA pixels per µm3) B

efor

e tre

atm

ent

0hr

4hr

8hr

12hr

24hr

Results


4.8 and mature biofilms treated

it

ig cell mass was rather uniformly

ist % biofilm, before and immediately

fte 4hrs later, the peak was shifted

.

ubsequently, the peak of bio-volume was progressively shifted towards the 20%-

0% region of the biofilm, maintaining at this spot until the 24th hr of exposure to

ALCO 7320.

igure 4.49(B) illustrated that peak bio-volume was found at 60%-80% region of

ature biofilm, before and up to the 8th hr of exposure to NALCO 7320.

ubsequently, the peak was shifted to 40%-60% region by the 12th hr of exposure

nd e tually to 20%-40% region by the 24th hr.

.5 Bio-volume distributions of developing

h NALCO 7320

ure 4.49(A) revealed that majority of the

ributed in the 20 -80% region of developing

r the addition of NALCO 7320. However,

w

F

d

a

upwards and was more concentrated at the 60%-80% region of the biofilm

S

4

N

F

m

S

a ven

Results


Effe

ct o

f NA

LCO

732

0 tr

eatm

ent o

n bi

o-vo

lum

e di

strib

utio

n in

dev

elop

ing

biof

ilm

0.00

1.00

2.00

3.00

4.00

5.00

6.00

7.00

8.00

x<20

% (bott

om)

Sect

ions

of b

iofil

m

Bio-volume (µm3µm

-2) B

efor

e tre

atm

ent

0hr

4hr

8hr

12hr

24hr

Effe

ct

bio-

volu

e di

2.00

3.00

4.00

5.00

6.00

7.00

8.00

tom)

of N

ALC

O 7

320

tm

strib

u i

a

biof

ilm

0.00

1.00

x<20

% (bot

reat

men

t on

tion

n m

ture

Sect

ions

of b

iofi

Bio-volume (µm3µm

-2)

lmB

efor

e tre

atm

ent

0hr

4hr

8hr

12hr

24hr

Figu

re

4.49

.N

ALC

O

7320

on

th

e di

strib

utio

n of

bi

o-vo

lum

e de

velo

ping

, an

d (B

) m

atur

e

aPA

O1-

CFP

bio

film

s. Th

e bi

o-vo

lum

e w

as o

btai

ned

stac

ks p

er c

oupo

n. T

he

stan

dard

dev

iatio

n of

4

coup

ons

from

2

inde

pend

ent e

xper

imen

ts.

Ef

fect

of

in

(A)

ugin

sa e

ent

P.er

o

from

at

le

ast

3 im

ag

erro

r ba

rs

repr

es

Results


4.8.6 treated with

AL

igu osity from 8.80 ± 1.95µm-3 to 14.8

0.4 fter the addition of NALCO 7320.

y t dropped and remained <4.50µm-3

.19µm-3 in mature biofilm was also observed immediately after the addition of

ALCO 7320. The level of porosity also dropped by the 4th hr of exposure and

mained <4.00µm-3 until the 24th hr.

igure 4.51(A) demonstrated that the porosity at the lower 60% of developing

iofilm increased drastically to >3.000µm-3 (with peak porosity at 20%-40%

gion of the biofilm) immediately after the addition of NALCO 7320 into the

BR. However, 4hrs later, the level of peak porosity dropped and remain within

e range of 1.000-1.500µm-3 at 40%-80% of the biofilm until the 24th hr of

enerally, NALCO 7320 had no observable effect on the porosity distribution of

ature biofilm where peak porosity was always found at 60%-80% region of the

iofilm (figure 4.51(B)). However, there was a noticeable increase in the porosity

f mature biofilm from peak porosity between 1.000-1.500µm-3 to >2.000µm-3),

mediately after the addition of NALCO 7320 into the CBR.

Porosity distributions of P. aeruginosa PAO1-CFP biofilms

CO 7320

re 4.50 illustrated a drastic increase of por

µm

N

F

-3 in developing biofilm immediately a

he 4

±

th hr of exposure, the level of porosityB

until the 24th hr. A slight increase of porosity from 3.49 ± 2.59µm-3 to 6.59 ±

1

N

re

F

b

re

C

th

exposure.

G

m

b

o

im

Results


Figure 4.50. P. aeruginosa

Porosity of PAO1-CFP biofilm in CBR treated with NALCO 7320. The porosity was obtained from at least 3 image stacks per coupon. The error bars represent standard deviation of 4 coupons from 2 independent experiments.

Porosity18.0

)

0.0

8.0

16.0

treatmenthr 8hr 12hr 24hr

er p

i p

e3

2.0

4.0

6.0

10.0

12.0

14.0

Before 0hr 4

Poro

sity

(Num

b o

f PI

xels

r µm

Developing biofilm (Day 4) Mature biofilm (Day 8)

Results


Figu

re

4.51

. Ef

fect

of

N

ALC

O

7320

on

po

rosi

ty

dist

ribut

ion

of

(A)

deve

lopi

ng,

and

(B)

mat

ure

P.

aeru

gino

sa

PAO

1-C

FP b

iofil

ms.

The

poro

sity

w

as

obta

ined

fr

om

at

leas

t 3

imag

e st

acks

per

cou

pon.

The

er

ror

bars

re

pres

ent

stan

dard

dev

iatio

n of

4

coup

ons

from

2

inde

pend

ent e

xper

imen

ts.

Effe

ct o

f NA

LCO

732

0 tr

eatm

ent o

n po

rosi

ty d

istr

ibut

ion

in d

evel

opin

g bi

ofilm

0.00

0.50

1.00

1.50

2.00

2.50

3.00

3.50

4.00

4.50

5.00

x<20

% (bott

om)

Sect

ions

of b

iofil

m

Porosity (Number of PI pixels per µm3) B

efor

e tre

atm

ent

0hr

4hr

8hr

12hr

24hr

Effe

ct73

20 tr

eatm

ent o

n po

rbu

tion

in m

atur

e of

ilm

0.00

0.50

1.00

1.50

2.00

2.50

3.00

3.50

4.00

4.50

5.00

x<20

% (bott

om)

24hr

of N

ALC

O

osity

dis

tri

bi

Sect

ions

of b

iofil

m

Porosity (Number of PI pixels per µm3)

12hr

8hr

4hr

t0h

rB

efor

e tre

atm

en

Discussion


Chapter 5: Discussion

Biofilm formation

To date, biofilm development is best studied in P. aeruginosa PAO1. The fusion

of an ecfp gene (encoding for enhanced cyan fluorescent protein) to a constitutive

promoter and subsequent insertion into a neutral intergenic region downstream of

the glmS gene on P. aeruginosa PAO1 genome (Klausen et al., 2003) allowed

CLSM observations of P. aeruginosa PAO1-CFP biofilms in the absence of

exogenous fluorescent dyes. The fluorescently tagged strain did not show any

phenotypic changes compared with the parental strain when tested in liquid

medium or flow chamber biofilms (Klausen et al., 2003). Similarly, the growth of

P. aeruginosa PAO1 and P. aeruginosa PAO1-CFP in minimal media supplied

with mannitol as the sole carbon source yielded indistinguishable growth curves

(Figure 4.2 and 4.3) in the present study.

The P. aeruginosa PAO1-CFP biofilm model was established at 30°C under high

shear, in the CBR system with continuous supply of minimal media containing

mannitol as the sole carbon source. After 6 days of growth under continuous

culture, both viable counts of P. aeruginosa PAO1-CFP in the biofilm (Figure

4.11) and the maximum thickness of the biofilm (Figure 4.14) reached a

reproducible plateau. This finding corroborated with a P. aeruginosa PAO1

biofilm development study, in which minimal medium containing glutamic acid as

the sole carbon source was used (Sauer et al., 2002). Thus in the present study,

mature biofilm is defined as one that has reached its maximum thickness on day 6

Discussion


while developing biofilm is one that has yet to reach its penultimate thickness

(before day 6).

Despite the attainment of maximum thickness on day 6 (Figure 4.14), the slight

increase in viable counts of planktonic (Figure 4.18) and biofilm (Figure 4.11) P.

aeruginosa PAO1-CFP, and in bio-volume (Figure 4.12) and average thickness

(Figure 4.13) of the biofilm on day 10 and 11, provided yet another experimental

evidence to support the hypothesis that biofilms never reached steady state

(Heydorn et al., 2000; Lewandowski et al., 2004). Early study by Bakke et al.

(1989) also demonstrated that older biofilms were continuously increasing their

density, even though their thickness remained constant. However, it is not possible

to differentiate between growth and attachment. Therefore, it is possible that

bacteria in the bulk fluid had been “captured” by mature biofilm. Additionally, no

sloughing event that might jeopardize the reproducibility of the biofilm structure

(Lewandowski et al., 2004), was observed during the course of this study.

As the P. aeruginosa PAO1-CFP biofilm matures, structural changes were

detected. The cell mass of biofilm started to move upwards on day 5, as suggested

by the drastic decrease in substratum coverage (Figure 4.15), the peak shift in

biomass distribution from 20%-40% to 40%-60% region of biofilm (Figure

4.26(A)), and drastic increase in SBR at bottom 20% of biofilm (Figure 4.27(A))

and in corresponding SBR ratio (bottom 20%: top 20% of biofilm) (Table 4.4).

Another upward movement of the cell mass was detected on day 9 when the

roughness coefficient of the biofilm dropped appreciably (Figure 4.17), a peak

Discussion


shift in biomass distribution from 40%-60% to 60%-80% region of the biofilm

was observed (Figure 4.26(B)), and the SBR at top 20% of biofilm decreased

noticeably (Figure 4.27(B)) resulting in the further increase in SBR ratio (Table

4.4). Eventually, the cell mass congregated at the 40%-80% region of the mature

P. aeruginosa PAO1-CFP biofilm with the least bio-volume found at the bottom

20% (Figure 4.26(B)). Expectedly, the region of biofilm with the lowest SBR

became increasingly prominent and coincided with that of the cell mass core on

day 5 onwards (Figure 4.27(B)). The roughness coefficient of P. aeruginosa

PAO1-CFP biofilm exhibited a general decreasing trend (Figure 4.17) thus

implying decreasing structural heterogeneity of the biofilm structure in the CBR

continuous flow system of this study. The production and redistribution of

biomass has been modeled in several investigations, where each model assumes a

different mechanism for biomass redistribution (Cogan and Keener, 2004;

Picioreanu et al., 2004; Alpkvist et al., 2006). However, in the absence of

empirical investigation, it is not clear how to judge the validity of the

redistribution mechanisms in the models.

Interestingly, despite the structural changes, the overall SBR remained relatively

uniform (Figure 4.16). Few studies applied the SBR function in COMSTAT in

their studies, but observations in this study corroborated with that of a previous

study in which P. aureofaciens, P. fluorescens and P. aeruginosa PAO1 each

exhibited relatively constant SBR throughout respective biofilm development and

structural changes (Heydorn et al., 2000). By extricating the overall SBR into 5

sections along the biofilm thickness, the present study demonstrated the

Discussion


usefulness of SBR distribution in providing insights into biofilm structural

differences.

Yang et al. (2000) first attempted to describe biofilm porosity using areal porosity.

Areal porosity is the ratio of the combined areas of the voids to the total area of

the image. However it is calculated from two-dimensional confocal images when

porosity characterizes three-dimensional space (Lewandowski, 2000). This study

presented an unprecedented method of quantifying porosity of paraformaldehyde

fixed biofilms by limiting the time of staining with PI (Chapter 3.4.4), obtaining

confocal image stacks under constant variables that may affect the quality of the

images (Chapter 3.8.7), applying constant threshold to all image stacks (Chapter

3.8.8.2) and quantifying the number of PI pixels per unit biomass (Chapter

3.8.8.4). PI has specificity for double stranded nucleic acids and bears a double

positive charge, thus readily enters and stains non-viable cells (Shapiro and Nebe-

von-Caron, 2004). Since with increasing incubation with 0.1mg/ml PI beyond 5

minutes resulted in excessive staining of the biofilm (Chapter 4.3.3), the amount

of PI molecules were not limiting. In this study, no viable bacteria were detected

from paraformaldehyde fixed biofilms, hence the possibility that the cells within

the biofilms remained viable (thus not picking up the PI dye) is eliminated.

Furthermore, extracellular DNA comprises <1-2% of the biofilm matrix

(Sutherland, 2001) therefore is not likely to contribute significantly to the number

of PI pixels detected. These imply that any variation in the number of PI pixels per

unit biomass is dependent on the accessibility of the biofilm cells to PI molecule,

thus reflecting the porosity of the biofilm in a three-dimensional context.

Discussion


Knowledge of the way in which substances are transported within biofilm is

essential for control or eradication. Based on the fact that biofilms consist of

microbial cell clusters separated by interstitial “voids”, “channels” or “pores”

(Lawrence et al., 1991; de Beer et al., 1994a and 1994b; Massol-Deya et al.,

1995), mass transport in the interstitial voids is mainly facilitated by convective

flow (Stoodley et al., 1994) and mass transfer inside the microbial clusters is

entirely due to molecular diffusion (de Beer et al., 1994a). On the contrary, Yang

and Lewandowski (1995) demonstrated that mass transfer coefficients were not

only found to vary both horizontally and vertically in the biofilm, they also

fluctuated significantly inside microbial cell clusters. This observation spurred the

proposal of a new conceptual model of biofilm microbial cluster structure, which

assumes the existence of flow channels with variable cross-sectional areas and

irregular orientations inside biofilm clusters. To the best of our knowledge, the

present study provided the first physical evidence of porous flow channels within

biofilm cell cluster (Figure 4.8).

Previous studies applied SBR to reflect the fraction of the biofilm that is exposed

to nutrient flow (Heydorn et al., 2000). Similar fraction of the biofilm has been

experimentally proven to be stained by PI (Figure 4.8) and can be represented by

the parameter “porosity” in the present study. On the contrary, overall SBR does

not correlate to overall porosity of P. aeruginosa PAO1-CFP biofilm (Chapter

4.5.6). Furthermore, there is no obvious correlation between the distribution of

SBR (Figure 4.27) and porosity (Figure 4.29). These observations suggest that the

structure of the biofilm alone is not enough to reflect the porosity of the biofilm.

Discussion


Although no attempts were made to detect EPS in the present study, it is well

established that biofilms comprise microbial cells within a matrix of EPS and

these microcolonies are separated by interstitial voids and channels. EPS, as the

major structural components of the biofilm matrix, has been implicated in the

protection of embedded microbial cells by either neutralizing or binding to toxic

substances, or merely serving as a physical barrier to environmental challenges

(Hall-Stoodley et al., 2004). Therefore, it is highly likely that the changes in the

quantity or nature of EPS had influenced the porosity of the biofilm in this study.

The overall porosity of the biofilm exhibited a general decreasing trend but

dropped significantly (p<0.01) on day 8 of development (Figure 4.28), suggesting

a drastic change in the quantity or property of EPS. However, throughout biofilm

development, the profile of porosity distribution remained comparable (Figure

4.29(A) and (B)). Thus suggesting the change in EPS was rather uniform

throughout the biofilm.

Discussion


Association of Legionella with biofilm

In this study, there was no significant difference (p=0.056) between the number of

legionellae adhering to developing and mature biofilm (Figure 4.20). The amount

of legionellae adhesion was found to be dependent on overall SBR of the biofilm

and independent on other biofilm parameters, especially porosity (Table 4.6). On

the contrary, legionellae adhesion patterns (Figure 4.23) did not emulate the

distribution patterns of SBR (Figure 4.27) for both developing and mature

biofilms. Interestingly, the legionellae adhesion patterns and biofilm porosity

distributions (Figure 4.23 and 4.29 respectively) were comparable, and the

attached legionellae were found co-localized with regions of high porosity even if

it was at the bottom of the biofilm (Figure 4.31(A) and (C)). These results

demonstrated that legionellae adhesion was dependent on the structure of the

biofilm, where biofilms with higher SBR can capture more planktonic legionellae,

but the adhesion might be hindered because legionellae only had access to biofilm

at areas of higher porosity. In a similar study, Langmark et al. (2005) found that

the accumulation of model pathogens (including L. pneumophila) was generally

independent of the biofilm cell density and was shown to be dependent on the

particle surface properties, where hydrophilic spheres accumulated to a larger

extent than hydrophobic ones. Taken together with the current study, the amount

and localization of legionellae adhering to biofilms may be determined by the

interplay of cell surface properties, biofilm structure and porosity.

In this study, figure 4.22 illustrated the 2-days delayed release of legionellae from

developing biofilm, until day 6 of biofilm development (Figure 4.21), which

Discussion


corresponded to biofilm maturation. The significant increase (p<0.01) in P.

aeruginosa PAO1-CFP biofilm detachment on day 6 (Figure 4.18) was likely the

cause of the sudden release of legionellae from P. aeruginosa PAO1-CFP biofilm

after the latter matured. This corroborated with another study which demonstrated

that detachment was one of the primary mechanisms affecting the loss of

microspheres and legionellae from biofilms within a pilot-scale distribution

system, as well as disinfection and biological grazing (Langmark et al., 2005).

Although the transport of particulates in biofilms has been largely neglected, it is

believed that in microbial competition in mixed population biofilms, slow

growing microorganisms are forced towards the biofilm surface and eventually

displaced (Okabe et al., 1996). In present study, legionellae release slowed down

(Figure 4.21 and 4.22) even though the bacteria was unable to replicate in the

continuous flow CBR system (which was fed with minimal media that supported

the growth of P. aeruginosa PAO1-CFP only) and the biofilm detachment

remained high, or even increased slightly on day 10 and 11 (Figure 4.18). In

addition, majority of remaining legionellae were found embedded in the biofilms,

away from porous regions (Figure 4.31(B) and (D)), implying reattachment of

planktonic legionellae to P. aeruginosa PAO1-CFP biofilm was not significant.

These indicate the existence of stable regions within P. aeruginosa PAO1-CFP

biofilm that harbored and protected legionellae from being desorbed. Figure 4.24

and 4.25 revealed that highest legionellae losses were found at the top 40% of the

biofilm while least legionellae losses were located at the bottom 60%, especially

at bottom 20%.

Discussion


The development of bimodal legionellae distribution 4 days after its adhesion to

developing biofilm (corresponding to day 7 of biofilm development) and

occurrence of alternate unimodal and bimodal distributions in mature biofilm

(Figure 4.23) revealed unbalanced advective transport of legionellae towards

biofilm surface took place after biofilm maturation. Similarly, Okabe et al. (1996)

observed that the trapped tracer beads were gradually transferred from the depth

of the biofilm to the surface but this advective transport was unbalanced. Since the

authors concluded that cell growth is an important factor for the entrapment and

release of the tracer beads, they attributed this phenomenon to unbalanced cell

growth. Therefore, it is likely that the concentration of biomass (thus cell growth)

near the substratum in developing biofilms (Figure 4.26(A)) resulted in unimodal

legionellae distributions (Figure 4.23(A)) and the faster loss of legionellae from

bottom 60% of developing biofilm (Figure 4.24) than from mature biofilm (Figure

4.25). On the other hand, the concentration of biomass in 40%-60% region of

mature biofilm (Figure 4.26(B)) was likely to result in bimodal legionellae

distributions (Figure 4.23), where legionellae from 40%-60% region of biofilm

were advected towards the surface of biofilm while legionellae loss at the bottom

slowed down (Figure 4.25). Therefore, the results from present study supported

the proposition of the existence of unbalanced cell growth in mature biofilm.

Discussion


Applications of biofilm-removing agents used in this study

Products from NALCO Company (www.nalco.com) such as NALCO 7320,

NALCO 7330, NALCO 7338, NALSPERSE® 7348 and NALCO 73550 are

registered as water treatment products to the NSF Registration Guidelines for

Proprietary Substances and Nonfood Compounds (www.nsf.org/usda) while

NALCO 90001 is registered to the New Zealand Food Safety Authority

(www.nzfsa.govt.nz). All the above products, except NALCO 73550, are

acceptable for treating boilers, steam lines and/or cooling systems where neither

the treated water nor the steam produced may contact edible products in and

around food processing areas. On the other hand, all the products, except NALCO

7320, 7330 and 90001, are acceptable for treatment of cooling and retort water in

and around food processing areas (www.nsf.org/usda). In addition, ACTI-PLUS

2818 is registered with U.S. Environmental Protection Agency (www.epa.gov),

under the Pest Control Products Act. It is an agent for controlling algal, bacteria

and fungal slime in condensing and cooling equipment to which recirculating

water is used as a cooling media. It can also be used to control bacterial and algal

slime in decorative fountains and brewery pasteurizers. Lastly, COOLING

TOWER QUARTERLY CLEANER was a product by Novapharm Research

(Australia) Pty Ltd., subsequently renamed and patented as Aeris-Guard

Enzymatic Coil Cleaner in 2003 (www.aerisguard.com). In March 2006 Quarterly

Report, Aeris Technologies Ltd. (www.aerisguard.com) reported several

successful applications of this product in both cooling towers and large industrial

water systems, and stated intentions to widen industrial applications in areas such

as mining operations and paper mills.

http://www.nalco.com/

http://www.nsf.org/usda

http://www.nsf.org/usda

http://www.aerisguard.com/

http://www.aerisguard.com/

Discussion


Effect of biocide NALCO 7320 on biofilm and associated legionellae

Without biofilm porosity as a concern, thin P. aeruginosa PAO1 biofilms of at

most 72hrs of age exhibited no correlation between initial cell density of the

biofilm and disinfection rate coefficient (Cochran et al., 2000). On the contrary, in

this study, the amount of viable P. aeruginosa PAO1-CFP cells in 8-days-old

mature biofilm decreased to a greater extent than in 4-days-old developing biofilm

during the first 8hrs of exposure to NALCO 7320, even when the former

contained more viable cells (Figure 4.37) and was relatively thicker than

developing biofilm (Figure 4.40). Since PI (molecular weight of 668.4) was able

to penetrate and stain the whole of 7-days-old mature biofilm within 30mins

(Figure 4.10), complete penetration of both developing and mature biofilms by

smaller 2,2-dibromo-3-nitrilopropionamide (DBNPA; molecular weight of 242)

was not likely to be hindered, if not faster. Thus the decreased resistance of the 8-

days-old mature biofilm to NALCO 7320 was most probably due to physiological

changes of biofilm organisms.

Such physiological changes could be explained by the fact that protein patterns for

dispersion stage biofilms (last stage of biofilm development) were reported to be

closer to the patterns observed from planktonic bacteria than for mature biofilms

(Sauer et al., 2002) and it is widely accepted that biofilm-grown cells are more

resistant to killing by biocides when compared with the same cells grown in

planktonic phase (Mah and O'Toole, 2001; Drenkard, 2003). Interestingly, biofilm

structures indicative of dispersion stage of biofilm development (Tolker-Nielsen

et al., 2000; Sauer et al., 2002) were observed occasionally and were earliest seen

Discussion


on day 7 of biofilm development, in the present study (Figure 4.19). To date, the

contribution of such dispersion mechanism to overall detachment of biofilm has

not been established.

Upon prolonged exposure to NALCO 7320 for 24hrs, no viable P. aeruginosa

PAO1-CFP cells were detected in developing biofilms while the amount of viable

cells in mature biofilms remained relatively constant since 8 hours of exposure

(Figure 4.37). Thus, suggesting the existence of either slow or non-growing cells,

or subpopulations of resistant phenotypes in mature biofilm. Occurrence of such

resistant P. aeruginosa subpopulations to DBNPA has been reported (Grobe et al.,

2002). Reflective of the bactericidal effects of NALCO 7320 on planktonic P.

aeruginosa PAO1 cells (Figure 4.36), no viable P. aeruginosa PAO1-CFP cells

were detected in the bulk fluid of CBR 8hrs after the addition of NALCO 7320

(Figure 4.38).

NALCO 7320 had different effects on the structures of developing and mature P.

aeruginosa PAO1-CFP biofilms. The effect on developing biofilm was immediate,

where the bio-volume (Figure 4.39), average thickness (Figure 4.40) and

maximum thickness (Figure 4.41) decreased immediately after NALCO 7320

addition, while those of mature biofilm remained relatively unchanged.

Additionally, the overall porosity of developing biofilm increased drastically

while that of mature biofilm only increased slightly (Figure 4.50). Upon closer

inspection, figure 4.51(A) demonstrated that the increase in porosity occurred at

bottom 60% of the developing biofilm but relatively uniform throughout mature

Discussion


biofilm. Four hours later, developing biofilm lifted off slightly from the

substratum but remained attached to the stainless steel coupons, as demonstrated

by the drop in substratum coverage (Figure 4.42), increase in average thickness

(Figure 4.40) and maximum thickness (Figure 4.41), and the shift in peak bio-

volume distribution to 60%-80% region of the biofilm (Figure 4.49(A)). As

established earlier in the discussion, changes in the quantity or nature of EPS

influence the porosity of the biofilm. Therefore, NALCO 7320 might have

immediate effect on the nature of EPS, which was then reflected by the increase in

porosity at the bottom 60% of developing biofilm where EPS accumulated, and

caused the lift off of developing biofilm from substratum. At the same time, the

increase in bio-volume suggested that portions of developing biofilm that had

detached might have been “recaptured” (Figure 4.39). Subsequently, majority of

the biomass accumulated at top 40% of the biofilm sloughed off by the 8th hr

(Figure 4.49(A)), resulting in a basal level of biomass (Figure 4.39), with low

average thickness (Figure 4.40) and substratum coverage (Figure 4.42), and high

SBR (Figure 4.43) and roughness coefficient (Figure 4.44). Interestingly, although

no viable P. aeruginosa PAO1-CFP cells were detected in developing biofilm at

24th hr (Figure 4.37), the basal biofilm structure remained (Figure 4.39).

By the 4th hr of treatment of NALCO 7320, the first significant drop (p<0.01) in

bio-volume (Figure 4.39) and average thickness (Figure 4.40) occurred, with

uniform decrease in bio-volume distribution throughout the mature biofilm

(Figure 4.49(B)). Thus, suggesting that the “outer layer” of mature biofilm had

peeled off. Interestingly, the bactericidal effects of NALCO 7320 continued to

Discussion


work on P. aeruginosa PAO1-CFP cells in mature biofilm while the structure

remained comparable for the subsequent 8hrs (Figure 4.39). Nevertheless, twenty-

four hours after the addition of NALCO 7320, the structure of the mature biofilm

was comparable with the basal structure of developing biofilm, where the peak of

biomass distribution was found at the 20%-40% region of the biofilms (Figure

4.49) and overall porosity remaining low (Figure 4.51). Taken together, it is

hypothesized that NALCO 7320 caused biofilm detachment by affecting the

nature of EPS that bound the microbial cells together as a microcolony, while


Persistence of biofilm basal structures after exposure to biocides, such as

hydrogen peroxide and isothiazolinone, was also reported by Schmid et al. (2004)

who investigated biocide efficacy by using photoacoustic spectroscopy for biofilm

monitoring. However, the present study further demonstrated that even when

majority of biofilm structure is removed, the existence of viable cells within the

basal biofilm is possible.

Legionellae cells persisted (Figure 4.45) within biofilms despite detachment of the

latter (Figure 4.39). The lack of significant difference (p=0.414) between

legionellae loss per unit biomass lost from developing and mature biofilms

(Chapter 4.8.3), together with the apparent lack of correlation between legionellae

loss from the biofilms and biofilm bio-volume loss (Table 4.8) suggests that the

persistence of legionellae in biocide-treated biofilms is dependent on factor(s)

other than biofilm structure, such as the nature of EPS, which may be affected by

NALCO 7320. Nevertheless, it was found that most legionellae persisted in 20%-

Discussion


60% and 40%-80% region of the remains of developing and mature biofilm

respectively, while least legionellae was found at top 20%, and both top and

bottom 20% of the remains of developing and mature biofilm respectively (Figure

4.48). This shows the existence of regions within the biocide-treated biofilms that

are more conducive for legionellae persistence. Furthermore, upon addition of

NALCO 7320, legionellae were found in the bulk fluid of CBR at high levels

(Figure 4.46), about more than 10× of that found in the bulk fluid of CBR without

NALCO 7320 added (Figure 4.42). This indicates high level of legionellae

detachment from biofilms treated with NALCO 7320, most likely together with

biomass from the P. aeruginosa PAO1-CFP biofilm.

Conclusion

In conclusion, the biofilm model set up in present study is reproducible and has

distinct developmental stages corroborating with that formed in other laboratory

(Sauer et al., 2002). To achieve greater insights of biofilm structure and properties,

certain data, such as bio-volume and SBR, were split up into 5 sections along the

biofilm thickness, and a method to quantify biofilm porosity was optimized and

applied in present study. Consequently, biofilm structures and development were

better described and the first physical evidence of porous flow channels within

biofilm cell cluster was discovered. Legionellae adhesion to biofilms was not

dependent on the developmental stage of the latter. Instead, biofilm structure and

porosity were found to determine the amount and even localization of legionellae

adhesion to biofilm. This opens up an unexplored possibility of controlling

legionellae colonization of existing biofilms in cooling towers by decreasing the

Discussion


porosity of biofilms. In addition, the bottom 60% of biofilms, especially at bottom

20%, was found to be stable regions within P. aeruginosa PAO1-CFP biofilm that

harbored and protected legionellae from being desorbed. Nevertheless, unbalanced

advective transport of legionellae towards biofilm surface took place after biofilm

maturation, and is most probably due to unbalanced cell growth. Thus, suggesting

that the bottom region of mature biofilm could harbor most legionellae eventually,

as compared to the rest of the biofilm. This further adds emphasis to the

overwhelming need of deep penetrating biocides to eradicate legionellae.

Developing P. aeruginosa PAO1-CFP biofilm was completely disinfected by

24hrs of exposure to NALCO 7320, which contained DBNPA as the active

ingredient, while a resistant subpopulation was found in the remains of mature

biofilm. NALCO 7320 exerted different effects on developing and mature

biofilms. From the porosity distribution data and biofilm structural analysis, it is

theorized that NALCO 7320 caused biofilm detachment by affecting the nature of

EPS matrix that bound the microbial cells together as a microcolony, while


Persistence of legionellae in biocide-treated biofilms was found to be independent

on the stage of biofilm development and loss of biomass, but there exists regions

of the biofilms in which legionellae best persist. Since EPS is a major component

in biofilm matrix, it may play an important role in legionellae persistence in

biocide-treated biofilms.

Discussion


Future Directions

Quick re-establishments of legionellae in biocide-treated cooling towers are

common (Kurtz et al., 1982). Therefore, it is necessary to improve on our

knowledge on legionellae persistence in biocide-treated biofilms. Since this study

suggests that EPS may play a role in legionellae persistence in biocide-treated

biofilms, further studies is required to verify this hypothesis.

In addition, it is also necessary to conduct further studies to determine whether

these persisting legionellae and even legionellae detached from the biofilm are

viable. Although 50ppm of NALCO 7320 was sufficiently bactericidal to

planktonic legionellae, it is unsure if 100ppm of NALCO 7320 is bactericidal to

legionellae detached from biofilms. This is because detaching biomass could

range from single cells to aggregate with a diameter of approximately 500µm

(Stoodley et al., 2001), which could provide shelter for legionellae from the

biocide in the bulk fluid. Problems can arise when concentrated numbers of such

biofilm-associated legionellae become detached from substrata (Stoodley et al.,

2001) where they have the potential to reach the consumer as an infective dose.

Last but not least, further studies should be conducted to determine the mode of

action of biofilm-removing agents used in this study, because this knowledge may

in turn provide insights into novel strategies of preventing public health problem.

However, since these biofilm-removing agents are proprietary products, closer

collaborations with the respective companies will be necessary.

References


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Appendix


Appendix Appendix I Edelstein BCYE liquid media: 2.0g Activated charcoal 10.0g Yeast extract 1L Deionized water Autoclaved at 121°C for 15mins. The media was allowed to cool before adding Legionella BCYE growth supplement (Oxoid Limited, UK) reconstituted as directed and filter sterilized. Appendix II Luria Bertani (LB) broth: 5g Yeast extract 10g Tryptone 10g NaCl 1L Deionized water Autoclaved at 121°C for 15mins. LB agar: Additional inclusion of 15g granulated agar in 1L LB broth and autoclaved at 121°C for 15mins. Appendix III Minimal media (MM): 10.5g K2HPO44.5g KH2PO42.0g (NH4)2SO42.0g Mannitol 0.2g MgSO4.7H2O 10mg CaCl25mg FeSO4.7H2O 2mg MnCl21L Deionized water Autoclaved at 121°C for 15mins. Appendix IV Phosphate Buffer Saline (PBS): 0.24g KH2PO41.44g Na2HPO48g NaCl 0.2g KCl 1L Deionized water Adjusted to pH 7.4 with 1N NaOH or 1M HCl, and autoclaved at 121°C for 15mins.

Appendix


Appendix V CFDA-SE stock solution (3.6mM): 1) Dissolve 2mg CFDA-SE (Molecular weight: 557) in 20μl DMSO 2) Top up to 1ml with ethanol (reagent grade) 3) Filter-sterilize & store at -20ºC in the dark 4) Working concentration: 10µM Appendix VI 4% Para-formaldehyde (PFA) solution: 1) Dissolved EM grade PFA in PBS with stir bar (4g to 100ml). 2) Add few drops of 1N NaOH and heat in hood (keep bottle cap loose) at 60°C

to dissolve. 3) Cool to room temperature and adjust to pH 7.4 with 1M HCl. *Prepare fresh prior to use.

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