Characterisation of Candida albicans, Actinomyces naeslundii ...

Characterisation of Candida albicans, Actinomyces naeslundii and Streptococcus

mutans interaction and its role in promoting oral carcinogenesis

By: Mohd Hafiz Arzmi

A thesis submitted in total fulfilment of the requirements for the degree of Doctor of Philosophy

April 2016

Melbourne Dental School Faculty of Medicine, Dentistry and Health Sciences

The University of Melbourne

1

ABSTRACT

Candida albicans has been widely reported in the aetiology of oral cancer.

However, the role of its interaction with members of the oral microbiome, such as

Actinomyces naeslundii and Streptococcus mutans, in promoting oral carcinogenesis

is still under investigation. The overall hypothesis of the present study is that

polymicrobial biofilms of C. albicans, A. naeslundii and S. mutans are involved in

oral carcinogenesis with the specific hypotheses as following: 1) Auto-aggregation

and co-aggregation of C. albicans is strain-dependent; 2) Polymicrobial biofilm

formation is C. albicans strain- and medium-dependent; 3) Polymicrobial interactions

within biofilms grown in flow-cells affects C. albicans biofilm formation; and 4) Oral

epithelial cells have an enhanced malignant phenotype when grown in the presence of

polymicrobial biofilm effluent.

The present study showed that C. albicans was able to auto-aggregate and co-

aggregate with A. naeslundii and/or S. mutans during planktonic growth. Co-

aggregation was shown to be variable between the eight strains of C. albicans with A.

naeslundii and S. mutans found to co-aggregate on both yeast and hyphae of C.

albicans. The static biofilm study showed that C. albicans formed yeast when grown

in 25% artificial saliva medium (ASM) and hyphae when grown in RPMI-1640.

Variability in biomass and metabolic activity was observed when C. albicans strains

were grown as mono-cultured and polymicrobial biofilms. In addition, ASM-grown

C. albicans, which predominantly forms yeast, was also able to form both mono-

cultured and polymicrobial biofilms in a flow-cell environment. Overall the biomass

of polymicrobial biofilms was found to be low relative to mono-cultured biofilms,

indicating antagonistic interactions between species. The present study showed that

biofilm effluent collected from flow-cell grown biofilms was able to promote oral

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carcinogenesis by increasing the adhesion of H357 cells (oral squamous cell

carcinoma cell line) to extracellular matrix molecules. Furthermore, the expression of

pro-inflammatory cytokines from H357 was found to increase when grown in

conditioned media suggesting that the biofilm effluent might have a role in the

promotion of oral carcinogenesis.

In conclusion, polymicrobial interactions of C. albicans A. naeslundii and S.

mutans promote oral carcinogenesis, thus supporting the hypothesis that


oral cancer by promoting carcinogenesis. Moreover, this carcinogenesis promoting

activity of polymicrobial biofilms is more likely to be C. albicans strain-specific.

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DECLARATION

This is to certify that:

i. The thesis comprises only my original work towards the PhD except where

indicated

ii. Due acknowledgement has been made in the text to all other material used

iii. The thesis is less than 100,000 words in length, exclusive of tables, maps,

bibliographies and appendices as approved by the Research Higher Degrees

Committee

Mohd Hafiz Arzmi

29th April 2016

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THE PREFACE

This PhD thesis comprises of seven chapters that determine the role of

polymicrobial interaction of C. albicans, A. naeslundii and S. mutans in promoting

oral cancer. The study was carried out under the supervision of Professor Michael

McCullough, Professor Stuart Dashper, Associate Professor Nicola Cirillo and

Associate Professor Neil O’Brien-Simpson. This study was done in collaboration

with Professor Eric Reynolds (Chapter 3 and 4), Deanne Catmull (Chapter 5), Dr.

Tanya D’Cruze (Chapter 5) and Dr. Jason Lenzo (Chapter 6). This is my original

work with all the experiments have been done myself. The analysis of data was

carried out with the contribution of collaborators. I have also contributed with more

than 80% of the published journals in this thesis.

The editorial assistance from Dr. Catherine Butler who is a knowledgeable

person in the academic discipline of the thesis has been very helpful during the

preparation of the thesis. Furthermore, two chapters of this thesis that have been

published in FEMS Yeast Research (Chapter 3) and Medical Mycology (Chapter 4),

were also thoroughly reviewed by the editors and reviewers of the journals.

Finally, I would like to acknowledge Ministry of Higher Education, Malaysia,

International Islamic University Malaysia (IIUM), Oral Health Cooperative Research

Centre (OHCRC), Melbourne Dental School, The University of Melbourne and

International Association of Dental Research (IADR), Australia for the scholarship

and research funding for this work.

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ACKNOWLEDGEMENT

I would like to thank to all my supervisors, Professor Michael McCullough,

Professor Stuart Dashper, Associate Professor Nicola Cirillo and Associate Professor

Neil O’Brien Simpson for all the input, knowledge and help in the completion of my

research and thesis. I also would like to thank Professor Eric Reynolds for his advice

and contribution especially in the publication of journals.

I would like to thank Deanne Catmull and Dr. Tanya D’Cruze for the assistant

in flow-cell biofilm experiment and Dr. Jason Lenzo particularly in flow cytometry

and Bio-Plex assays.

My gratitude to my parents, Arzmi Mansor and Safiah Abdul Aziz, my mother

in law, Norhayati Abdul Aziz and my younger brother, Muhammad Hazwan Arzmi

who have been giving a lot of supports and doa’ throughout my study.

Thanks also to my beloved wife, Nurul ‘Izzah Zulkifli, my beloved daughters,

Iffah Humaira Mohd Hafiz, Iffah Huriyya Mohd Hafiz and Iffah Huwayna Mohd

Hafiz for being very patient while Abi was so tired and moody. Thank you for

everything that you have sacrificed.

Finally, I would like to thank to all my colleagues, especially Dr. Ali

Alnuaimi, Dr. Antonio Celentano, Dr. Catherine Butler and Dr. Tami Yap for being

very supportive.

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ABBREVIATIONS

ANOVA Analysis of variance

ASM Artificial saliva medium

ATCC American Type Culture Collection

CV Crystal violet

EPS Extracellular polysaccharides

EMT Epithelial mesenchymal transition

FISH Fluorescence in situ hybridisation

g Gram

GM-CSF Granulocyte macrophage colony-stimulating factors

h Hour

IADR International Association of Dental Research

IL Interleukin

M Molar

mg Milligram

min Minute

mL Milliliter

mm Millimeter

nm Nanometer

OD Optical density

OSCC Oral squamous cell carcinoma

PBS Phosphate buffer saline

RNA Ribonucleic acid

rpm Revolution per minute

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RPMI Roswell Park Memorial Institute medium

s Second

SD Standard deviation

SDA Sabouraud’s dextrose agar

TEMED N,N,N,N-tetramethylethylenediamine

TNF Tumour necrosis factor

v/v volume/volume

WHO World Health Organization

XTT Tetrazolium salt, 2, 3-bis (2-methoxy-4-nitro-5-sulfophenyl)-5-

[(phenylamino) carbonyl]-2H-tetrazolium hydroxide

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TABLE OF CONTENTS

ABSTRACT ................................................................................................................... 1

DECLARATION ........................................................................................................... 3

THE PREFACE ............................................................................................................. 4

ACKNOWLEDGEMENT ............................................................................................. 5

ABBREVIATIONS ....................................................................................................... 6

TABLE OF CONTENTS ............................................................................................... 8

LIST OF TABLES ....................................................................................................... 14

LIST OF FIGURES ..................................................................................................... 17

CONFERENCES AND PUBLICATIONS.................................................................. 20

LITERATURE REVIEW ............................................................................................ 21

1.1 General Introduction ........................................................................................ 22

1.2 Oral Microorganisms ....................................................................................... 22

1.2.1 Candida species ........................................................................................... 23

1.2.1.1 Virulence factors of Candida species .................................................. 23

1.2.1.2 Prevalence of C. albicans in the oral cavity ........................................ 27

1.2.1.3 Growth requirements ........................................................................... 28

1.2.1.4 Clinical manifestation .......................................................................... 29

1.2.2 Actinomyces species ..................................................................................... 33

1.2.3 Streptococcus species................................................................................... 33

1.3 Intra-kingdom and inter-kingdom interaction of Candida spp. ....................... 35

1.3.1 C. albicans and non-albicans Candida spp. ................................................ 40

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1.3.2 Candida spp. and Actinomyces spp. ............................................................. 40

1.3.3 Candida spp. and Streptococcus spp. .......................................................... 41

1.3.4 Candida spp. and Pseudomonas spp. ........................................................... 41

1.3.5 Candida spp. and Porphyromonas spp. ....................................................... 42

1.3.6 Candida spp. and Prevotella spp. ................................................................ 43

1.4 Oral biofilms .................................................................................................... 43

1.4.1 Importance of oral fluids in the oral biofilm development .......................... 44

1.4.2 Importance of salivary flow rate in oral biofilm development .................... 46

1.5 Epidemiology of oral cancer ............................................................................ 47

1.6 Risk factors for oral squamous cell carcinoma ................................................ 48

1.6.1 Tobacco ........................................................................................................ 49

1.6.2 Alcohol ......................................................................................................... 50

1.6.3 Betel quid ..................................................................................................... 50

1.6.4 Dietary and genetic factors .......................................................................... 52

1.6.5 Microbial infection....................................................................................... 52

1.7 Cytokines and carcinogenesis .......................................................................... 53

1.7.1 Clinical significance of cytokines in carcinogenesis ................................... 54

1.8 Hypotheses ....................................................................................................... 60

1.9 Aims ................................................................................................................. 61

MATERIALS AND METHODS ................................................................................. 62

2.1 Growth of microorganisms .............................................................................. 63

2.2 Aggregation assay ............................................................................................ 63

2.3 Scanning Electron Microscopy (SEM) imaging .............................................. 65

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2.4 Static biofilm formation ................................................................................... 65

2.5 Gram stain ........................................................................................................ 66

2.6 Crystal violet (CV) assay ................................................................................. 67

2.7 XTT reduction assay ........................................................................................ 68

2.8 Flow-cell preparation ....................................................................................... 68

2.9 Flow-cell biofilm formation ............................................................................. 69

2.10 Flow-cell gel acrylamide preparation .............................................................. 70

2.11 Fluorescent In Situ Hybridisation (FISH) staining .......................................... 71

2.12 Confocal Laser Scanning Microscopy (CLSM) and image analysis ............... 72

2.13 Cell lines and culture ....................................................................................... 73

2.14 Preparation of test cell growth media .............................................................. 74

2.15 Cell-extracellular matrix (ECM) adhesion assay ............................................. 75

2.16 Preparation of cell suspensions for EMT and Bio-Plex assays........................ 76

2.17 EMT assay using flow cytometry .................................................................... 76

2.18 Bio-Plex assays ................................................................................................ 77

CO-AGGREGATION OF CANDIDA ALBICANS, ACTINOMYCES NAESLUNDII

AND STREPTOCOCCUS MUTANS IS CANDIDA ALBICANS STRAIN-

DEPENDENT .............................................................................................................. 79

3.1 Abstract ............................................................................................................ 80

3.2 Introduction ...................................................................................................... 81

3.3 Materials and methods ..................................................................................... 84

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3.4 Results .............................................................................................................. 85

3.4.1 Morphology of C. albicans in RPMI-1640 and 25% ASM ......................... 85

3.4.2 Auto-aggregation ......................................................................................... 85

3.4.3 Inter-kingdom co-aggregation ..................................................................... 88

3.4.4 Scanning Electron Microscopy analyses ..................................................... 88

3.5 Discussion ........................................................................................................ 92

3.6 Conclusion ....................................................................................................... 95

POLYMICROBIAL BIOFILM FORMATION BY CANDIDA ALBICANS,

ACTINOMYCES NAESLUNDII AND STREPTOCOCCUS MUTANS IS CANDIDA

ALBICANS STRAIN AND MEDIUM DEPENDENT ................................................ 96

4.1 Abstract ............................................................................................................ 97

4.2 Introduction ...................................................................................................... 98

4.3 Materials and methods ................................................................................... 101

4.4 Results ............................................................................................................ 103

4.4.1 Morphology of C. albicans bioflms in RPMI-1640 and 25% ASM .......... 103

4.4.2 Effect of microbial interaction and medium on biofilm biomass .............. 103

4.4.3 Effect of microbial interaction and medium on metabolic activity ........... 107

4.5 Discussion ...................................................................................................... 111

4.6 Conclusion ..................................................................................................... 115


ACTINOMYCES NAESLUNDII AND STREPTOCOCCUS MUTANS IN A FLOW

ENVIRONMENT ...................................................................................................... 116

5.1 Abstract .......................................................................................................... 117

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5.2 Introduction .................................................................................................... 119


5.4 Results ............................................................................................................ 123

5.4.1 Mono-cultured biofilms of C. albicans, A. naeslundii and S. mutans ....... 123

5.4.2 Polymicrobial biofilms of C. albicans, A. naeslundii and S. mutans......... 123

5.4.3 Effect of polymicrobial interaction on C. albicans, A. naeslundii and S.

mutans biofilms .............................................................................................. 124

5.5 Discussion ...................................................................................................... 131

5.6 Conclusion ..................................................................................................... 135

BIOFILM EFFLUENT OF CANDIDA ALBICANS, ACTINOMYCES NAESLUNDII

AND STREPTOCOCCUS MUTANS AFFECT THE ADHESION, EPITHELIAL

MESENCHYMAL TRANSITION AND CYTOKINE EXPRESSION OF NORMAL

AND MALIGNANT ORAL KERATINOCYTES .................................................... 136

6.1 Abstract .......................................................................................................... 137

6.2 Introduction .................................................................................................... 139


6.4 Results ............................................................................................................ 146

Part I: Adhesion assay ............................................................................................ 146

6.4.1 Adhesion of OKF6 to ECM ................................................................... 146

6.4.2 Adhesion of H357 to ECM .................................................................... 146

6.4.3 Comparison of adhesion to ECM between OKF6 and H357 ................. 147

Part II: Epithelial-mesenchymal transition (EMT) ................................................ 151

6.4.4 Percentage of cells expressing E-cadherin and vimentin ....................... 151

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6.4.5 Mean fluorescence intensity (MFI) ........................................................ 154

Part III: Cytokine assay .......................................................................................... 158

6.4.6 Expression of cytokines by OKF6 and H357 ........................................ 158

6.4.6.1 Interleukin 2 (IL-2) ........................................................................ 163

6.4.6.2 Interleukin 4 (IL-4) ........................................................................ 165

6.4.6.3 Interleukin 6 (IL-6) ........................................................................ 167

6.4.6.4 Interleukin 8 (IL-8) ........................................................................ 169

6.4.6.5 Interleukin 10 (IL-10) .................................................................... 171

6.4.6.6 Granulocyte-macrophage colony-stimulating factor (GM-CSF) ... 173

6.4.6.7 Interferon gamma (IFN-γ) .............................................................. 175

6.4.6.8 Tumour necrosis factor alpha (TNF-α) .......................................... 176

6.4.6.9 Overall............................................................................................ 177

6.5 Discussion ...................................................................................................... 181

6.6 Conclusion ..................................................................................................... 186

DISCUSSION AND CONCLUSION ....................................................................... 187

7.1 Discussion ...................................................................................................... 188

7.2 Conclusion and future studies ........................................................................ 196

REFERENCES .......................................................................................................... 197

APPENDICES ........................................................................................................... 245

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LIST OF TABLES

Table 1.1 Microorganisms isolated from oral cavity ................................................... 36

Table 1.2 Interaction of C. albicans with various important species of microorganisms

in the oral cavity .......................................................................................... 38

Table 1.3 Summary of cytokines that possibly have a role in the progression of OSCC

..................................................................................................................... 59

Table 3.1 Auto and co-aggregation scores of pairs of eight strains of RPMI-grown

(hyphal form) and ASM-grown (yeast form) C. albicans, A. naeslundii and

S. mutans. Percentage aggregation as measured by OD620nm change over 1

h ................................................................................................................... 90

Table 4.1 Static biofilm biomass scores of eight strains of RPMI-grown and ASM-

grown C. albicans, A. naeslundii (An) and S. mutans (Sm) as measured by

OD620nm after 72 h incubation .................................................................... 106

Table 4.2 Static biofilm metabolic activity scores of eight strains of RPMI-grown and

ASM-grown C. albicans, A. naeslundii (An) and S. mutans (Sm) as

measured by OD450nm-OD620nm after 72 h incubation .................................. 109

Table 4.3 Mono-culture metabolic activity per biofilm biomass (XTT/CV) scores of

eight strains of RPMI-grown (hyphal form) and ASM-grown (yeast form)

C. albicans, A. naeslundii (An) and S. mutans (Sm) ................................. 110

Table 5.1 Total biomass (µm3 µm-2) of ASM-grown C. albicans, A. naeslundii and S.

mutans after 24 h incubation in a flow-cell (3 mL h-1) at 37 °C in mono-

cultured biofilm and polymicrobial biofilms ............................................. 129

Table 5.2 Surface roughness, average and maximum thickness and percentage surface

colonisation of ASM-grown C. albicans, A. naeslundii and S. mutans after

24 h incubation in a flow-cell (3 mL h-1) at 37 °C .................................... 130

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Table 6.1A Adhesion of OKF6 and H357 in 80% serum free medium (SFM)

containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A.

naeslundii (AN), S. mutans (SM) and polymicrobial (TRI) biofilm effluents

................................................................................................................... 148

Table 6.1B Fold change of OKF6 and H357 adhesion when incubated with C.

albicans (ALC3), A. naeslundii (AN), S. mutans (SM) and polymicrobial

(TRI) biofilm effluents compared to non-effluent (NE) ........................... 149

Table 6.2A Percentage positive of OKF6 and H357 cells treated with 80% serum free

medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3),

A. naeslundii (AN), S. mutans (SM) and polymicrobial (TRI) biofilm

effluents at 37 °C, 5% CO2 for 2 h and 24 h.............................................. 152

Table 6.2B Percentage difference of positive OKF6 and H357 cells expressing

vimentin and E-cadherin between 2 h and 24 h incubated in NE, ALC3,

AN, SM and TRI effluents at 37 °C, 5% CO2 for 2 h and 24 h ................. 153

Table 6.3A Mean fluorescence intensity (MFI) of vimentin and E-cadherin of OKF6

and H357 cells treated with 80% serum free medium containing 20% of

non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S.

mutans (SM) and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2

for 2 h and 24 h .......................................................................................... 156

Table 6.3B Percentage difference of mean fluorescence intensity (MFI) of OKF6 and

H357 cells expressing vimentin and E-cadherin between 2 h and 24 h

incubated in NE, ALC3, AN, SM and TRI effluents at 37 °C, 5% CO2 for 2

h and 24 h .................................................................................................. 157

Table 6.4A Cytokines expressed by OKF6 (pg mL-1) incubated with 80% serum free


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effluent at 37 °C, 5% CO2 for 2 h .............................................................. 159

Table 6.4B Cytokines expressed by OKF6 (pg mL-1) incubated with 80% serum free



effluent at 37 °C, 5% CO2 for 24 h ............................................................ 160

Table 6.4C Cytokines expressed by H357 (pg mL-1) incubated with 80% serum free



effluent at 37 °C, 5% CO2 for 2 h .............................................................. 161

Table 6.4D Cytokines expressed by H357 (pg mL-1) incubated with 80% serum free



effluent at 37 °C, 5% CO2 for 24 h ............................................................ 162

Table 6.5A Percentage difference of cytokines expressed by OKF6 incubated in NE,

ALC3, AN, SM and TRI effluents at 37 °C, 5% CO2 between 2 h and 24 h

incubation .................................................................................................. 179

Table 6.5B Percentage difference of cytokines expressed by H357 incubated in NE,

ALC3, AN, SM and TRI effluents at 37 °C, 5% CO2 between 2 h and 24 h

incubation .................................................................................................. 180

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LIST OF FIGURES

Figure 2.1 Flow-cell system ......................................................................................... 71

Figure 3.1 Gram-stained of C. albicans cultures observed under light microscopy at

1000x magnification .................................................................................. 86

Figure 3.2 Percentage auto-aggregation in RPMI-1640 (A) and 25% ASM (B) grown

C. albicans after 1 h incubation in co-aggregation buffer ......................... 87

Figure 3.3 SEM of C. albicans (strain ALT4) auto-aggregation (A & E), inter-

kingdom interaction with A. naeslundii (B & F), S. mutans (C & G) and

both bacteria (D & H). C. albicans was grown in RPMI-1640 (A, B, C &

D) and 25% ASM (E, F, G & H) ............................................................... 91

Figure 4.1 Gram-stained biofilms of C. albicans strain ALC3 observed under light

microscope at 200x magnification after 72 h incubation at 37 °C in 24-well

plate at 90 rpm ......................................................................................... 105

Figure 5.1A Representative CLSM image of mono-cultured C. albicans as observed

using a 63x objective at 512 x 512 pixels magnification ......................... 125

Figure 5.1B Representative CLSM image of mono-cultured A. naeslundii as observed

using 63x objective at 512 x 512 pixels magnification............................ 126

Figure 5.1C Representative CLSM image of mono-cultured S. mutans as observed

using a 63x objective at 512 x 512 pixels magnification ......................... 127

Figure 5.1D Representative CLSM image of polymicrobial biofilms as observed using

a 63x objective at 512 x 512 pixels magnification (Red: C. albicans;

Green: A. naeslundii; Blue: S. mutans) .................................................... 128

Figure 6.1 Fold change of OKF6 and H357 adhesion when incubated with C. albicans

(ALC3), A. naeslundii (AN), S. mutans (SM) and polymicrobial (TRI)

biofilm effluents compared to non-effluent (NE) .................................... 150

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Figure 6.2A Fold change of IL-2 expression by OKF6 (left) and H357 (right) cells

incubated with 80% serum free medium containing 20% of non-effluent

ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and

polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h.

Fold change were the ratio of mean cytokine expression at 2 h and 24 h of

biofilm effluents (ALC3, AN, SM and TRI), to NE ................................ 164

Figure 6.2B Fold change of IL-4 expression by OKF6 (left) and H357 (right) cells






Figure 6.2C Fold change of IL-6 expression by OKF6 (left) and H357 (right) cells






Figure 6.2D Fold change of IL-8 expression by OKF6 (left) and H357 (right) cells






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Figure 6.2E Fold change of IL-10 expression by OKF6 (left) and H357 (right) cells






Figure 6.2F Fold change of GM-CSF expression by OKF6 (left) and H357 (right)

cells incubated with 80% serum free medium containing 20% of non-

effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans

(SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h

and 24 h. Fold change were the ratio of mean cytokine expression at 2 h

and 24 h of biofilm effluents (ALC3, AN, SM and TRI), to NE ............. 174

Figure 6.2G Fold change of TNF-α expression by OKF6 (left) and H357 (right) cells



polymicrobial (TRI) biofilm effluent at 37 °C, 5% CO2 for 2 h and 24 h.



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CONFERENCES AND PUBLICATIONS

1) Arzmi MH, Dashper S, Reynolds EC & McCullough M (2013) Intregenic and

intergenic coaggregation of Candida albicans, Actinomyces naeslundii and

Streptococcus mutans. New Zealand Microbiology Society, University of

Waikato, New Zealand.

2) Arzmi MH, Dashper S, Reynolds EC & McCullough M (2014) Co-

aggregation: Interaction between Candida albicans, Actinomyces naeslundii

and Streptococcus mutans. 54th Annual Scientific Meeting of the Australian &

New Zealand Division of the IADR, Brisbane, Australia.

3) Arzmi MH, Dashper S, Reynolds EC & McCullough M (2015) Polymicrobial

biofilms are Candida albicans strain and morphology dependent. 55th Annual

Scientific Meeting of the Australian & New Zealand Division of the IADR,

Dunedin, New Zealand.

4) Arzmi MH, Alshwaimi E, Harun WW, Razak FA, Farina F, McCullough M &

Cirillo N (2014) Gaining more insight into the determinants of Candida

species pathogenicity in the oral cavity. Eur J Inflamm 12: 227-235.

5) Arzmi MH, Dashper S, Catmull D, Cirillo N, Reynolds EC & McCullough M

(2015) Coaggregation of Candida albicans, Actinomyces naeslundii and

Streptococcus mutans is Candida albicans strain-dependent. FEMS Yeast Res

15.

6) Arzmi MH, Alnuaimi AD, Dashper S, Cirillo N, Reynolds EC & McCullough

M (2016) Polymicrobial biofilm formation by Candida albicans, Actinomyces

naeslundii and Streptococcus mutans is Candida albicans strain and medium

dependent. Med Mycol 54 (8): 856-864.

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CHAPTER 1

LITERATURE REVIEW

22

1.1 General Introduction

The oral cavity contains a multitude of microorganisms, including monera and

fungi (Dewhirst et al., 2010). They live in an ecological shared space, a community

in the human oral cavity (Nobbs and Jenkinson, 2015). The soft mucosal surfaces of

the tongue, gingival sulci, cheeks, lips, palate and tonsils, and the hard surfaces of

teeth provide convenient habitat niches for more than 700 oral microbial species

including Candida species, Actinomyces species and Streptococcus species

(Kolenbrander et al., 2010; Chandra et al., 2016).

The oral microbiome can exist in both planktonic and biofilm (plaque) forms

(Kolenbrander et al., 2010). An oral biofilm is defined as a community of

microorganisms that attach to surfaces in the oral cavity and are encapsulated within

extracellular polymeric substances (EPS), derived from both the microorganisms and

the oral environment (Filoche et al., 2010). It is suggested that cell-cell signalling

mechanisms between members of polymicrobial biofilms contributes to the successful

colonisation of bacteria in periodontal pockets (Yamada et al., 2005). For example,

Neisseria, an aerobic bacterium, has been found to metabolise the surrounding oxygen

and release carbon dioxide intensely, thus contributing to the survival of obligate

anaerobic bacteria such as Porphyromonas gingivalis (Kolenbrander et al., 2006;

Marsh, 2015).

1.2 Oral Microorganisms

It has been shown that more than 2000 bacterial taxa exist in the oral cavity

with a large number of opportunistic pathogens found to be involved in periodontal

and systemic diseases (Dewhirst et al., 2010; Warinner et al., 2014). These oral

microorganisms are classified into different kingdoms including monera (bacteria),

23

fungi (yeast) and protozoa. Previously, these microorganisms were classified

according to their morphology, fermentation of simple sugar and chemical analyses,

however, with the advance of DNA analysis technology, classification is now based

upon comparative 16S rRNA gene sequencing (Chaffin, 2008).

1.2.1 Candida species

Candida spp. belong to the Eukaryota domain and are known as imperfect

fungi within the family of Cryptococcaceae (Odds, 1979). These unicellular yeasts

possess globose, ellipsoidal and occasionally triangular microscopic morphology with

the sizes of the blastopores, hyphae and pseudohyphae varying between candidal

species (Kurtzman et al., 2011). There are at least seven Candida spp. that are

significant in human pathogenesis including C. albicans, C. kefyr, C. glabrata, C.

krusei, C. parapsilosis, C. dubliniensis and C. stellatoidea (Sida et al., 2016). C.

albicans has been reported to be the most prevalent in the oral cavity (Akdeniz et al.,

2002; Nejad et al., 2013). Some Candida spp. such as C. albicans and C.

dubliniensis, have the ability to form septate hyphae (Samaranayake, 2006). A few

Candida spp., such as C. krusei, have been shown to form pseudohyphae (Arzmi et

al., 2012). The cell wall of Candida spp. has been shown to possess ß-glucan,

mannoprotein and chitin that can assist in adhesion to oral surfaces and co-

aggregation with other bacteria, both in planktonic and biofilm growth (Chaffin,

2008).

1.2.1.1 Virulence factors of Candida species

Virulence factors of Candida spp. vary between species (Haynes, 2001). At

least seven significant virulence factors of Candida spp. have been reported, including

the ability to undergo phenotypic switching, cell surface hydrophobicity, production

24

of hydrolytic enzymes, dimorphism, candidalysin, quorum sensing and biofilm

formation (Williams et al., 2011; Kragelund et al., 2016; Sida et al., 2016).

Phenotypic switching is one of the most important virulence factors of C.

albicans, C. glabrata, C. dubliniensis and C. krusei (Anderson et al., 1987; Soll,

1992; Jones, et al., 1994; Lachke et al., 2000; Lachke et al., 2002; Vargas, et al.,

2004; Arzmi et al., 2012; Kragelund et al., 2016). The significance of the switching

strategy has similarities to human immune function and aims to counter threats in the

host’s environment.

Rapid phenotypic switching has been shown to enhance the survivability of

Candida spp. as an adaptive response to the stressful environment of the oral cavity

(Hellstein et al., 1993; Haynes, 2001). In a suppressed environment, it is postulated

that Candida spp. employs two mechanisms; mitotic recombination and subsequently,

phenotypic switching. A direct consequence of mitotic recombination is the loss of

heterozygosity throughout the entire genome. This deletion of the genome does,

however, affect the viability of Candida spp., especially in situations of multiple

modified conditions (Vargas et al., 2004). On the other hand, phenotypic switching is

a mechanism of adaptation that avoids alteration of the candidal genome. This spares

the alteration expression of genes, such as those involved in adhesion, and resistance

of Candida spp. to phagocytosis by polymorphonucleur leukocytes. Thus,

heterozygosity of the entire genome of Candida spp. is conserved (Marsh et al.,

2009).

In addition, cell surface hydrophobicity, another important virulence factor,

allows adherence of Candida spp. to host surfaces. The hydrophobicity among

Candida spp. varies, likely due to a difference in hydrophobic-associated surface

protein expression. There is a positive correlation between hydrophobicity and

25

interaction of Candida spp. with itself, endothelial cells and inert surfaces (Razak et

al., 2006; Silva-Dias et al. 2015).

The production of hydrolytic enzymes, such as aspartyl proteinase,

phospholipases, lipases, phosphomonoesterase and hexosaminidase has been reported

to contribute to the pathogenicity of Candida spp. in oral candidosis (Williams et al.,

2011). Aspartyl proteinase, coded by the secreted aspartyl proteinase (SAP) genes, is

considered central in the development of candidal infections. C. albicans and C.

krusei derived aspartyl proteinases have the ability to penetrate the host cell, thus

causing candidal infection (Samaranayake and Ferguson, 1994).

Another important hydrolytic enzyme in host invasion is phospholipase.

There are 4 types of phospholipase (type A, B, C and D). C. albicans produces

phospholipase A and C (Samaranayake and Ferguson, 1994). Phospholipase A can

attack cell membranes and is found on the cell surface, especially at bud formation

sites. Furthermore, the enzyme activity is enhanced when the hyphae are in direct

contact with host tissue (Williams et al., 2011).

Dimorphism is another important virulence factor of Candida spp. It is

defined as the ability of the microorganism to switch between yeast and hyphal

morphology depending on the environment and requirement. This is modulated by

gene expression within the cell (Nantel et al., 2002). A hypha (plural, hyphae) is a

long branching filamentous structure of fungus that consists of one or more cells

surrounded by tubular chitin-made cell walls. It is classified as the main mode of

vegetative fungal growth. Hyphae prefer to grow together in a compact tuft or

mycelium, particularly during the germination phase. In addition, this configuration

has also been shown to assist in the adhesion and colonisation of the host tissue by C.

26

albicans (Tronchin et al., 1988; Hawser and Douglas, 1994; San Millan et al., 1996;

Madigan et al., 2012).

The ability of Candida spp. to produce quorum sensing molecules has recently

been categorised as a virulence factor of this microorganism. It has been reported that

quorum sensing molecules are also involved in the colonisation of C. albicans in the

oral cavity (Shih and Huang, 2002). For example, Farnesol, secreted by C. albicans,

can freely pass across membranes and has been reported to be a biofilm-limiting agent

which inhibits the conversion of yeast to hyphal form in biofilm once its threshold is

reached (Ramage et al., 2002). Via these mechanisms, the number of microorganisms

within the consortia is optimised and overpopulation, which increases competition for

nutrients, oxygen, water and sites, is avoided. It has been suggested that one of the

factors that determines dissemination of Candida spp. infection within the oral cavity

is the secretion of Farnesol (Ramage et al., 2002). Detached Candida spp. cells will

adhere on the substratum where no Farnesol is detected (Hornby et al., 2001; Ramage

et al., 2002).

Candidalysin is newly discovered cytolytic peptide toxin that has been

classified as a virulence factor of C. albicans. This toxin is secreted by C. albicans

hyphae and has been shown to disrupt epithelial membranes, activate epithelial

immunity and trigger a danger response signalling pathway (Hofer, 2016). A mutant

strain of C. albicans lacking the candidalysin did not activate or damage epithelial

cells and was also shown to exhibit less tissue damage and neutrophil infiltration in

the tongues of mice than the wild-type, thus supporting the important role of

candidalysin as a virulence factor in the oral cavity (Moyes et al., 2016).

27

The ability to form biofilm on the oral surfaces is an important virulence

factor of Candida spp. in the oral cavity. Candida spp. adhesion initiates colonisation

of both hard and soft tissue surfaces in the oral cavity (Höfs et al., 2016). In general,

this adhesion occurs via non-specific and specific interactions. Non-specific

interactions consist of physicochemical forces such as Van der Waals forces,

electrostatic forces and acid-base interactions. Specific interactions involve protein

mediation between Candida spp. and the substratum (Van Oss, 1995).

Following adhesion, the attached cells grow and form a structured community

known as a biofilm. An oral biofilm or dental plaque is defined as a thin layer

comprised of various microbial communities encapsulated within EPS and attached to

a hard, soft or prosthetic surface (Holmes et al., 2002; Samaranayake et al., 2002).

The details of biofilm development will be further discussed in Section 1.4.

1.2.1.2 Prevalence of C. albicans in the oral cavity

C. albicans is a commensal yeast that inhabits the epidermis, vagina, gastro-

intestinal tract, nails and oral cavity (Williams et al., 2011; Chandra et al., 2016). It is

estimated that 70% of people with a healthy oral cavity have either a transient or

permanent residence of C. albicans (Mitchell, 2007; Thein et al., 2007). Several

factors that promote the conversion of C. albicans from commensal to opportunistic

microorganism in patients are impaired salivary gland function, a high carbohydrate

diet, tobacco smoking, drug abuse and use of broad-spectrum antibiotics. This is

commonly seen in the immunocompromised, including those using long-term

corticosteroids and common endocrine disorders such as diabetes mellitus (Williams

et al., 2011). It has also been associated with oral mucosal presentations and diseases

such as oral cancer and may play a role in symptoms of oral burning (Samaranayake,

2006; Scardina et al., 2007; Williams et al., 2011; Cavalcanti et al., 2016).

28

C. albicans infection can be local and/or systemic. Systemic infection of C.

albicans can be very severe and lead to fatality with mortality rates up to 60% (Leroy

et al., 2009). The treatment of the infection is difficult and its role may sometimes

only be determined by post-mortem. In the oral cavity, C. albicans has been found to

colonise mucosal surfaces including buccal and labial mucosa, dorsum or lateral

borders of tongue, hard and soft palate regions, as well as tooth surfaces and denture-

bearing areas (Harriott and Noverr, 2011). Virulence factors that contribute to the

successful colonisation of C. albicans in the oral cavity are previously discussed in

Section 1.2.1.1.

1.2.1.3 Growth requirements

There are many factors involved in the growth of Candida spp. in the oral

cavity. However, the two main factors are nutrients and host temperature.

The role of nutrients as a growth requirement has been widely discussed.

Candida is a chemoheterotrophic organism that requires carbon and nitrogen for

growth (Marsh et al., 2009). Carbohydrates are the most readily utilised form of

carbons in both oxidative and non-oxidative pathways. Thus, the presence of

carbohydrates influences the colonisation of Candida spp. in the oral cavity. Certain

carbohydrates, such as sucrose and glucose, have been shown to increase the adhesion

potential of C. albicans on to hard and soft oral surfaces (Samaranayake et al., 1986;

Jin et al., 2004). Glucose is an acid promoter that leads to the reduction of pH in the

oral environment, which activates acid proteinase and phospholipase enzymes, and

enhances the adherence capability of Candida spp. (Jin et al., 2004). Additionally, the

production of mannoprotein surface layer in the environment where glucose is present

has been shown to assist the adherence capability of Candida spp. (Modrzewska and

Kurnatowski, 2015; Demirezen et al., 2016).

29

Furthermore, body temperature has also been shown to influence the growth

of Candida spp. Candida spp. has been shown to grow at the optimal temperature of

37 ºC (Singh et al., 2002). This is also the optimal temperature for various pathogenic

microorganisms in the oral cavity, such as S. mutans and Actinomyces spp.

(MacFarlane and Samaranayake, 2014). Any alteration in normal body temperature

may influence the competitiveness of organisms within the normal microbiome, thus

enhancing the growth of opportunistic Candida spp. Many experimental assays are

conducted at 37 ºC and this is generally accepted as the standard incubation

temperature for candidal species (Marsh et al., 2009).

1.2.1.4 Clinical manifestation

Various classifications of candidosis and Candida spp. related disease have

been proposed (Meurman, 2016). The earliest classification was acute

pseudomembraneous candidosis (oral thrush), acute atrophic candidosis, chronic

atrophic candidosis and chronic hyperplastic candidosis (Scully et al., 1994). Oral

candidosis has also been described as primary oral candidosis, a candidal infection

confined to the oral and perioral tissue, and secondary oral candidosis, where the oral

presentation is a manifestation of systemic infection (Samaranayake et al., 1990).

Some keratinised primary lesions super-infected by Candida spp. have now been

classified under the category of primary oral candidosis such as leukoplakia, lichen

planus and lupus erythematous (Axéll et al., 1997). Furthermore, in later

classifications, the terminology of ‘acute’ and ‘chronic’ have been removed and

thought to have little bearing on the causality and treatment of the condition

(McCullough and Savage, 2005).

Primary oral candidosis has been divided into three major clinical variants,

pseudomembraneous, erythematous and hyperplastic. Furthermore, there are a further

30

four lesions that have been classified as primary oral candidosis that are denture-

associated erythematous stomatitis, chronic hyperplastic, angular cheilitis and median

rhomboid glossitis.

Pseudomembraneous candidosis (oral thrush) is a common disease of neonates

and elderly debilitated persons at rates of 5 to 10% (Samaranayake et al., 2009). The

lack of microbiota in the oral cavity of infants allows Candida spp. to flourish. In

elderly people, the disease can be caused by debilitation, xerostomia or atrophy of the

host immune system (Reichart et al., 2000). The disease is characterised by the

presence of white curd-like patches on the tongue, labial mucosal, cheeks, palate and

lips, that consist of dead mucosal cells, hyphal white plaques, blastospores,

inflammatory cells, fibrin and desquamated epithelial cells. The patches can be easily

removed leaving an erythematous background (Samaranayake et al., 2009). Oral

thrush can disseminate to the surfaces of oesophagus and pharynx, which leads to

feeding difficulties especially in infants (Klein and Klein, 1985).

The presentation of erythematous candidosis has previously been described in

association with the usage of broad-spectrum antibiotic therapy including tetracycline

(Williams et al., 2011). The surfaces affected may include the buccal mucosal, palate

and tongue. The mucosa may appear erythematous and atrophic (Millsop and Fazel,

2016). The condition can be caused by the overgrowth of commensal Candida spp.

due to the usage of broad-spectrum antibiotic therapy. This may inhibit the growth of

other antagonistic microorganisms to Candida spp., thus increasing the survival of the

yeast within the oral environment (Ito et al., 2015).

Denture-associated erythematous stomatitis (DAES) presents an area of

erythematous mucosa corresponding to the fitting surface of a prosthesis (Williams et

al., 2011). It is suggested to occur in up to 75% of denture wearers and is typically

31

asymptomatic (Webb et al., 1998b). Nocturnal denture wear as well as poor denture

hygiene are major predisposing factors (Budtz-Jörgensen, 1975; Williams et al.,

2011). These habits, including the accompanying limited flow of saliva over the area,

may allow an established biofilm that includes the overgrowth of Candida spp.

Biofilms on the oral prosthesis have been shown to be critical in the development of

DAES (Manfredi et al., 2013). Increasing surface roughness of dentures has been

shown to correlate with Candida spp. biofilm development (Lamfon et al., 2005).

Additionally, frictional irritation due to denture surface roughness may damage

normal mucosa if the denture is ill-fitting, which may allow the infiltration of

Candida spp. (Williams et al., 2011).

Another primary oral candidosis is hyperplastic candidosis (Candida

leukoplakia). The clinical presentation of this disease may include areas of keratosis,

particularly in the post commissure region of the buccal mucosa (Scardina et al.,

2007; Millsop and Fazel, 2016). The risk factors of the disease include impaired

salivary gland function, drugs, dentures, high carbohydrate diet, diabetes mellitus,

Cushing’s syndrome, malignancies and immunosuppressive conditions of patients

(Scardina et al., 2007). Hyperplastic candidosis usually develops as isolated white

patches intraorally and may be hard to differentiate between other causes of oral

diseases such as leukoplakia (Lamey and Samaranayake, 1988). The patches cannot

be removed with scraping and histopathologically will include hyphal elements

(Walker et al., 1990; de Lima et al., 2015). Marsh and Martin (1992) demonstrated

the production of nitrosamines in the saliva of patients that had been infected by C.

albicans. Nitrosamines are known carcinogens and the role of Candida spp. in the

development of oral cancer continues to be hypothesised, particularly in relation to

hyperplastic candidosis.

32

Angular cheilitis, which is classified as primary oral candidosis, may affect

individuals of any age. However, it is commonly seen in elderly patients

(Samaranayake and Holmstrup, 1989). Clinically, it presents as erythema and

ulceration of the angles of the mouth (Millsop and Fazel, 2016). Angular cheilitis has

also been reported to be associated with concomitant intraoral candidosis and worn

dentures (Budtz-Jörgensen et al., 1990; Farah et al., 2010).

Finally, median rhomboid glossitis (MRG) is characterised by a rhomboidal

shaped area of papillary atrophy of the mid-posterior dorsum of the tongue, anterior to

the circumvallate papillae. There may additionally be a corresponding lesion on the

palate. It is generally asymptomatic, symmetric and a flat/smooth lesion, although it

can be lobulated or nodular (Manfredi et al., 2013). Previously, Candida spp. were

suggested to be aetiological agents of MRG due to candidal hyphae observed

penetrating the superficial layers of the epithelium histopathologically (Scully et al.,

1994). However, the role of Candida spp. as causative agents in this disease remains

undetermined (Manfredi et al., 2013).

Chronic mucocutaneous candidosis is a rare disease that occurs in young

children and elderly males and is classified as a secondary oral candidosis (Williams

et al., 2011). The infection may be associated with disorders of T-cell production.

This type of candidosis commonly affects the oral cavity; it is usually infiltrated with

Candida spp. in thick granulomatous plaques (Farah et al., 2000; Liu and Hua, 2007).

It is important to understand that the removal of the candidosis may lead to other

infections by opportunistic bacteria including Pseudomonas spp. and Staphylococcus

spp. Chronic mucocutaneous candidosis may become a lifetime superinfection,

however the systemic imidazole antifungal agent ketonazole may aid in combating the

disease (Marsh and Martin, 1992).

33

1.2.2 Actinomyces species

Actinomyces spp. are facultative anaerobic Gram-positive rod bacteria that can

be easily isolated from the oral cavity (Schaal and Yassin, 2015). The bacteria have

been classified as part of the oral microbiome with some species categorised as

primary colonisers, such as A. naeslundii, that form fimbriae to assist in attachment to

oral surfaces (Kolenbrander et al., 2010).

A. naeslundii is the most common isolate of Actinomyces spp. in the oral

cavity and has been considered to be an early oral coloniser (Arai et al., 2015). The

genospecies of the bacterium is subdivided into genospecies 1 and 2. Genospecies 2

is now classified as A. oris (Yamane et al., 2013). Thus, A. naeslundii denotes what

was previously designated as genospecies 1. Some A. naeslundii strains are able to

produce urease, which modulates pH in the oral biofilm, as well as neuraminidase,

which can modify receptors in the acquired-pellicle to aid bacterial adhesion (Yaling

et al., 2008). Additionally, fimbriae that are present on the surface of A. naeslundii

contribute to colonisation of the oral cavity through cell-to-cell and cell-to-surface

adhesion, thus aiding in the formation of polymicrobial biofilms (Kolenbrander et al.,

2010).

1.2.3 Streptococcus species

The genus Streptococcus is comprised of facultative anaerobic Gram-positive

cocci bacteria. The nomenclature of Streptococcus spp. is in reference to their coccal

morphology in contrast to short rods or cocco-bacilli (Parks et al., 2015).

Streptococcus spp. have a tenacious binding ability for enamel surfaces and are

pathogens with high cariogenic potential. They form voluminous amounts of EPS

that facilitate the formation of oral biofilms (Samaranayake, 2006). The majority of

Streptococcus spp. are α-haemolytic on blood agar and referred to as the viridans-

34

group, that are clustered into four main groups: the mitis-group, salivarius-group,

anginosus-group and mutans-group (mutans streptococci) (Marsh et al., 2009).

The mutans-group of streptococci remains of great interest due to its role in

the aetiology of dental caries. Mutans streptococci include nine serotypes (a-h, and k)

which are determined by specific carbohydrate antigens present on the cell wall of the

bacteria. The major serotypes usually isolated from the human host are serotypes c, e,

f and k (Nakano et al., 2013).

S. mutans is the most common mutans-group isolate and is epidemiologically

implicated as the primary pathogen of enamel caries (Leme et al., 2006). It possesses

a cell wall with carbohydrate antigens, lipoteichoic acid and lipoproteins. Antigen I/II

is an example of an antigenic protein of S. mutans that is involved in the initial

adherence to the salivary pellicles on oral surfaces (Chuzeville et al., 2015). S.

mutans also produces polysaccharides that are comprised of fructan, glucan and

mutan metabolised from sucrose (Koo et al., 2010). These polysaccharides, in

association with S. mutans, help the formation and maturation of dental plaque (oral

biofilm) (Leme et al. 2006). Additionally S. mutans is able to communicate with

other members of the same group by releasing diffusible signalling molecules that

lead to the transfer of genes and provides a convenient condition for the growth of

other bacteria in the oral biofilm matrix (Marsh et al., 2009).

Even though infection by S. mutans is scarce on oral mucosal surfaces, reports

have shown a significant proliferation of the bacteria in disrupted mucosa and in

association with modified host defences such as in neutropenic patients (Tunkel and

Sepkowitz, 2002). Table 1.1 shows a range of other microorganisms that can be

isolated from the oral cavity.

35

1.3 Intra-kingdom and inter-kingdom interaction of Candida spp.

Candida spp. have been reported to be involved in both intra-kingdom and

inter-kingdom interactions (Morales and Hogan, 2010). Intra-kingdom interaction is

defined as communication between microorganisms of the same kingdom (such as

yeast to yeast) whereas inter-kingdom interaction is the communication between at

least two different kingdoms (such as yeast to bacteria). These interactions can occur

during planktonic growth and within the biofilm (Marsh et al., 2009).

36

Kingdom Bacteria

Gram-positive cocci Gram-negative cocci

Streptococcus anginosus

S. gordonii

S. mitis

S. mutans

S. oralis

S. salivarius

S. sanguinis

Staphylococcus spp.

Moraxella spp.

Neisseria mucosa

N. sicca

N. subflava

Veillonella parvula

V. dispar

V. atypica

Gram-positive rod Gram-negative rod

Actinomyces israelii

A. meyeri

A. naeslundii genospecies 1

A. naeslundii genospecies 2

A. odontolyticus

Bifidobacterium dentium

B. denticolens

Eubacterium minutum

E. nodatum

Lactobacillus acidophilus

L. casei

L. oris

L. salivarius

Actinobacillus actinomycetemcomitans

Bacteroides capillosus

B. forsythus

B. fragilis

Kingdom Fungi (yeast) Kingdom Protozoa

Candida albicans

C. glabrata

C. gulliermondi

C. krusei

C. parapsilosis

C. tropicalis

Entamoeba gingivalis

Giardia lamblia

Trichomonas tenax

Table 1.1 Microorganisms isolated from the oral cavity (Marsh et al., 2009).

37

Interactions of Candida spp. with bacteria have been reported to occur via co-

aggregation and co-adhesion (Kolenbrander et al., 2010). Co-aggregation and co-

adhesion are two important interactions in the oral cavity (Thein et al., 2007). Co-

aggregation is defined as the association of genetically distinct microorganisms such

as bacterium and yeast during planktonic growth (Gibbons and Nygaard, 1970; Bos et

al., 1996; Kolenbrander, 2000; Kolenbrander et al., 2002; Rickard et al., 2003; Al-

Ahmad et al., 2007; Ledder et al., 2008). This mechanism may involve the

interaction of proteins on the yeast surfaces and carbohydrate-containing molecules

on the bacterial surfaces (Arzmi et al., 2015).

Candida spp. co-adhesion is defined as the interaction between

microorganisms in planktonic growth with those adhered to an oral surface (Bos et

al., 1996; Kolenbrander, 2000; Kolenbrander et al., 2002; Rickard et al., 2003; Al-

Ahmad et al., 2007). This is significant for non-primary colonisers in biofilm

development, as C. albicans has been reported to co-adhere with S. gordonii which is

a primary coloniser. S. gordonii has been shown to produce high molecular mass cell

surface polypeptides, encoded by cshA and cshB that assist in the co-adhesion of the

bacterium to the yeast (Holmes et al., 1995). Many separate studies have investigated

the interactions between C. albicans and bacteria in the oral cavity and these are

summarised in Table 1.2. The following discussion will outline the key important

factors in the interaction between intra-kingdom Candida spp., between Candida spp.

and Streptococcus spp., Candida spp. and Actinomyces spp., Candida spp. and

Pseudomonas spp., Candida spp. and Porphyromonas spp., and finally Candida spp.

and Prevotella spp.

38

References Microorganisms Interaction Results

Purohit et al.,

1977

C. albicans and

L. acidophilus Antagonistic

L. acidophilus inhibited

growth of C. albicans.

Collins and

Hardt, 1980

C. albicans and

L. acidophilus Antagonistic

L. acidophilus produced

peroxidase and induced

growth retardation of C.

albicans.

Makrides and

MacFarlane,

1982

C. albicans and

E. coli Synergistic

Adherence of C. albicans

to epithelial cells was

increased in the presence

of fimbriae on E. coli.

Bagg and

Silverwood, 1986

C. albicans and

S. sanguinis Synergistic

C. albicans co-aggregated

S. sanguinis.

Verran and

Motteram, 1987

C. albicans and

S. sanguinis /

S. salivarius

Synergistic


to acrylic increased after

pre-incubation with S.

sanguinis / S. salivarius.

Branting et al.,

1989

C. albicans and

S. mutans Synergistic


increased with S. mutans

co-culture.

Jenkinson et al.,

1990

C. albicans and

S. sanguinis /

S. gordonii

Synergistic

Co-aggregation occurred

but inhibited after heat

and protease treatments.

Nair and

Samaranayake,

1996a; 1996b

C. albicans and

S. sanguinis /

S. salivarius

Antagonistic

Treatment of C. albicans

with S. sanguinis

decreased its adherence to

acrylic surfaces.

S. salivarius reduced

adherence of C. albicans

to human buccal epithelial

cells.

Table 1.2 Interaction of C. albicans with various important species of microorganisms in the oral cavity.

39

Nair and

Samaranayake,

1996a; 1996b

C. albicans and

E. coli Synergistic

Resistance of C. albicans

was increased.

C. albicans and

P. gingivalis Antagonistic


to acrylic and human

buccal epithelial cells was

suppressed.

C. krusei and

P. gingivalis Synergistic

Exposure of C. krusei to

P. gingivalis increased its

adherence to an acrylic

surface.

Busscher et al.,

1997

C. albicans and

S. thermophilus Synergistic

S. thermophilus mediated

adhesion of C. albicans to

silicone rubber.

Millsap et al.,

2000

C. albicans and

A. naeslundii /

S. gordonii /

S. sanguinis /

S. aureus

Antagonistic

A. naeslundii suppressed

C. albicans adherence to

an acrylic surface.

Nair et al., 2001

C. albicans and

L. casei Synergistic

Germ tube of C. albicans

was stimulated.

C. albicans and P.

intermedia Antagonistic

Germ tube formation of C.

albicans was suppressed.

Adam et al., 2002

C. albicans and

Staphylococcus

spp.

Synergistic

Staphylococcus spp.

adhered to C. albicans and

induced hyphal formation.

Hogan and

Kolter, 2002

C. albicans and P.

aeruginosa Antagonistic

P. aeruginosa formed a

dense biofilm and

adsorbed nutrients from

the hyphae of C. albicans

before killing.

(Continued)

40

1.3.1 C. albicans and non-albicans Candida spp.

Although C. albicans is the most significant Candida sp. in pathogenesis, non-

albicans Candida (NAC) species can additionally play a role. The occurence of NAC

species such as C. dubliniensis, C. glabrata and C. tropicalis has increased in

awareness among medical practitioners (Reichart et al., 2005). Surprisingly, in some

conditions, the carriage of NAC can outnumber that of C. albicans (Joshi et al., 1991;

Tzar et al., 2015). With regards to a mixed biofilm of Candida spp., research has

shown a competitive interaction between C. albicans and NAC in a polystyrene tube

model (El-Azizi et al., 2004). Furthermore, interaction of C. albicans with C. krusei

has been shown to vary depending on the nutrients supplied during the experiments

(Thein et al., 2007).

1.3.2 Candida spp. and Actinomyces spp.

Actinomyces spp. are Gram-positive bacteria that have been recognised as an

important human pathogen in the oral cavity (Schaal and Yassin, 2015; Steininger C

and Willinger, 2016). These microorganisms have been previously reported to be

involved in root surface caries and gingivitis (Shen et al., 2005). Candida spp. and

Actinomyces spp. have been found to co-aggregate in varying degrees with co-

aggregation of C. albicans with A. naeslundii reported as being C. albicans strain-

dependent (Grimaudo et al., 1996; Arzmi et al., 2015). This phenomenon has been

supported by subsequent findings that identified the interaction of protein complexes

on fungal surfaces with carbohydrate-containing moieties on A. naeslundii surfaces

(Grimaudo et al., 1996). Even though co-aggregation of C. albicans and Actinomyces

spp. has been widely reported (Bagg and Silverwood 1986; Grimaudo et al., 1996;

Arzmi et al., 2015), the adhesion of C. albicans to polymethylmethacrylate (PMMA)

in a flow-cell system when grown in TNMC buffer (1 mM Tris-HCl, 0.15 mM NaCl,

41

1 mM MgCl2, 1 mM CaCl2 in 1 L) has been shown to decrease when pre-cultured

with A. naeslundii T14V-J1 (Millsap et al., 2000). Chapters 3 and 4 will discuss in

more detail the interaction of C. albicans with A. naeslundii during planktonic and

biofilm growth, respectively.

1.3.3 Candida spp. and Streptococcus spp.

Streptococcus spp. form part of the human oral microbiome, with the mutans-

group being the most well-known for their high cariogenic potential (Klein et al.,

2015). Co-aggregation of C. albicans with S. sanguinis, and C. albicans with S.

mutans have been reported, suggesting that the interaction may be due to the presence

of streptococcal cell surface proteins that co-aggregate with cell surface carbohydrates

of C. albicans (Bagg and Silverwood, 1986; Arzmi et al., 2015). A similar finding

has been reported where protein-carbohydrate interactions have been shown to exist

between C. albicans with S. sanguinis, S. anginosus, S. oralis and S. gordonii

(Jenkinson et al., 1990). This mutualistic interaction is important to microorganisms

in dental plaque as it enhances the development of food chains using metabolic by-

products such as glycoproteins (Marsh, 1994). Chapters 3 and 4 will discuss in more

detail the interaction of C. albicans with S. mutans during planktonic and biofilm

growth, respectively. Chapters 3 and 4 will discuss in more detail the interaction of

C. albicans with S. mutans in planktonic and biofilm, respectively.

1.3.4 Candida spp. and Pseudomonas spp.

Pseudomonas spp. are Gram-negative bacilli that belong to the genus

Enterobacteriacea. These saprophytic aerobic bacteria possess virulence factors

including extracellular proteases, exotoxins and endotoxins (Samaranayake, 2006).

Studies have found that P. aeruginosa has the ability to suppress the growth of C.

albicans (Grillot et al., 1994; Kerr, 1994). The bacterium has been shown to form a

42

dense biofilm on C. albicans and receive nutrients from the candidal hyphae, which

later killed the fungus (Hogan and Kolter, 2002). P. aeruginosa has also been found

to inhibit C. albicans biofilm formation by synthesising anti-candidal agents such as

extracellular bacterial glycocalyx, pyrrolnitrin and 3-oxo-C12-homoserine lactone

(Hogan et al., 2004; Thein et al., 2006).

1.3.5 Candida spp. and Porphyromonas spp.

Porphyromonas spp. are Gram-negative coccobacilli that can be isolated from

dental plaque. These anaerobic, black-pigmented bacteria are classified as part of the

normal oral microbiome (Wade, 2013). P. gingivalis is highlighted as the most

important among these species due to its frequent isolation from sub-gingival sites

and identification as a major periodontal pathogen (Thein et al., 2007). For instance,

P. gingivalis has been shown to produce proteases that destroy haemolysin, haeme-

sequestering proteins, collagens, immunoglobulins and complements (Popova et al.,

2013; Antipa et al., 2015). The presence of fimbriae on the cell surface of the

bacteria mediates adhesion to the host cell while the capsule has been shown to resist

phagocytosis, which contributes to the virulence of P. gingivalis in periodontal

disease (Samaranayake, 2006). P. gingivalis has been found to suppress adhesion of

C. albicans to acrylic surfaces (Thein et al., 2006). This is supported by a subsequent

study that observed a significant inhibition of germ-tube formation of C. albicans by

P. gingivalis (Nair et al., 2001). The suppression of the adhesion of C. albicans to

denture acrylic surfaces by P. gingivalis suggests antagonistic interactions between

the microorganisms which may also occur in the oral cavity (Nair and Samaranayake,

1996b). It is suggested that metabolites of P. gingivalis and other anaerobic bacteria

may inhibit the colonisation of C. albicans in the gingival crevicular area (Thein et

al., 2006). Further study is required to identify the specific inhibitors.

43

1.3.6 Candida spp. and Prevotella spp.

Prevotella spp. are Gram-negative bacilli found in the human oral cavity. These

species, in particular P. intermedia, have been associated with the development of

periodontal diseases (Nair et al., 2001). Furthermore, P. intermedia has been shown

to suppress the development of germ-tube formation in C. albicans thus affecting

adhesion to the host cell (Nair et al., 2001). It is suggested that metabolites of P.

intermedia and other anaerobic bacteria may inhibit the colonisation of C. albicans in

the gingival crevicular area (Thein et al., 2006). Further study is required to identify

the specific inhibitors, which have not been previously reported.

1.4 Oral biofilms

The oral microbiome exists as both planktonic cells and oral biofilm

(Kolenbrander et al., 2010). An oral biofilm is defined as a community of

microorganisms that attach to oral surfaces and is encapsulated within EPS of

microbial and salivary origin (Donlan and Costerton, 2002). It is suggested that cell-

cell signalling mechanisms between members of a polymicrobial biofilm contribute to

the successful colonisation of bacteria. For example, Neisseria spp. have been found

to metabolise surrounding oxygen in the periodontal pocket releasing carbon dioxide

that contributes to the survival of obligate anaerobic bacteria (Kolenbrander et al.,

2006; Marsh, 2015).

Biofilms have been reported to increase the resistance up to 1000-fold of

opportunistic oral microorganisms, such as C. albicans, to antimicrobial agents

including fluconazole, thus increasing their pathogenicity towards the host (Stewart

and Corteston, 2001; Tobudic et al., 2012). It has been suggested that the EPS

encapsulating biofilms are a barrier limiting the penetration of antimicrobial agents,

44

subsequently reducing the susceptibility of the targeted microorganisms.

Furthermore, several enzymes produced by microorganisms within the biofilm have

been reported to neutralise active antimicrobial compounds, further reducing

susceptibility of targeted pathogens (Gilbert et al., 2002). Nutrient limitation has

been shown to occur within biofilms in the basal region that reduced microbial growth

and changed cell surface composition (Nadell et al., 2016; Ng et al., 2016). The

susceptibility to antimicrobial agents is thus modulated by biochemical pathways

associated with actively growing microorganisms (Mah and O'Toole, 2001, Donlan

and Costerton, 2002). Finally, slower growing microorganisms could represent

‘persister cells’ (LaFleur et al., 2006; Lewis, 2007; Lewis, 2010). It has been

suggested that persister cells possess a resistant phenotype which reduces their

susceptibility to antimicrobial agents (LaFleur et al., 2006).

1.4.1 Importance of oral fluids in the oral biofilm development

The oral cavity contains oral fluids such as saliva and gingival crevicular fluid.

These fluids are important in the first line of host defence as well as allowing the

proliferation of oral biofilms due to their nutrient content.

Saliva is an exocrine secretion comprised of water, proteins and electrolytes,

such as sodium, phosphate, calcium, magnesium, bicarbonate, phosphate and chloride

(Berkovitz et al., 2002). Saliva is produced by the major and minor salivary glands.

The major salivary glands consist of paired parotid, submandibular and sublingual

glands. The minor salivary glands are found throughout the lower lip, tongue, palate,

cheeks and pharynx (de Almeida et al., 2008).

The major roles of saliva include maintaining the integrity of teeth, a buffering

agent, and cleaning, protecting and lubricating the oral cavity (Dawes, 2008). Saliva

has been shown to maintain the physicochemical integrity of tooth enamel by

45

modulating remineralisation and demineralisation with calcium, fluoride and

phosphate and this has been reported to be the main factor controlling the stability of

enamel hydroxyapatite (de Almeida et al., 2008). Saliva also is an effective buffer

that neutralises potentially damaging acids of dietary and biofilm origin. The

composition of secretions from each gland differs while biochemical compounds,

including phosphates, bicarbonate and peptides, found in normal whole saliva, result

in a mean pH of 6.75 to 7.25 that prevents enamel demineralisation (Dawes, 2008).

The viscosity of saliva has been shown to provide mechanical cleansing of planktonic

microorganisms and food debris, thus limiting the nutrient intake by the oral biofilm

(de Almeida et al., 2008).

Saliva forms a seromucosal covering that lubricates and protects the oral

surfaces from microbial infection (Stack and Papas, 2001; Nagler, 2004). Research

has shown that saliva forms a thin film approximately 70 μm to 100 μm deep over all

external surfaces in the oral cavity, known as the salivary pellicle (Dawes, 2008).

Mucins, which are proteins with a high carbohydrate content, have been reported to

provide lubrication and protection to oral epithelial cells from dehydration, while

other salivary proteins are also involved in host defence mechanisms, such as

aggregation of exogenous microorganisms, thus facilitating clearance during

swallowing or expectoration (de Almeida et al., 2008).

Gingival crevicular fluid (GCF) is the exudate originating from plasma that

passes through the gingiva. This fluid influences the development of biofilms in the

oral cavity (Wade, 2013). GCF has been shown to assist the development of

subgingival plaque around and below the gingival margin (Marsh and Devine, 2011).

GCF contains a higher total protein content than saliva that may be converted to

nutrients such as peptides, amino acids and carbohydrates by bacterial enzymes

46

(Marcotte and Lavoie, 1998). In addition, GCF also contains antibodies that are both

non-specific and specific for a variety of periodontal disease important

microorganisms (Barros et al., 2016).

1.4.2 Importance of salivary flow rate in oral biofilm development

Circadian rhythms can affect salivary flow and composition, both of which are

important for diminishing colonisation by microorganisms (Edgar, 1992; Humphrey

and Williamson, 2001). The circadian rhythm results in the lowest flow rate

occurring during sleep (de Almeida et al., 2008; Dawes, 2008). Throughout the year,

the lowest flow rates are observed in summer whilst the highest flow rates are in

winter (Edgar, 1990). It has been suggested that this variability affects the

distribution of proteins such as mucin and agglutinin that serve as the receptors for

microorganisms on oral surfaces and thus assists the formation of oral biofilms

(Willet et al., 1991; Holmes et al., 2002). For example, the total concentration of

proteins in resting whole saliva is estimated at 220 μg mL-1 whereas in stimulated

saliva, it is approximately 280 μg mL-1 (Marsh et al., 2009). Furthermore, parotid

saliva alone has been shown to support the adherence of S. mutans to hydroxyapatite

beads (Carlen et al., 1996).

Flow rates also differ between individuals of various backgrounds. On

average, flow rate of un-stimulated saliva is 0.3 mL min-1 during waking hours with

an average of 300 mL for 16 hours when waking, and may drop to nearly zero during

sleeping hours (Edgar, 1990). Un-stimulated salivary flow rate is considered normal

above 0.1 mL min-1 whereas for stimulated saliva, the minimum volume accepted as

normal increases to 0.2 mL min-1 (Humphrey and Williamson, 2001). Any un-

stimulated flow rate that is below 0.1 mL min-1 is considered as hypo-salivation

(Edgar, 1990; Humphrey and Williamson, 2001). Meanwhile, stimulated saliva can

47

be up to 7 mL min-1 and has been reported to be 80% to 90% of the average salivary

production (Edgar, 1990).

Low flow rate has been shown to reduce the protective function of saliva in

the oral cavity, thus increasing the incidence of caries and the colonisation of

microorganisms such as Candida spp., Actinomyces spp. and Streptococcus spp.

(Loesche, 1986; Humphrey and Williamson, 2001; Holmes et al., 2002; Kolenbrander

et al., 2010; Pandey et al., 2015).

1.5 Epidemiology of oral cancer

In 2008, 12.4 million new cases and 7.6 million deaths due to cancer were

reported worldwide (Ariyawardana and Johnson, 2013). Of these, 263,000 cases of

lip and oral cavity cancer, and 135,000 pharyngeal cases were reported representing

2.1% and 1.1% of all new cancers, respectively (Ferlay et al., 2010). Oral cancer has

been classified as the sixth most common cancer in the world (Warnakulasuriya,

2009) and globally ranks as the eighth most common cancer in males and the 13th in

females (Parkin et al., 2005). It is estimated that the incidence of oral cancer is

around 275,000 cases with two-thirds of these reported in developing countries

(Ferlay et al., 2004).

The highest incidences of oral cancer have been reported in South-east Asia

(Taiwan), South Asia (India and Pakistan), Western Europe (e.g. France), Eastern

Europe (e.g. Slovakia, Slovenia and Hungary), parts of Latin America and the

Caribbean (e.g. Puerto Rico and Brazil) and Pacific regions (Papua New Guinea and

Melanesia) (Banoczy and Squier, 2004; Wünsch-Filho and de Camagro, 2001). In the

United States, oral cancer accounts for nearly 2.3% of all cancers and has a relatively

low five-year survival rate with almost 30,000 new oral cancer cases diagnosed every

48

year with 8,000 associated deaths (Silverman, 2001; Greenlee et al., 2001). Even

though surgical advances have improved the life quality for patients, the overall

mortalities remain unchanged (Casiglia and SB, 2001; Casto et al., 2009).

There were a total of 60,826 new cases of lip, oral cavity and oropharyngeal

cancer diagnosed between 1992 and 2008 in Australia. This corresponds to 2.9% of

the total cancer load and 1.6% of all cancer deaths in Australia (Farah et al., 2014). In

2009, oral cancer was classified as the eighth most diagnosed cancer in Victoria

(Alnuaimi et al., 2014). The most common site of new oral cancer cases was found to

be the lip, followed by the tongue (Sugerman and Savage, 2002; Farah et al., 2014).

Even though the incidence in Australia has been reducing in the past three decades, no

significant change was observed in the mortality rate (Ariyawardana and Johnson,

2013; Farah et al., 2014).

In Malaysia, oral cancer was found to be the 20th most common cancer for

females and 28th for males in 2006, with squamous cell carcinoma reported to be the

most common type of oral cancer (Omar et al., 2006; Ashazila et al., 2011; Helen-Ng

et al., 2012). In regard to ethnicity, individuals of Indian descent were found to have

the highest prevalence at 4.0%, followed by the indigenous people of Sabah and

Sarawak at 2.5% and the lowest prevalence was among those of Chinese descent at

0.5% (Zain et al., 1997; Zain; 2001).

1.6 Risk factors for oral squamous cell carcinoma

Oral squamous cell carcinoma (OSCC) has been classified as the most

common type of oral cancer (Ferlay et al., 2004), accounting for greater than 90% of

malignancies originating from the oral cavity (Casiglia and SB, 2001). It has been

49

reported that the average five-year survival rate following a diagnosis of oral cancer is

less than 50% (Zakrzewska, 1999).

OSCC is associated with a number of aetiological factors, including the use of

tobacco and heavy alcohol consumption. Other factors suggested to have a role in

OSCC formation include areca nut/betel quid chewing, dietary intake and microbial

infections (Warnakulasuriya et al., 2005).

1.6.1 Tobacco

Asia, Australia and the Far East are the largest tobacco consumers followed by

Americans, Eastern Europe and Western Europe (Petti, 2009). Global data on

smoking prevalence showed that almost one billion men in the world smoke with 35%

of those found in developed countries and 50% found in developing countries (Petti,

2009). Meanwhile, approximately 250 million women in the world are daily smokers

with 22% found in developed and 9% found in developing countries (Petti, 2009).

Even though cigarette smoking amongst women has been shown to be declining in

some developed countries such as Australia and Canada, several countries in

Southern, Central and Eastern Europe have shown increased or stagnant rates

(Mackay and Eriksen, 2002; Petti, 2009).

Tobacco smoking is a strong independent risk factor for oral cancer (Johnson,

2001; Winn, 2001; Vineis et al., 2004; Warnakulasuriya et al., 2005; Vallecillo

Capilla et al., 2007; Hirota et al., 2008; Pelucchi et al., 2008, Stucken et al., 2010). It

has been estimated that 43% to 60% of oral cancers are attributable to tobacco

smoking (Sasco et al., 2004) with the risk of developing oral cancer proportional to

the number of cigarettes smoked and duration of smoking. Pipe and cigar smokers

are at a higher risk for oral cancer development when compared to cigarette smokers

(Franceschi et al., 1990; Talamini et al., 2000; Lubin et al., 2009). The most

50

important carcinogens are tobacco-specific nitrosamines such as 4-

(methylnitrosamino-1-(3-pyridyl)-1-butanone, N-nitrosonornicotine, benzo[a]pyrene,

and aromatic amines (Hecht, 2003).

1.6.2 Alcohol

According to the World Health Organisation (WHO) report in 2014, it was

estimated that per year, an average of 6.2 L of pure alcohol was consumed by

individuals aged 15 years or older, with the highest consumption levels reported in the

developed world, particularly in the European region and the Americas (World Health

Organisation, 2014). In Australia, cancer has been reported to be attributable to the

consumption of alcohol. It is estimated that 3,208 cancers, which is equivalent to

2.8% of all cancers occurring in Australian adults, may be due to alcohol consumption

(Pandeya et al., 2015).

This is due to the breakdown of ethanol to the carcinogen acetaldehyde, which

has an established role in carcinogenesis (Schlecht et al., 2001; Huang et al., 2003;

Seitz et al., 2004; Boccia et al., 2009; Ogden, 2009; Warnakulasuriya 2009). In

animal studies, ethanol has been shown to have a synergistic effect when combined

with other carcinogens such as nitrosamines (Hooper et al., 2009). Furthermore, the

effect of both alcohol consumption and tobacco smoking has been reported to have a

multiplicative effect on carcinogenesis when compared to each factor in isolation.

(Pelucchi et al., 2008; Hashibe et al., 2009).

1.6.3 Betel quid

Piper betle is a plant belonging to the Piperaceae, which originated from

South East Asia including India, Sri Lanka and Bangladesh (Hoque et al., 2012). The

betel leaf itself is known as Sireh (Malay), Paan (Urdu and Hindi), Vetrilai (Tamil)

and Ikmo (Tagalog) (Datta et al., 2011). P. betle is an evergreen plant with glossy

51

heart-shaped leaves and white catkins. Usually, the leaf is chewed together with areca

nut, lime and gambier leaves (Johnson et al., 2011). The nut gives the reddish colour

to the saliva and thus darkens the teeth.

In South-East Asia and the Pacific Islands, 600 million people have been

reported to practice betel quid chewing, similar to the activity in South America of

chewing coca leaves or tobacco (Gupta and Ray, 2004). It is estimated that 10% to

20% of the world’s population, corresponding to 600 to 1200 million people, use betel

quid (Gupta and Ray, 2004). It is classified as the fourth most frequently consumed

psychoactive substance after nicotine, ethanol and caffeine (Pickwell et al., 1994;

Norton, 1998; Gupta and Ray, 2004).

Betel quid has been classified as an oral carcinogen in humans by the

International Agency for Research on Cancer, with evidence for a dose-response

relationship (Petti, 2009). It has been reported that betel quid chewing may produce

carcinogenic nitrosamines such as 3-methylnitrosopropionitrile (Petti, 2009).

Frequent and long-term betel quid consumption has been shown to increase oral

cancer risk. However, the risk of low to moderate betel quid chewing remains unclear

(Subapriya et al., 2007; Petti, 2009).

Even though betel quid has been classified as an oral carcinogen in humans, it

is culturally believed that the leaves can be a treatment for various diseases including

bad breath, headache, boils, conjunctivitis, itches, mastitis, mastoiditis and ringworm

(Chopra et al., 1956). The essential oil of P. betle was reported to contain

antibacterial, antiprotozoan and antifungal properties (Indu and Ng, 2002). Research

has shown that the plant may produce bacteriostatic and fungistatic effects against

Salmonella typhi, Escherichia coli and C. albicans respectively (Guha and Jain, 1997;

Indu and Ng, 2002). P. betle was found to be effective as an anti-dermatophyte

52

against C. albicans, Microsporum gypseum and Trichosporon beigelii and phyto-

pathogens such as Sclerotium rolfsii, Alternaria solani and Phytophthora infestans

(Rahman et al., 2005). Crude aqueous extracts of P. betle have been reported to

reduce the cell surface hydrophobicity of S. sanguinis, S. mitis and Actinomyces spp.

(Razak et al., 2006).

1.6.4 Dietary and genetic factors

Thirty to forty per cent of global cancer cases are attributable to unhealthy diet,

obesity and lack of physical activity and 10% to 15% of cases are associated with low

fruit and vegetable intake (Popkin, 2007). It has been suggested that the antioxidant

and anti-carcinogenic properties that exist in plants, such as vitamin A, C and E,

carotenoids, flavonoids, phytosterols, folates and fibres, play an essential role in

counterbalancing the detrimental effects of other carcinogenic substances, such as

betel quid chewing, alcohol consumption and tobacco smoking (Serdula et al., 1996;

Agudo et al., 1999).

1.6.5 Microbial infection

Since the 1960’s, Candida spp. have been suggested to be associated with oral

leukoplakic lesions (Cawson, 1969a). Candida spp. have been recognised as an

independent risk factor in the development of oral carcinoma (Cawson, 1969b). An

aetiological role for Candida spp. in the progression of oral mucosal keratoses to

carcinoma was first suggested by Cawson in 1966 (Cawson, 1969a). Researchers

have found that the majority of non-homogenous leukoplakias that are most often

invaded by C. albicans, have higher malignant transformation potential than the

homogenous types (Renstrup, 1970; Banoczy and Sugar; 1972, Cawson and Binnie,

1980; Pindborg, 1980; Axéll et al., 1984).

53

Candida spp., when associated with dysplasia, may represent a secondary

infection of a pre-existing altered epithelium (Barrett et al., 1998; Kragelund et al.,

2016). Furthermore, C. albicans was more commonly isolated from oral biofilms on

OSCC sites when compared to the control sites (Nagy et al., 1998). There is

additionally a correlation between oral yeast carriage and the presence of oral

epithelial dysplasia (McCullough et al., 2002). Despite many clinical and

experimental conclusions describing an association between C. albicans and

malignant transformation, the exact role of the yeast in the development of dysplastic

changes remains unclear (Supriya et al., 2016).

C. albicans produces C. albicans alcohol dehydrogenases (CaADH), the

enzyme that converts alcohol to acetaldehyde, which may play a role in oral

carcinogenesis. Bakri et al. (2010) showed that CaADH1 could utilise the pro-

carcinogenic substrate ethanol, using reversible conversion mechanisms, to produce

highly carcinogenic acetaldehyde. Acetaldehyde, a carcinogenic substrate, alters

epithelial cells, thus potentiating the formation of OSCC (Väkeväinen et al., 2002;

Salaspuro, 2003; Kurkivuori et al., 2007; Alnuaimi et al., 2015).

1.7 Cytokines and carcinogenesis

Inflammation induced by pathogens may be involved in carcinogenesis,

particularly after the classification of Helicobacter pylori as a class-1 carcinogen in

humans by the WHO International Agency for Research on Cancer (IARC) (Peek and

Blaser, 2002; Björkholm et al., 2003; Correa and Houghton, 2007). One factor that

may induce inflammation is the increase of pro-inflammatory cytokines and growth

factors due to microbial infection (Fantini and Pallone, 2008).

54

Cytokines are soluble proteins released by cells and are both autocrine and

paracrine in nature, and facilitate communication between cells (Lázár-Molnár et al.,

2000). Cytokine signals are received at the cell surface, not only as single messages,

but also in complex, subtle, synergistic and antagonistic combinations that coordinate

processes, including the stimulation of haematopoiesis, orchestration of directed

leukocyte migration (chemokinesis), activation of various inflammatory cells,

stimulation of lymphocyte development and maturation, and processes related to the

immune response (Budhu and Wang, 2006). However some circumstances, such as

failure to resolve an injury, might provoke excessive immune cell infiltration that then

leads to persistent cytokine production. As a result, the host may respond to the

persistent cytokine expression by enhancing cancer formation and progression (Budhu

and Wang, 2006).

1.7.1 Clinical significance of cytokines in carcinogenesis

Among cytokines synthesised by human epithelial cells, IL-6, IL-8, GM-CSF

and TNF-α have been widely studied (Kitadai et al., 2000; Riedel et al., 2005;

Kishimoto, 2006; Duffy et al., 2008; Lederle et al., 2011). It is suggested that these

cytokines may be involved in the progression of oral carcinoma.

IL-6 is a pleiotropic cytokine involved in the acute phase of inflammation and

is a major inducer of C-reactive protein (Kishimoto, 2006). This cytokine has been the

major mediator linking inflammation to cancers together with TNF-α (Kundu and

Surh, 2008; Balkwill and Mantovani, 2012). IL-6 has been reported to promote

malignant growth of skin squamous cell carcinoma by regulating a complex cytokine

and protease network (Lederle et al., 2011). IL-6 is one of the main chemokines

present in serum samples of head and neck cancer patients and elevated IL-6 levels

can independently predict tumour recurrence, poor survival, and tumour metastasis

55

(Riedel et al, 2005; Duffy et al., 2008). IL-6 in serum has been reported to be higher

in patients with OSCC compared to controls and has been proposed as an additional

marker in early detection of oral cancer (Jablonska et al., 1997; Dawes and Dong,

1995; Wu et al., 2015; Rao et al., 2016).

Even though much research has been performed to understand the role of IL-6

in carcinogenesis, very little is known about the direct role of this cytokine in head

and neck tumour metastasis and epithelial mesenchymal transition (EMT), which is

characterised by the loss of cell-cell junctions and cell polarity (Yadav et al., 2011).

IL-8 is an important cytokine that can be isolated from lipopolysaccharide

(LPS)-stimulated peripheral blood mononuclear cells (Oppenheim et al., 1991).

Research has shown that the cytokine is produced by various cells including

endothelial cells, fibroblasts, lymphocytes, neutrophils, keratinocytes, epithelial cells,

hepatocytes and lung macrophages (Kunkel et al., 1991; Kondo et al., 1993;

Bersinger et al. 2011). In addition, IL-8 can be produced by a variety of tumours,

both constitutively and in response to cytokines (Kolář et al. 2012). This cytokine has

also been detected in surgical specimens, fresh cultured cell lines and well-defined

cell lines of head and neck squamous cell carcinoma (Chen et al., 1999; Cohen et al.,

1995).

Two receptors for IL-8, IL-8RA (CXCR1) and IL-8RB (CXCR2), have been

identified on human neutrophils, both of which are members of the seven

transmembrane domain family of G-protein-associated receptors (Oppenheim et al.,

1991). Both of these receptors bind IL-8 with high affinity, but IL-8RA is more

specific for IL-8 whereas IL-8RB will also bind with similar affinity to other CXC

chemokines that possess a specific N-terminal motif. IL-8RA is expressed on a

variety of cells including neutrophils, T cells, monocytes and fibroblasts, whereas the

56

expression of IL-8RB is rather more restricted. Interestingly, these receptors of IL-8

have also been identified on tumour cells including squamous cell carcinoma (SCC)

(Cohen et al., 1995; Wang et al., 1996; Reiland et al., 1999; Brew et al., 2000). IL-8

stimulates many physiopathological functions in various tumour cells. Tumour cells

such as prostate carcinoma (Reiland et al., 1999; Inoue et al., 2000), melanoma (Luca

et al., 1997), breast carcinoma (Youngs et al., 1997) and gastric carcinoma (Kitadai et

al., 2000) cell lines respond chemotactically to IL-8. Furthermore, high expression of

IL-8 in cancerous liver tissue has been shown to associate with a higher frequency of

portal vein, venous, and bile duct invasion in hepatocellular carcinoma (HCC)

patients with surgical resection and may therefore be important in invasion and

metastasis (Akiba et al., 2001).

IL-18 is suggested to have a role in carcinogenesis. This cytokine has been

shown to be synthesised as a 24 kDa inactive precursor in cells such as macrophages,

dendritic cells, Kupffer cells and some tumour cells (Sugawara, 2000, Cho et al.,

2002). The inactive cytokine is cleaved by the IL-1β converting enzyme (caspase-1)

in the cytoplasm and then secreted as an 18 kDa active protein, which can induce the

formation of interferon gamma (IFN-γ) (Ushio et al., 1996; Sugawara, 2000; Gracie et

al., 2003). Enhanced IL-18 expression is positively correlated with the pathogenesis

of malignant skin tumours (Park et al., 2001). It has also been shown that this

cytokine regulates hepatic melanoma metastasis by increasing the adherence of

melanoma cells and the expression of vascular cell adhesion molecule-1 (Vidal-

Vanaclocha et al., 2000). Furthermore, IL-18 secreted by the B16 murine melanoma

cell line has been shown to be involved in the immune escape of murine melanoma

cells (Cho et al., 2000). Therefore, it is suggested that IL-18 may be important in the

formation of OSCC.

57

Granulocyte macrophage colony-stimulating factor (GM-CSF) has been

shown to be important in carcinogenesis. It is termed a growth factor since it supports

the colonisation of granulocyte, macrophage, erythroid, megakaryocyte and

eosinophil progenitor cell lines (Burgess and Metcalf, 1980). GM-CSF is a 127

amino acid monomer with a mass ranging from 14 to 35 kDa depending on the

amount of glycosylation in vivo (Cantrell et al., 1985). On mature haematopoietic

cells, GM-CSF activates the effector functions of granulocytes,

monocytes/macrophages and eosinophils (Morrissey et al., 1987). It is produced and

released by various cell lines in response to immune and/or inflammatory stimuli,

including activated T cells (Kharkevitch et al., 1994), B cells (Pistoia et al., 1993),

mast cells (Levi-Schaffer et al., 1998), endothelial cells (Sieff et al., 1987)

and

fibroblasts (Koeffler et al., 1987). Recently, GM-CSF has been reported to have a

functional role on non-haematopoietic cells by inducing human endothelial cells to

migrate and proliferate (Budhu and Wang, 2006). Interestingly, GM-CSF can also

stimulate the proliferation of a number of tumour cell lines, including breast cancer

cell lines (Park et al., 2007). Cancer cells have been shown to produce GM-CSF

(Burgdorf et al., 2009). Therefore, increased expression of this cytokine can be used

as a biomarker in the detection of OSCC.

Since 1987, tumour necrosis factor (TNF)-α has been reported to be involved

in breast cancer with TNF mRNA and protein detected in malignant and stromal cells

in human biopsies (Spriggs et al., 1987). Malignant cell-derived TNF has been

reported to enhance the growth and spread of syngeneic-, xenogeneic- and

carcinogenic-induced tumours of the bowel, pancreas, skin and ovary (Kundu and

Surh, 2008; Balkwill, 2009). Research has found that TNF-α in HCC patients was

higher in metastatic liver carcinoma than the healthy individual (Nakazaki, 1992).

58

Furthermore, higher levels of TNF-α were found in the tissue surrounding HCC and

hepatic metastasis than in the tumour (Bortolami et al., 2002). This pro-inflammatory

and pro-angiogenic cytokine has also been shown to increase from OSCC tumour

cells together with IL-1, IL-6 and IL-8 compared to normal cells (SahebJamee et al.,

2008). The mechanisms of the increase of TNF-α during tumour growth are not fully

defined. In an ovarian cancer model, TNF-α was an important component of the

malignant cell-autonomous network of inflammatory cytokines, including

chemokines stromal cell-derived factor 1 (SDF1) and C-C chemokine ligand 2

(CCL2), which have been suggested to aid in the proliferation and survival of

malignant cells, stimulating angiogenesis and metastasis (Kulbe et al., 2007).

Other pro-inflammatory cytokines involved in carcinogenesis are IL-4, IL-10

and interferon-gamma (IFN-γ). IL-4 has been reported as both a pro- and anti-

inflammatory cytokine (Brieland et al., 2001; Hosoyama et al., 2011). A study using a

mammary carcinoma model has shown that Th2-derived IL-4 is responsible for the

promotion of metastasis (DeNardo et al., 2009). IL-4 has also been reported to

accelerate tumour cell growth from rhabdomyosarcoma (RMS) tissue, suggesting that

the cytokine could potentially be a growth factor acting on tumour cells directly or

indirectly through tumour associated macrophages (TAMs) (Hosoyama et al., 2011).

In addition, IL-10 has been shown to promote tumour progression together with

regulatory B cells during squamous carcinogenesis (Schioppa et al., 2011). IL-10 was

found to inhibit major histocompatibility complex type II dependent antigen

presentation, activation of type I helper T cells and autologous T cell specific tumour

lysis from melanoma cells, suggesting that a mechanism of escape of tumour cells

from the human immune system may have been activated through secretion of the

cytokine (Tartour and Fridman, 1998). Furthermore, interferon-gamma (IFN-γ) was

59

found to synergistically act with TNF-α to induce IL-8 production from a human

gastric cancer cell line, which has been reported to be involved in cancer cell

metastasis (Yasumoto et al., 1992). Table 1.3 summarises the functions of

inflammatory cytokines that might be involved in oral carcinogenesis.

Cytokines Functions

IL-4 Pro-inflammatory, pro-metastasis and accelerates tumour growth from

rhabdomyosarcoma tissue.

IL-6 Pro-inflammatory, neovascularisation, pro-metastasis, B cell activation

and acute phase response.

IL-8 Neo-vascularisation, pro-metastasis, increases cell migration and

activation.

IL-10 Promotes tumour progression during squamous carcinogenesis.

IL-18 Up-regulates inflammatory mediators and innate immune responses.

Up-regulates Th1 response that assists clonal expansion of cytotoxic

T-lymphocytes (CTL) and Th2 response that increases antibody

production.

TNF-α Pro-inflammatory, pro-atherogenic and anti-inflammatory.

Up-regulates inflammatory mediators and innate immune responses.

GM-CSF Pro-inflammatory, anti-inflammatory, pro-metastasiss

Table 1.3 Summary of cytokines that possibly have a role in the progression of OSCC.

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1.8 Hypotheses

The overall hypothesis is that polymicrobial biofilms of C. albicans, A. naeslundii

and S. mutans are involved in oral carcinogenesis. The null hypothesis is that

polymicrobial biofilms of C. albicans, A. naeslundii and S. mutans have no role in

oral carcinogenesis.

The specific hypothesis of the first study is that the auto-aggregation and co-

aggregation of C. albicans is strain-dependent. The null hypothesis is that auto-

aggregation and co-aggregation of C. albicans is not strain-dependent. The hypothesis

of the second study is that polymicrobial biofilm formation is C. albicans strain- and

medium-dependent. The null hypothesis is that polymicrobial biofilm formation is

not C. albicans strain- and medium-dependent. The hypothesis of the third study is

that the polymicrobial interactions within biofilms grown in a flow-cell affects C.

albicans biofilm formation. The null hypothesis is that polymicrobial interactions in

flow-cell biofilms do not affect C. albicans biofilm formation. Finally, the hypothesis

of the fourth study is that oral epithelial cells have an enhanced malignant phenotype

when grown in the presence of polymicrobial biofilm effluent. The null hypothesis is

that the presence of polymicrobial biofilms does not enhance the malignant potential

of oral epithelial cells.

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1.9 Aims

The aim of the aggregation study was to assess the auto-aggregation and co-

aggregation of C. albicans, A. naeslundii and S. mutans during planktonic growth.

The aim of the static biofilm study was to assess the effect of mono- and co-culture of

C. albicans, A. naeslundii and S. mutans on static biofilm formation. The aim of the

flow-cell biofilm study was to assess polymicrobial interactions within biofilms of C.

albicans, A. naeslundii and S. mutans in a flow-cell environment. Finally, the aim of

the adhesion, epithelial-mesenchymal-transition and cytokine studies was to assess

the malignant phenotype of oral epithelial cells when grown in medium containing

biofilm effluent of mono-cultured and polymicrobial C. albicans, A. naeslundii and S.

mutans. The malignant phenotype was measured by adherence to extracellular matrix

molecules, epithelial to mesenchymal transition and cytokine expression. Oral

epithelial cell lines derived from both normal oral epithelium (OKF6) and oral

squamous cell carcinoma (H357) were used to assess the interaction.

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CHAPTER 2

MATERIALS AND METHODS

63

2.1 Growth of microorganisms

C. albicans American Type Culture Collection (ATCC) 32354 (ALT1), ATCC

MYA-2876 (ALT2), ATCC 90234 (ALT3), ATCC 18804 (ALT4), genotype A

isolated from AIDS patient (ALC1), genotype B isolated from AIDS patient (ALC2),

oral cancer isolate 1 (ALC3) and oral cancer isolate 2 (ALC4) were used in this study.

C. albicans strains were sub-cultured on Sabouraud’s dextrose agar (SDA) (Difco,

USA) and incubated at 37 °C aerobically for 24 h.

To grow bacteria, stock cultures of A. naeslundii (NCTC 10301) and S.

mutans (Ingbritt), provided by the Oral Health Cooperative Research Centre,

Melbourne Dental School, The University of Melbourne, were revived by sub-

culturing onto blood agar (40 g L-1 blood agar base and 100 mL L-1 defibrinated horse

blood; Microbiology Medium Preparation Unit, Australia) and Todd-Hewitt yeast

extract (THYE) agar (36.4 g L-1 Todd-Hewitt broth, 8 g L-1 yeast extract and 15 g L-1

Bacto agar), respectively. The agar plates were incubated at 37 °C for 48 h.

2.2 Aggregation assay

A semi-quantitative spectrophotometric assay based on that outlined by

Ledder et al. (2008) and Nagaoka et al. (2008) was used to analyse the aggregation of

the microorganisms. Initially, 24 h cultures of C. albicans grown aerobically in

RPMI-1640 or 25% ASM (0.625 g L-1 type II porcine gastric mucin, 0.5 g L-1

bacteriological peptone, 0.5 g L-1 tryptone, 0.25 g L-1 yeast extract, 0.088 g L-1 NaCl,

0.05 g L-1 KCl, 0.05 g L-1 CaCl2 and 0.25 mg mL-1 haemin, pH 7.0 supplemented with

2.5 mM DTT and 0.5 g L-1 sucrose) to stationary phase were harvested by

centrifugation at 12,000 g for 5 min and washed twice using co-aggregation buffer

(0.1 mM CaCl2, 0.1 mM MgCl2, 150 mM NaCl, 3.1 mM NaN3 dissolved in 1 mM

Tris buffer and adjusted to pH 7.0). The supernatant was discarded and the pellet was

64

re-suspended in co-aggregation buffer. A similar protocol was repeated for S. mutans

and A. naeslundii, except these microorganisms were grown in heart infusion broth

(HIB) to stationary phase.

To determine auto-aggregation, C. albicans, A. naeslundii and S. mutans were

standardised in co-aggregation buffer to give a final cell density of 106 cells mL-1, 107

cells mL-1 and 108 cells mL-1 respectively in separate sterile 2 mL Eppendorf tubes,

equivalent to an optical density of 0.5 at 620 nm wavelength (OD620nm). The number

of cells was enumerated using the colony forming unit (cfu) counting method on SDA

(C. albicans), blood agar (A. naeslundii) and THYE agar (S. mutans) during

optimisation to confirm the cell density at absorbance OD620nm of 0.5. Different cell

densities were chosen to mimic the variability of the microorganisms in the oral

cavity (Lamfon et al., 2005). Each suspension was mixed thoroughly using a vortex

mixer for 30 s and the OD620nm at time (t) = 0 h was measured. The inoculum was

incubated at room temperature for 1 h to allow aggregation and the OD620nm was

recorded. Sterile co-aggregation buffer was used as a standard blank. Percentage

aggregation was calculated using the following equation:

% Auto-aggregation =

([OD620nm (t = 0 h) - OD620nm (t = 1 h)] / OD620nm (t = 0 h)) x 100

Percentage auto-aggregation was calculated for classification of auto-

aggregation; 1) high (more than 40%), 2) intermediate (30% to 40%) and 3) low auto-

aggregation (less than 30%).

A similar protocol was repeated for the study of co-aggregation by inoculating

C. albicans, A. naeslundii or/and S. mutans (inter-kingdom), and A. naeslundii and S.

mutans (intra-kingdom) into a sterile 2 mL Eppendorf tube with the same cell density

65

used for auto-aggregation. The suspension was mixed thoroughly using a vortex

mixer and the OD620nm at t = 0 h recorded. The suspension was incubated at room

temperature for 1 h followed by the measurement of optical density at OD620 nm. The

OD620nm at time (t) = 0 h of dual-culture and tri-culture were 1.0 and 1.5, respectively.

Percentage co-aggregation was assessed using the following equation:

% Co-aggregation = ([OD620nm (t = 0 h) - OD620nm (t = 1 h)] / OD620nm (t = 0 h)) x 100

2.3 Scanning Electron Microscopy (SEM) imaging

The 0 h and 1 h suspensions (100 μL sample) of a selected representative C.

albicans strain, A. naeslundii (NCTC 10301) and S. mutans (Ingbritt), prepared as

above, were transferred onto cover slips and fixed with 1% osmium tetroxide (OsO4)

vapour. The specimens were dehydrated thoroughly in a freeze-drying system,

sputter coated with palladium gold to a thickness of approximately 20 nm and

observed using a scanning electron microscope (XL 30 Series, Philips, Japan).

2.4 Static biofilm formation

A quantitative assay based on that outlined by Yamada et al. (2005) and

Alnuaimi et al. (2013) was used to analyse static biofilm formation by the

microorganisms. To study intra-kingdom biofilms, streak diluted cultures of C.

albicans, A. naeslundii and S. mutans were grown on SDA, blood agar and THYE

agar respectively, for 24 h at 37 °C and several single colonies were resuspended in

RPMI-1640 (Alnuaimi et al., 2013) or 25% ASM, and standardised to give a final cell

density of 106 cells mL-1, 107 cells mL-1 and 108 cells mL-1 respectively in separate

sterile 2 mL Eppendorf tubes, equivalent to an absorbance of 0.5 at 620 nm

wavelength (OD620nm). The suspensions were mixed thoroughly using a vortex mixer

for 30 s. Subsequently, 200 µL of each suspension, resulting in 2 x 105 cells (C.

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albicans), 2 x 106 cells (A. naeslundii) and 2 x 107 cells (S. mutans) of initial

inoculum, was pipetted into each well of a sterile 96-well plate (Nunc, Denmark).

The number of cells mono- and co-cultured was optimised using the cfu counting

method on SDA (C. albicans), blood agar (A. naeslundii) and THYE agar (S. mutans)

to confirm the ratio of the cells in the inocula. Different cell densities were chosen to

mimic the variability of the microorganisms in the oral cavity (Lamfon et al., 2005).

Finally, the 96-well plate was incubated in an orbital shaker at 90 rpm for 72 h at 37

°C (Alyos, Thermo Fisher Scientific, Australia) to mimic the dynamic oral

environment (Alnuaimi et al., 2013). The medium was replenished aseptically every

24 h.

A similar protocol was used to study inter-kingdom biofilm formation by

inoculating C. albicans, A. naeslundii or/and S. mutans into a sterile 2 mL Eppendorf

tube with a similar cell density as in the intra-kingdom assay resulting in 2 x 105 cells

(C. albicans), 2 x 106 cells (A. naeslundii) and 2 x 107 cells (S. mutans) for each

combination per well. The number of cells was confirmed using the cfu counting

method. The suspension was mixed thoroughly using a vortex mixer and 200 µL of

the suspension was pipetted into a sterile 96-well plate. The plate was incubated

aerobically for 72 h at 37 °C in an orbital shaker at 90 rpm and the medium was

replenished aseptically every 24 h. The resultant 72 h biofilm was assessed by the

crystal violet assay (Section 2.6) and the XTT reduction assay (Section 2.7).

2.5 Gram stain

Gram stain was performed on C. albicans ALC3 strain following growth in

RPMI-1640 and 25% ASM for 72 h at 37 °C for the determination of morphology.

Initially, 1 mL of suspension of RPMI-1640 or ASM-grown C. albicans containing 2

x 105 cells was pipetted into each well of a 12-well plate and incubated at 37 °C in an

67

orbital shaker at 90 rpm. The medium was replenished aseptically every 24 h

incubation. Following incubation, the supernatant was discarded and each well was

washed carefully with phosphate buffered saline (PBS) (Sigma-Aldrich, USA) twice

to remove non-adherent cells. Later, Gram staining was performed by adding 1 mL

of methanol to each well for fixation and incubated for 15 min at 25 °C. The

supernatant was then discarded and the plate was air-dried for 45 min. 1 mL of 0.1%

(w/v) crystal violet (CV) solution was added into each well and incubated for a

further 1 min at 25 °C. Subsequently, the plate was washed gently twice under

running distilled water. 1 mL of 70% (v/v) ethanol was pipetted to de-stain for 10 sec

and washed immediately under running water. 1 mL of 1% (w/v) safranin was

pipetted and left for 1 min prior to final washing. The plate was air-dried and

observed under the light microscope (CH Series, Olympus, Australia) (Madigan et al.,

2012). A similar protocol was repeated for A. naeslundii and S. mutans to confirm

each species prior to experiment.

2.6 Crystal violet (CV) assay

Crystal violet (CV) assays were performed according to the protocol outlined

by Alnuaimi et al. (2013). Initially, the biofilm in each well of a 96-well plate was

washed twice with sterile PBS to remove non-adherent cells. 200 µL of methanol was

added to each well for fixation and incubated for 15 min at 25 °C. The supernatant

was then discarded and the plate was air-dried for 45 min. 200 µL of 0.1% (w/v) CV

solution was added into each well and incubated for a further 20 min at 25 °C. The

plate was washed gently twice using running distilled water and 200 µL of 33% (v/v)

acetic acid added to de-stain the biofilm. The plate was incubated for 5 min at room

temperature. A 100 µL aliquot of this solution was transferred to a new sterile 96-

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well plate and the absorbance was measured at OD620nm using a microtiter plate reader

(Victor3, Perkin-Elmer, Australia).

2.7 XTT reduction assay

An XTT reduction assay was performed according to the protocol provided by

the manufacturer (Sigma-Aldrich, USA). Briefly, the biofilm-coated wells were

washed twice with sterile PBS to remove non-adherent cells. Subsequently, 160 µL

of sterile PBS and 40 µL of 4% XTT salt containing 1% phenazine methosulphate

(Sigma-Aldrich, USA) were pipetted into each well to give a final volume of 200 µL.

The plate was incubated at 37 °C for 3 h in the dark. Following incubation, 100 µL of

the suspension was transferred into a new sterile 96-well plate and the absorbance at

OD450nm and OD620nm wavelengths were measured using a microtiter plate reader.

Measurement at the reference wavelength of OD620nm was subtracted from OD450nm to

account for background fluorescence.

2.8 Flow-cell preparation

A single-track flow-cell (40 mm long, 16 mm wide and 2 mm deep) milled

into a high-density polyethylene block was used to examine biofilm formation (Zhu et

al., 2013; Department of Engineering, University of Melbourne, Australia). A

standard-sized 24 mm x 60 mm uncoated glass coverslip (Menzer-Glaser, Germany)

served as the substratum and was secured to the flow-cell using a silicone adhesive

(GE Silicones, General Electric Company, Waterford, NY). Sodium hypochlorite

with 0.5% available chlorine was pumped through the system at a flow rate of 3 mL h-

1 overnight to ensure sterility. Subsequently, overnight rinsing with sterile milliQ

water delivered at the same flow rate was performed to remove the bleach. The flow-

69

cell system was then treated with 25% ASM (Section 2.2) for 2 h at 37 °C to

condition the glass surface with medium prior to inoculation.

2.9 Flow-cell biofilm formation

A quantitative assay based upon that outlined by Zhu et al. (2013) was used to

examine flow-cell biofilm formation. C. albicans ALC3 was chosen for use in the

flow-cell biofilm study as the strain was isolated from a patient with OSCC (Section

2.1). To study intra-kingdom biofilms, C. albicans was grown in 25% ASM, while A.

naeslundii and S. mutans were grown in HIB to stationary phase, washed twice,

resuspended in separate sterile 15 mL tubes in 25% ASM and standardised to give a

final cell density of 106 cells mL-1, 107 cells mL-1 and 108 cells mL-1 respectively. The

suspension was mixed thoroughly using a vortex mixer for 30 s. Subsequently, 1 mL

of this suspension was added with 2 mL of 25% ASM, resulting in 1 x 106 cells (C.

albicans), 1 x 107 cells (A. naeslundii) and 1 x 108 cells (S. mutans), of initial

inoculum which was injected into the system aseptically. These final cell densities

have been chosen to replicate the microbial composition of the previous co-

aggregation (Chapter 3; Arzmi et al., 2015) and static biofilms studies (Chapter 4).

Finally, the system was incubated statically and inverted to allow cells to attach to the

glass substratum for 1 h prior to constant flow (3 mL h-1) of 25% ASM for 23 h to

give a total of 24 h incubation at 37 °C. The experiment was conducted in three

different flow-cells with each flow-cell representing one biological replicate (three in

total).

A similar protocol was used in the study of polymicrobial biofilms formation

by inoculating C. albicans, A. naeslundii and S. mutans into a sterile 15 mL tube at

the same cell density as in the intra-kingdom for each combination per flow-cell. The

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suspension was mixed thoroughly using a vortex mixer and 3 mL of the suspension,

resulting in 1 x 106 cells (C. albicans), 1 x 107 cells (A. naeslundii) and 1 x 108 cells

(S. mutans), was injected into the system aseptically with a subsequent 1 h static

incubation prior to constant flow (3 mL h-1) of 25% ASM for 23 h to give a total of 24

h incubation at 37 °C. The experiment was conducted in three different flow-cells

representing three biological replicates.

The effluent was collected in a sterile 250 mL Schott bottle (Schott, Australia)

on ice, followed by filter sterilisation and storage at -80 °C prior to use.

2.10 Flow-cell gel acrylamide preparation

At the completion of the 24 h incubation period, the biofilms were rinsed in

situ with sterile PBS (Sigma-Aldrich, Australia) to remove non-adherent cells.

Subsequently, the biofilms were fixed with 50% ethanol for 1 h at room temperature

followed by washing with PBS for 30 min. The fixed biofilms were embedded in

20% acrylamide with 0.02% of ammonium persulfate and 0.8% of N,N,N,N-

tetramethylethylenediamine (TEMED), and incubated at room temperature for 30 min

to solidify the gel. The coverslip was then removed and the biofilm embedded in the

polymerised acrylamide slab was stored in PBS at 4 °C prior to fluorescent in situ

hybridisation (FISH) staining.

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Figure 2.1 Flow-cell system.

2.11 Fluorescent In Situ Hybridisation (FISH) staining

An 8 mm2 portion of the polymerised acrylamide slab was excised and placed into

a 4-well Nunclon Surface multidish plate (ThermoFisher Scientific, Australia).

Customised specific probes (Life Technologies, USA) were then added at a final

concentration of 15 μM for C. albicans and S. mutans probes, and 30 μM for A.

naeslundii probes in the presence of 15% formamide in hybridisation buffer (0.9 M

NaCl, 20 mM Tris-HCl, 0.01% SDS adjusted to pH 7.3) (Zainal-Abidin et al., 2012).

Species-specific probe GCC AAG GCT TAT ACT CGC T with the 5’ end labelled

with Alexa Fluor 555 was used for the detection of C. albicans, CGG TTA TCC AGA

AGA AGG GG with the 5’ end labelled with Alexa Fluor 488 was used in the

detection of A. naeslundii and ACT CCA GAC TTT CCT GAC with the 5’ end

labelled with Alexa Fluor 647 was used for S. mutans detection (Life Technologies,

USA).

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2.12 Confocal Laser Scanning Microscopy (CLSM) and image analysis

Fluorescently labelled biofilms were visualised by CLSM (LSM 510 Meta,

Carl Zeiss, Germany) with an inverted stage as described by Dashper et al. (2013) and

Zhu et al. (2013). Horizontal (xy) optodigital sections were taken through the depth

of the biofilm (z) every 2 µm for mono-cultured C. albicans biofilm and every 1 µm

for mono-cultured A. naeslundii, S. mutans and polymicrobial biofilms. Each stack

was imaged using a 63x objective at 512 x 512 pixels, with each frame at 0.28 µm (x)

x 0.28 µm (y). To determine reproducibility, 5 image stacks in random positions

were obtained at wavelengths of 555 nm (C. albicans), 488 nm (A. naeslundii) and

647 nm (S. mutans) for each channel from each of three biological replicates. All

images were analysed with COMSTAT software to determine the biometric

parameters of the biofilms including roughness coefficient, biofilm biomass, average

thickness, maximum thickness and percentage surface colonisation (Heydorn et al.,

2000). Three-dimensional reconstructed images were produced using Zeiss LSM

image browser software (Carl Zeiss, Germany).

Roughness coefficient (Ra) is calculated based on the thickness distribution of

the biofilm that provides a measurement of the variable thickness of the biofilm,

which is an indicator of biofilm heterogeneity (Murga et al., 1995; Heydorn et al.,

2000).

The biofilm biomass is defined as the number of biomass pixels in all images

of a stack multiplied by the voxel size [(pixel size)x x (pixel size)y x (pixel size)z] and

divided by the substratum area of the image stack (Heydorn et al., 2000). The

resulting value is biomass volume divided by substratum area (µm3 µm-2). Bio-

volume represents the overall volume of the biofilm, and also provides an estimate of

the biomass in the biofilm.

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Average biofilm thickness is a measurement of the spatial size of the biofilm

whereas maximum thickness is the thickness over a given location, ignoring pores and

voids inside the biofilm (Heydorn et al., 2000). The percentage of substratum

coverage is the fraction of the area occupied by biomass in each image of a stack.

The substratum coverage is the area coverage in the first image of the stack, which

reflects the efficiency of microbial colonisation on the substratum (Heydorn et al.,

2000).

The percentage proportion of C. albicans, A. naeslundii and S. mutans in

polymicrobial biofilms was calculated using the following equation:

% Proportion =

(Biomass of specific microorganism / Total biomass of microorganisms) x 100

2.13 Cell lines and culture

OKF6, a normal human oral epithelial cell line provided by the Oral Health

Cooperative Research Centre (OHCRC), The University of Melbourne, Australia was

grown in 12 mL of keratinocyte serum-free medium (k-SFM) (Invitrogen, Australia)

containing 25 μg mL-1 bovine pituitary extract and 0.2 ng mL-1 human recombinant

epidermal growth factor 1-53, supplemented with 0.4 mM CaCl2 in a 75 cm2 flask

(Dalley et al., 2014). H357 (Sigma-Aldrich, Australia), a cancerous cell line isolated

from the tongue of a patient with OSCC, was grown in 12 mL of Dulbecco’s

Modified Eagle’s Medium (DMEM)/F12 (Sigma, Australia) containing 10% (v/v)

fetal bovine serum, 0.6 μg mL-1 L-glutamine (Sigma, Australia), and 0.5 μg mL-1

hydrocortisone (Sigma-Aldrich, Australia) in a 75 cm2 flask (Minter et al., 2003).

Both OKF6 and H357 were incubated at 37 °C, 5% CO2 for five to seven days to

74

reach 80% confluence. The medium was replenished every two days for the optimum

growth of the cells. Cells were detached from the flask using 0.25% trypsin/EDTA

(Sigma-Aldrich, Australia) for the adhesion assay, and 10 mM of EDTA in PBS

(Sigma-Aldrich, Australia) for the EMT and Bio-Plex assays, for approximately 5 min

at 37 °C, 5% CO2. The concentration of cells was finally standardised in a prepared

test cell growth medium (Section 2.14) prior to adhesion (Section 2.15), EMT and

cytokine assays (Section 2.16).

2.14 Preparation of test cell growth media

Optimisation of test cell growth media has been conducted at three different

concentrations of serum free medium (SFM) containing biofilm effluent (100% SFM,

80% SFM and 50% SFM). To assess cell viability, 10 μL of the seeded suspension in

a 12-well plate (Corning, NY) as described in Section 2.16, was immediately

collected and mixed with 10 μL of 0.4% trypan blue dye (Bio-Rad, CA) followed by

quantification using a TC10 automated cell counter (Bio-Rad, CA). 80% SFM (v/v)

was selected in the present study based on the optimisation that exhibited

approximately 95% of cells were viable.

To prepare test cell growth media, SFM (DMEM/F12 and k-SFM for H357

and OKF6 respectively) was diluted in the biofilm effluent (Section 2.9) of (1) C.

albicans (ALC3), (2) A. naeslundii (AN), (3) S. mutans (SM) and poly-microbial

(TRI) biofilms, and non-effluent 25% ASM (NE), to give a final concentration of

80% (v/v) SFM (Steele and Fidel, 2002). Subsequently, each solution was mixed

thoroughly and warmed in a water bath at 37 °C for 5 min prior to adhesion, EMT and

Bio-Plex assays.

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2.15 Cell-extracellular matrix (ECM) adhesion assay

Cell-extracellular matrix (ECM) adhesion assays were carried out using the

CytoSelect 48-well Cell Adhesion Assay ECM Array kit (Cell Biolabs, USA). The

kit provides five types of ECM protein (fibronectin, collagen I, collagen IV, laminin

and fibrinogen) coated wells and eight wells of bovine serum albumin coated

substratum that serves as the negative control. The protocol provided by the

manufacturer was followed. Initially, 150 μL of the cell line suspension was

standardised to 1.5 x 104 cells in test cell growth media and added into each well of

the 48-well plate and incubated at 37 °C, 5% CO2 for 90 min. The supernatant was

discarded from each well followed by four times washing with 250 μL of PBS

(Sigma-Aldrich, Australia) containing 2 mM CaCl2 and 2 mM MgCl2. The

suspension was then aspirated and 200 μL of Cell Stain Solution (Cell Biolabs, USA)

was added followed by 10 min incubation at room temperature. The stain solution

was removed and each well was washed four times with 500 μL of sterile milliQ

water. The plate was air-dried at room temperature for 1 h and 200 μL of Extraction

Solution (Cell Biolabs, USA) was added into each well followed by incubation at

room temperature for 10 min on an orbital shaker at 45 rpm (Alyos, Thermo Fisher

Scientific, Australia). Finally, 150 μL of the extracted samples was transferred into a

96-well plate and measured at 570 nm wavelength (OD570nm) using an automated plate

reader (Victor3, Perkin-Elmer, Australia).

Fold change was calculated using the following equation:

Fold change =

Adhesion of cells incubated in biofilm effluent/adhesion of cells in NE.

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2.16 Preparation of cell suspensions for EMT and Bio-Plex assays

To prepare cell suspensions, OKF6 and H357 were standardised in

DMEM/F12 and k-SFM respectively, to a concentration of 1 x 106 cells mL-1. 500 μL

of cell suspension, equivalent to 5 x 105 cells, was seeded into the wells of a 12-well

plate (Corning, NY) followed by incubation at 37 °C, 5% CO2 (Thermo-Fisher,

Australia) for 24 h to reach approximately 80% confluence. The cells were seeded in

triplicate in two different 12-well plates for each cell line. Following incubation, 500

μL of each prepared 80% (v/v) SFM test cell growth medium was added to different

wells and incubated for 2 h and 24 h at 37 °C, 5% CO2 prior to EMT and Bio-Plex

assays.

2.17 EMT assay using flow cytometry

Expression of E-cadherin and vimentin was determined by staining with

fluorescein isothiocyanate (FITC)-conjugated mouse anti-human E-cadherin (67A4)

and phycoerythrin (PE)-conjugated mouse anti-human vimentin (SPM576) (Novus

Biologicals, CO, USA) as per manufacturer’s instructions. Initially, the supernatant

(conditioned media) was collected from wells (Section 2.16) for Bio-plex assays

(Section 2.18) and the cells were incubated with antibodies in FACS buffer (PBS, 2%

BSA, 2 mM EDTA) for 20 min on ice before washing once in PBS by centrifugation

for 5 min at 720 g. The supernatant was removed and the cells were re-suspended in

FACS buffer. Cells were read on a LSR Fortessa X-20 (Becton Dickinson,

Australia). A typical forward and side scatter gate was set to exclude dead cells and

aggregates and a total of 1 x 105 events in the gate were collected. Flow cytometry

data was analysed using FlowJo analysis software (FlowJo, OR, USA). The

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percentages of cells expressing vimentin and E-cadherin, and mean fluorescence

intensity (MFI) have been measured for both OKF6 and H357 cell lines.

Percentage difference of cells expressing EMT markers between 2 h and 24 h

was calculated using the following equation:

% Difference of cells expressing markers =

[(Cell expressing markers at 24 h –cell expressing markers at 2 h) /

cell expressing markers at 2 h] x 100

Whereas, percentage difference of EMT markers expressed by cells between 2

h and 24 h was calculated using the following equation:

% Difference of MFI =

[(MFI at 24 h – MFI at 2 h) / MFI at 2 h] x 100

2.18 Bio-Plex assays

To quantify the amount of cytokines secreted by epithelial cells in response to

biofilm effluent, the conditioned medium was collected and analysed using the Bio-

Plex protein array system and Bio-Rad cytokine multi-plex panel (Bio-Rad). The

method of Bio-Plex analysis was based on Luminex technology and simultaneously

measures IL-2, IL-4, IL-6, IL-8, IL-10, TNF-α, GM-CSF and IFN-γ. In brief, anti-

cytokine/chemokine antibody-conjugated beads were added to individual wells of a

96-well filter plate and adhered using vacuum filtration. The wells were washed and

50 µL of pre-diluted standards or samples were added and the filter plate was adjusted

to shake at 300 rpm. Thereafter, the filter plate was washed and 25 µL of pre-diluted

multiplex biotin-conjugated antibodies were added. After washing, 50 µL of pre-

78

diluted streptavidin-conjugated PE was added, followed by additional washing and

the addition of 125 µL of Bio-Plex buffer to each well. The filter plate was analysed

using the Bio-Plex protein array system and the concentration of each cytokine and

chemokine was determined using Bio-Plex protein array system on a Bio-Plex 200

system (Biorad).

Fold change was calculated using the following equation:

Fold change =

Cytokines expressed by cells incubated in biofilm effluent / cytokines expressed by

cells incubated in NE

Percentage difference of cytokines expressed by cells between 2 h and 24 h

was calculated using the following equation:

% Difference of cytokines expression =

[(Cytokines expressed at 24 h – cytokines expressed at 2 h) / cytokines expressed at 2

h] x 100

For IFN-γ, less than the lowest detectable measure (LLD) was standardised at 0.4 pg

mL-1, as this was the lowest detected cytokine using the Bio-Plex.

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CHAPTER 3

CO-AGGREGATION OF CANDIDA ALBICANS, ACTINOMYCES

NAESLUNDII AND STREPTOCOCCUS MUTANS IS CANDIDA ALBICANS

STRAIN-DEPENDENT

80

3.1 Abstract

Microbial interactions are necessarily associated with the development of

polymicrobial oral biofilms. The aim of this study was to determine the co-

aggregation of eight strains of C. albicans with A. naeslundii and S. mutans. In auto-

aggregation assays, C. albicans strains were grown in either RPMI-1640 or 25%

artificial saliva medium (ASM) whereas bacteria were grown in heart infusion broth

(HIB). C. albicans, A. naeslundii and S. mutans were suspended to give 106 cells mL-

1, 107 cells mL-1 and 108 cells mL-1, respectively, in co-aggregation buffer followed by

a 1 h incubation. The absorbance difference at 620 nm (ΔAbs) between 0 h and 1 h

was recorded. To study co-aggregation, the same protocol was used, except

combinations of microorganisms were incubated together. The mean ΔAbs% of auto-

aggregation of the majority of RPMI-1640-grown C. albicans was higher than in

ASM-grown. Co-aggregation of C. albicans with A. naeslundii and/or S. mutans was

variable among C. albicans strains. Scanning electron microscopy (SEM) images

showed that A. naeslundii and S. mutans co-aggregated with C. albicans in dual- and

tri-culture. In conclusion, the co-aggregation of C. albicans, A. naeslundii and S.

mutans is C. albicans strain-dependent.

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3.2 Introduction

Auto-aggregation is defined as the adherence ability of microorganisms

belonging to the same species (Boris et al., 1997), while co-aggregation is the ability

of genetically distinct microorganisms to adhere to each other (Ledder et al., 2008).

Both auto-aggregation and co-aggregation have been classified as important

mechanisms in the development of oral biofilms and postulated to provide protective

mechanisms to the microbial inhabitants against shear forces that occur within the

oral cavity (Handley et al, 2001). Aggregation contributes to the integration of new

microbial species into biofilms, facilitating the exchange of genes and metabolic

products that in turn supports survival of microorganisms against variable

environmental conditions (Gibbons and Nygaard, 1970; Bos et al., 1996;

Kolenbrander, 2000; Kolenbrander et al., 2002; Rickard et al., 2003; Al-Ahmad et al.,

2007; Ledder et al., 2008).

Furthermore, co-aggregation has been shown to improve the colonisation of

oral epithelial cells by C. albicans, as pre-incubation of buccal epithelial cells with

fimbriated strains of E. coli or Klebsiella pneumoniae increases the adherence and

subsequent attachment of C. albicans (Bagg and Silverwood, 1986). Pre-adherence of

S. sanguinis and S. gordonii to the hard surfaces of the oral cavity provides adhesion

sites for C. albicans, which supports the importance of polymicrobial inter-kingdom

interactions in the oral cavity (Jenkinson et al., 1990; Bamford et al., 2009; Shirtliff et

al., 2009).

The oral microbiome comprises a wide variety of microorganisms such as

yeasts (C. albicans) and bacteria (Actinomyces spp. and Streptococcus spp.). Candida

spp. that belong to kingdom fungi, especially C. albicans, have been found to colonise

approximately 40% to 50% of healthy oral cavities (Manfredi et al., 2013). The

82

number increases in immunocompromised patients with diseases such as AIDS and

diabetes (Grimaudo et al., 1996; Thein et al., 2009). The human oral microbiome is

also comprised of over 600 prevalent taxa at species level although only half of these

have been cultured in the laboratory (Dewhirst et al., 2010). Among the important

oral bacteria, A. naeslundii is an early oral coloniser that can constitute up to 27% of

supragingival dental plaque (Nyvad and Kilian, 1987; Li et al., 2004). The ability of

this species to co-aggregate with other oral microorganisms has been well recognised

(Grimaudo et al., 1996; Li et al., 2001). S. mutans, an acidogenic and aciduric gram-

positive oral bacterium, is widely regarded as a causative agent of dental caries

(Peters et al., 2012).

The majority of in vitro studies of oral microbial co-aggregation have assessed

dual-species oral bacteria interactions (Grimaudo et al., 1996; Cisar et al., 1979; Eke

et al., 1989; Umemoto et al., 1999; Handley et al., 2001; Foster and Kolenbrander,

2004; Shen et al., 2005; Rosen and Sela, 2006; Ledder et al., 2008), and information

of inter-kingdom interactions is limited. Further, as yet, no study utilising ASM for

the growth of C. albicans has been undertaken to assess inter-kingdom co-

aggregation. This is clinically relevant as C. albicans grows as yeast in 25% ASM and

as hyphae in RPMI-1640, and this dimorphism has a role in the virulence of the

species (Arzmi et al., 2012; Arzmi et al., 2014). The yeast form of C. albicans can

adhere to the host cell surfaces by the expression of adhesins, which trigger yeast-to-

hyphae transition, followed by the expression of invasins by the hyphal form that

mediate the uptake of the fungus by the host cell through endocytosis (Kim and

Sudbery, 2011; Gow et al., 2011; Mayer et al., 2013). In addition, research has also

found that S. salivarius strain K12 preferred to co-aggregate to the hyphal region of

C. albicans than the yeast after 3 h incubation in RPMI-1640 during planktonic

83

growth (Ishijima et al., 2012). A similar interaction was also observed between S.

gordonii and C. albicans in which more bacteria co-aggregated at the hyphal region

of the yeast (Bamford et al., 2009).

The aim of the present study was to determine the co-aggregation of C.

albicans, A. naeslundii and S. mutans with the hyphotheses that auto-aggregation and

co-aggregation are C. albicans strain-dependent.

84

3.3 Materials and methods

C. albicans strains were grown on Sabouraud’s dextrose agar (SDA) (Difco,

USA) and incubated at 37 °C aerobically for 24 h, whereas, A. naeslundii (NCTC

10301) and S. mutans (Ingbritt), were revived by sub-culturing onto blood agar

(Difco, USA) and Todd-Hewitt yeast extract agar (Difco, USA), respectively. The

agar plates were incubated at 37 °C for 48 h. To study auto-aggregation and co-

aggregation, a semi-quantitative spectrophotometric assay based on that outlined by

Ledder et al. (2008) and Nagaoka et al. (2008) was used to analyse the aggregation of

the microorganisms (Section 2.2). To verify the co-aggregation of microorganisms,

SEM was conducted for 0 h and 1 h suspensions (Section 2.3). All experiments were

run in triplicate (three biological replicates) with each replicate comprised of three

technical replicates. All data were statistically analysed using SPSS software version

22.0 using independent t-test to compare between the auto-aggregation of C. albicans

in RPMI-1640 and 25% ASM. The analyses were considered statistically significant

when P < 0.05.

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3.4 Results

3.4.1 Morphology of C. albicans in RPMI-1640 and 25% ASM

C. albicans was shown to be predominantly in the hyphal form when grown in

RPMI-1640 medium after 24 h incubation whereas the yeast form was the most

observed in 25% ASM after the same period of incubation (Figure 3.1).

3.4.2 Auto-aggregation

Variation in auto-aggregation of RPMI-1640 grown C. albicans strains

(hyphal growth) was observed with a group of four strains (ALT3, ALT4, ALC1 and

ALC3) exhibiting high auto-aggregation (over 40%), two strains (ALT1 and ALC4)

exhibiting intermediate auto-aggregation (30% to 40%), and two strains (ALT2 and

ALC2) exhibiting low auto-aggregation (Table 3.1; Figure 3.2A). The auto-

aggregation values of A. naeslundii and S. mutans were also classified as low with

11.4% and 7.4%, respectively (Table 3.1).

Four strains of ASM-grown C. albicans (ALT2, ALT3, ALC1 and ALC4)

(yeast growth) exhibiting intermediate auto-aggregation while the remainder strains

(ALT1, ALT4, ALC2 and ALC3) were classified as exhibiting low auto-aggregation

(Table 3.1; Figure 3.2B).

There were four strains of C. albicans that exhibited significantly more auto-

aggregation when grown in RPMI-1640 (hyphal growth) (ALT1, ALT4, ALC1 and

ALC3) compared to 25% ASM (yeast growth) (P < 0.05). Two strains (ALT2 and

ALC2) showed significantly more auto-aggregation when grown in 25% ASM than

RPMI-1640 (P < 0.05) and two strains (ALT3 and ALC4) exhibited no difference in

auto-aggregation regardless of the media type (Figure 3.2).

86

Figure 3.1 Gram-stained of C. albicans cultures observed under light microscopy at 1000x magnification. Left: C. albicans (ALT4) grown in RPMI-1640 after 24 h incubation at 37 °C; >75% of C. albicans cells were present in hyphal form in this medium. Right: C. albicans (ALT4) grown in 25% ASM after 24 h incubation at 37 °C; 100% of C. albicans displaying yeast morphology in this medium.

87

Figure 3.2 Percentage auto-aggregation in RPMI-1640 (A) and 25% ASM (B) grown C. albicans after 1 h incubation in co-aggregation buffer. The study was conducted in three biological replicates with each replicate consisted of three technical replicates. Data were analysed using independent t-test and considered as significantly different when P < 0.05. * indicates significantly more auto-aggregation between the two growth media.

05

101520253035404550

ALT1 ALT2 ALT3 ALT4 ALC1 ALC2 ALC3 ALC4

% A

uto-

aggr

egat

ion

% Auto-aggregation in RPMI-1640 A

* * *

*

0

10

20

30

40

50


% A

uto-

aggr

egat

ion

% Auto-aggregation in ASM B

* *

88

3.4.3 Inter-kingdom co-aggregation

All strains of RPMI-grown C. albicans (hyphal growth) were found to co-

aggregate with A. naeslundii ranging from 9.9 ± 0.5% (ALT3) to 26.2 ± 0.4%

(ALC3). Co-aggregation of RPMI-grown C. albicans with A. naeslundii and S.

mutans were also observed for all strains of the yeast ranging from 2.2 ± 0.3%

(ALT3) to 17.0 ± 0.6% (ALC1). Our study showed that ASM-grown C. albicans

strains (yeast form) co-aggregated with A. naeslundii ranging from 9.6 ± 0.7%

(ALT2) to 23.0 ± 0.1% (ALC3). ASM-grown C. albicans strains were observed to co-

aggregate S. mutans ranging from 9.9 ± 0.2% (ALT3) to 28.1 ± 0.1% (ALT4) (Table

3.1). Co-aggregation of ASM-grown C. albicans with A. naeslundii and S. mutans

were observed in all strains of the yeast ranging from 12.9 ± 0.4% (ALT2) to 25.8 ±

0.5% (ALT1) (Table 3.1).

3.4.4 Scanning Electron Microscopy analyses

SEM analysis of RPMI-grown C. albicans ALT4 strain exhibited auto-

aggregation in co-aggregation buffer after 1 h incubation (Figure 3.3A). Co-

aggregation was observed between C. albicans and A. naeslundii (Figure 3.3B). In

addition, an SEM image also revealed that S. mutans co-aggregated with C. albicans

mostly at the hyphal region of the yeast (Figure 3.3C). The co-aggregation of RPMI-

grown ALT4 C. albicans with A. naeslundii and S. mutans showed that A. naeslundii

and S. mutans were partially aggregating with C. albicans at the hyphal region. A.

naeslundii was also observed to co-aggregate with S. mutans (Figure 3.3D).

SEM analysis showed that ASM-grown C. albicans ALT4 strain (yeast

growth) had auto-aggregation (Figure 3.3E) and A. naeslundii was found to co-

aggregate on the yeast surface after 1 h incubation (Figure 3.3F). Co-incubation of

ALT4 C. albicans with S. mutans revealed that there was inter-kingdom co-

89

aggregation between the two microorganisms with clumps of bacteria attached to the

yeast surface of ALT4 C. albicans (Figure 3.3G). In addition, an SEM image of the

interaction between ASM-grown ALT4 C. albicans with both bacterial species

showed that A. naeslundii and S. mutans co-aggregated on the surface of the yeast.

Finally, the image also revealed that S. mutans cells were co-aggregating with A.

naeslundii after 1 h incubation (Figure 3.3H).

Taken together, the data demonstrate that the auto-aggregation and inter-

kingdom co-aggregation of C. albicans, A. naeslundii and S. mutans are C. albicans

strain-dependent.

90

Table 3.1 Auto and co-aggregation scores of pairs of 8 strains of RPMI-grown (hyphal form) and ASM-grown (yeast form) C. albicans, A. naeslundii and S. mutans. Percentage aggregation as measured by OD620nm change over 1 h (see materials and methods section). Data are means from three separate experiments (SD are given in parenthesis). The study was conducted in three biological replicates with each replicate consisted of three technical replicates. *Auto-aggregation scores representative of interaction between cells from the same culture. # A. naeslundii and S. mutans were grown in BHI respectively.

Strains RPMI-1640 25% ASM Auto-aggregation An Sm An and Sm Auto-aggregation An Sm An and Sm

ALT1 *37.0 (0.2)

24.6 (0.4)

18.2 (0.1)

5.4 (0.1)

*21.5 (0.1)

17.6 (0.2)

17.8 (0.2)

25.8 (0.5)

ALT2 *27.6 (0.4)

17.6 (0.4)

16.4 (0.1)

13.7 (0.3)

*33.3 (0.9)

9.6 (0.7)

24.8 (0.5)

12.9 (0.4)

ALT3 *41.6 (0.4)

9.9 (0.5)

15.4 (0.4)

2.2 (0.3)

*39.7 (0.5)

14.6 (0.4)

9.9 (0.2)

23.3 (0.2)

ALT4 *41.7 (0.5)

17.7 (0.5)

17.3 (0.5)

10.9 (0.1)

*17.9 (0.7)

14.8 (0.1)

28.1 (0.1)

22.0 (0.9)

ALC1 *47.4 (0.3)

18.7 (0.4)

20.0 (0.2)

17.0 (0.6)

*37.3 (0.2)

15.5 (0.2)

10.3 (0.5)

16.7 (0.3)

ALC2 *20.5 (0.3)

19.7 (0.1)

12.3 (0.2)

8.1 (0.2)

*25.1 (0.5)

20.5 (0.3)

10.5 (0.1)

21.9 (0.3)

ALC3 *40.9 (0.5)

26.2 (0.4)

19.5 (0.2)

13.7 (0.3)

*17.2 (0.5)

23.0 (0.1)

19.3 (0.5)

17.3 (0.3)

ALC4 *35.7 (0.6)

18.3 (0.2)

22.7 (0.4)

15.5 (0.2)

*35.7 (0.2)

14.6 (0.5)

22.0 (0.2)

21.7 (0.3)

An# *11.4 (0.7)

9.6 (1.1) *11.4

(0.7) 9.6

(1.1)

Sm# *7.4 (0.6)

9.6 (1.1)

*7.4 (0.6)

9.6 (1.1)

91

A) B) C) D)

E) F) G) H) Figure 3.3 SEM of C. albicans (strain ALT4) auto-aggregation (A & E), inter-kingdom interaction with A. naeslundii (B & F), S. mutans (C & G) and both bacteria (D & H). C. albicans was grown in RPMI-1640 (A, B, C & D) and 25% ASM (E, F, G & H). Magnification is as shown on each image (6500x and 10000x).

92

3.5 Discussion

Co-aggregation is a mechanism that induces the development of a complex

architecture of oral biofilms, which assists the attachment of secondary colonisers

such as S. mutans (Kolenbrander, 2000; Min and Rickard, 2009).

We have shown that inter-kingdom co-aggregation was strain-dependent. The

co-aggregation of the majority of RPMI-grown (hyphal growth) C. albicans strains,

when grown with S. mutans and A. naeslundii either alone or in combination, resulted

in variable co-aggregation. The observed variability of co-aggregation in C. albicans

may be attributable to the different abundances of specific molecules that are

important in adhesion and quorum sensing (eg. Farnesol) from different strains, which

have been suggested to have a role in inter-kingdom interactions of C. albicans and

bacteria (Morales and Hogan, 2010). Furthermore, the variability of co-aggregation

observed in ASM-grown C. albicans (yeast growth) supports our hypothesis that the

co-aggregation of C. albicans to A. naeslundii and S. mutans is highly dependent on

the individual yeast strain.

Variability of co-aggregation was observed when ASM-grown C. albicans

strains were co-incubated with S. mutans and A. naeslundii. This variability suggests

that S. mutans might have induced the formation of binding sites on the yeast surface

that allow the co-aggregation of A. naeslundii to ASM-grown C. albicans when co-

cultured. Previous study has shown that C. albicans that has been co-cultured with S.

mutans increased the binding site of the yeast (Webb et al., 1998a; Calderone et al.,

2000; Falsetta et al., 2014). These results support our hypothesis that co-aggregation

is highly dependent on the C. albicans strain. It cannot be related to the production of

glucan by S. mutans glucosyltransferases as no sucrose was present, however it may

93

be that specific proteins are induced on the surface of C. albicans due to the

interaction with S. mutans that promotes further interaction with A. naeslundii

(Holmes et al., 1995; Koo et al., 2010; Falsetta et al., 2014). Further research is

necessary to assess this hypothetical possibility.

It can be postulated that the observed variability in co-aggregation may be

related to that specific strain’s ability to produce both non-specific (adhesins) and

specific (lectin-saccharide) cell surface receptors (Kolenbrander and Williams, 1981;

McIntire et al., 1982; Rickard et al., 2003; Rosen and Sela, 2006; Ledder et al., 2008).

The specific co-aggregation between C. albicans and A. naeslundii is due to the

presence of mannose-containing adhesin protein on the yeast cell surface (Grimaudo

et al., 1996). This same study also showed variation in the co-aggregation of A.

naeslundii with four different yeast strains, which supports the present study.

However, the study does not include the co-aggregation ability of C. albicans’ hyphae

and the tri-cultured polymicrobial interaction, which has been conducted in the

present study. Furthermore, other research has shown significant strain variation of

the cell wall biogenesis in C. albicans, that may have a role in the observed variation

in aggregation ability (Ragni et al., 2011). Further analysis of the cell wall structure

of a range of C. albicans strains is necessary to fully elucidate the mechanism of this

observed variability.

The sum of auto-aggregation of C. albicans, A. naeslundii and S. mutans

cannot be combined to produce expected co-aggregation value in order to determine

the increase or decrease co-aggregation when co-incubated. This is due to the total

number of cells present in each Eppendorf tube is significantly higher for the

coaggregation studies, which will dramatically alter the dynamics of cell-cell

interactions. Thus, the co-aggregation data has been used to compare the relative

94

coaggregation capabilities of each C. albicans strain.

It has previously been suggested that, due to the limitation of nutrients present

in RPMI-1640, growth in this media induces yeast-hyphae transition leading to

predominant hyphal growth (Urban et al., 2006). Our light microscope images

confirmed this with greater than 75% of C. albicans cells growing in hyphal form in

RPMI-1640. No previous study has assessed the form of growth at SEM level when

C. albicans is grown in 25% ASM. The present study is the first to observe C.

albicans cellular morphology in 25% ASM using SEM which confirmed that in this

media, C. albicans does not grow in hyphal form.

Future assessment of co-aggregation of C. albicans, A. naeslundii and S.

mutans requires animal studies to assess oral biological factors, such as salivary flow

and immunological components that exist in the oral cavity, which may influence

aggregation. These in vivo studies of co-aggregation are likely to enhance our

understanding of the mutual interaction of microorganisms in the oral cavity, a

process likely to be critical in chronic infection and potentially oral carcinogenesis.

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3.6 Conclusion

In conclusion, auto-aggregation and inter-kingdom co-aggregation of C.

albicans have been shown to be strain-dependent and this is likely to be important in

polymicrobial oral biofilm formation.

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CHAPTER 4


ACTINOMYCES NAESLUNDII AND STREPTOCOCCUS MUTANS IS

CANDIDA ALBICANS STRAIN AND MEDIUM DEPENDENT

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4.1 Abstract

Oral biofilms comprise of extracellular polysaccharides and polymicrobial

microorganisms. The aim of this study was to determine the effect of polymicrobial

interactions of C. albicans, A. naeslundii and S. mutans on biofilm formation with the

hypotheses that biofilm biomass and metabolic activity are both C. albicans strain and

medium dependent. To study monospecific biofilms, C. albicans, A. naeslundii and

S. mutans were inoculated into 25% artificial saliva medium (ASM) and RPMI-1640

in separate vials, whereas to study polymicrobial biofilm formation, the inoculum

containing microorganisms was prepared in the same vial prior inoculation into a 96-

well plate followed by 72 h incubation. Finally, biofilm biomass and metabolic

activity were measured using crystal violet (CV) and XTT assays, respectively. Our

results showed variability of mono-cultured and polymicrobial biofilm biomass

between C. albicans strains and medium. Based on cut-offs, out of 32, seven RPMI-

grown biofilms had high biofilm biomass (HBB), whereas, in ASM-grown biofilms,

14 out of 32 were HBB. Of the 32 biofilms, 21 were high metabolic activity (HMA),

whereas in 25% ASM, there was no biofilm had exhibiting HMA. Significant

differences were observed between 25% ASM and RPMI-grown biofilms with respect

to metabolic activity (P < 0.01). In conclusion, biofilm biomass and metabolic

activity were both C. albicans strain and growth medium dependent.

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4.2 Introduction

The oral cavity is a habitat for various microorganisms including yeast and

bacteria (Morales and Hogan, 2010). This oral microbiome provides a balanced oral

environment however perturbation of this homeostasis may lead to the development

of dysbiosis and oral disease (Atanasova and Yilmaz, 2015).

Candida spp., Actinomyces spp. and Streptococcus spp. are common

inhabitants of the human oral cavity (Wade, 2013; O'Donnell et al., 2015;

Kolenbrander et al., 2010). Candida spp. have been found to colonise approximately

50% of healthy human oral cavities (Manfredi et al., 2013). C. albicans is the most

frequently isolated Candida spp. from the oral cavity, especially in

immunocompromised patients with diseases such as AIDS and diabetes (Thein et al.,

2006; Nobile and Johnson, 2015). Actinomyces spp. And Streptococcus spp. are the

normal components of the human oral microbiome, with some species associated with

dental caries initiation and development (Wade, 2013). A. naeslundii is categorised as

an early oral coloniser that can constitute up to 27% of supragingival dental plaque

(Arai et al., 2015; Cheaib et al., 2015; Cavalcanti et al., 2016). Streptococcus mutans

is an acidogenic and aciduric Gram-positive oral bacterium that is widely regarded as

a pathogen that initiates dental caries in association with other oral bacteria (Peters et

al., 2012; Wade, 2013).

Dimorphism is an important virulence factor of C. albicans. It is defined as

the ability of Candida spp. to change morphology between yeast and hyphal forms

(Arzmi et al., 2012; Arzmi et al., 2014). C. albicans is predominantly in the yeast

form during early colonisation of the oral cavity, however, subsequent invasion of

oral epithelial cells is predominantly by the hyphal form. The yeast form of C.

albicans can adhere to host cell surfaces by the expression of adhesins, which trigger

99

yeast-to-hyphae transition, followed by the expression of invasins by the hyphal form

that mediate the uptake of the fungus by the host cell through induced endocytosis

(Gow et al., 2011; Kim and Sudbery, 2011; Mayer et al., 2013).

The majority of in vitro studies of biofilms have been with mono-cultured and

dual-cultured oral microorganisms (Cisar et al., 1979; Eke et al., 1989; Umemoto et

al., 1999; Handley et al., 2001; Foster and Kolenbrander, 2004; Shen et al., 2005;

Ledder et al., 2008; Zhu et al., 2013; Dutton et al., 2014), and information from tri-

culture polymicrobial biofilms remains limited (Zainal-Abidin et al., 2012; Dashper et

al., 2014; Cavalcanti et al., 2016). As yet, no study utilising artificial saliva medium

for the growth of C. albicans has been undertaken to assess polymicrobial biofilms.

This is clinically relevant as C. albicans grows as yeast in 25% ASM and as hyphae in

RPMI-1640.

Crystal violet (CV) and 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-5-

[(penylamino)carbonyl]-2H-tetrazolium hydroxide (XTT) assays are two methods for

biofilm quantification. CV assay measures the microbial biofilm biomass where the

dye interacts with negatively charged molecules present on the surface of the

microorganisms and extracellular polysaccharide (Cheaib et al., 2015). The XTT

assay is a colorimetric-based assay of cell metabolic activity using tetrazolium

hydroxide (McCluskey et al., 2005). Tetrazolium hydroxide is an active compound

that is converted to formazan by the activity of dehydrogenases involved in the

metabolic pathways of microbial cells (Peeters et al., 2008). Succinate

dehydrogenases of prokaryotic cells and mitochondrial dehydrogenases of eukaryotic

cells are examples of dehydrogenase activity that can be detected by XTT

(McCluskey et al., 2005; Moffa et al., 2016).

100

The aims of the present study were to determine the effect of interactions of C.

albicans, A. naeslundii and S. mutans on the formation of polymicrobial biofilms and

to assess this interaction when biofilms were grown in 25% ASM for predominantly

yeast growth and in RPMI-1640 for predominantly hyphal growth. We hypothesized

that this polymicrobial biofilm formation is C. albicans strain- and growth medium-

dependent.

101


A quantitative assay based on that outlined by Yamada et al. (2005) and

Alnuaimi et al. (2013) was used to analyse static intra-kingdom and inter-kingdom

biofilms formation by C. albicans, A. naeslundii and S. mutans (Section 2.4). Gram

staining was performed on C. albicans ALC3 strain for the determination of

morphology (Section 2.5). Crystal violet (CV) assay was performed according to the

protocol outlined by Alnuaimi et al. (2013) to assess biofilm biomass (Section 2.6)

and XTT reduction assay was performed according to the protocol provided by the

manufacturer (Sigma-Aldrich, USA) to assess metabolic activity of biofilm (Section

2.7).

All biofilms containing C. albicans were divided into terciles according to

biofilm biomass and metabolic activity for CV and XTT assays, respectively. This

division provided the cut-offs to classify strains as high, moderate and low biofilm

biomass (HBB, MBB and LBB); and high, moderate and low metabolic activity

(HMA, MMA and LMA) (Marcos-Zambrano et al., 2014).

All experiments were run in triplicate (three biological replicates) with each

replicate comprised of three technical replicates. Using SPSS software version 22.0,

all data were statistically analysed by applying chi-square test to compare between the

categories (high, medium and low) for each assay. Since comparison was made

between independent groups that were not normally distributed, thus, non-parametric

chi-square was chosen for the statistical analysis. Meanwhile, to compare between

ATCC and clinical strains biofilm biomass, two-tailed independent t-test analysis was

conducted as the data were normally distributed. Comparison between a group of

ATCC isolates (ALT1, ALT2, ALT3 and ALT4) and a group of clinical isolates

(ALC1, ALC2, ALC3, ALC4) of C. albicans was analysed using two-tailed

102

independent t-test since the data were normally distributed. Finally, multiple

comparisons between mono-cultured with polymicrobial biofilms were analysed using

ANOVA post hoc Tukey test since the equal variances were assumed as analysed

using Levene’s test.

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4.4 Results

4.4.1 Morphology of C. albicans bioflms in RPMI-1640 and 25% ASM

C. albicans biofilm growth was predominantly in the hyphal form when grown

in RPMI-1640, and in the yeast form when grown in 25% ASM after 24 h incubation

as observed by Gram staining (Figure 4.1).

4.4.2 Effect of microbial interaction and medium on biofilm biomass

Biofilm biomass was categorised into terciles using the following CV

measurement cut-offs: LBB < 2.280, MBB 2.280-2.535, HBB > 2.535. None of

mono-cultured C. albicans was categorised as HBB, however, when co-cultured with

A. naeslundii three C. albicans strains (ALT1, ALT2 and ALT3) were categorised as

HBB (Table 4.1). Only ALT1 was categorised as HBB when co-cultured with S.

mutans in RPMI-1640 whereas in tri-cultured biofilms, three strains of C. albicans

(ALT1, ALT2 and ALT3) were categorised HBB (Table 4.1).

None of mono-cultured ASM-grown C. albicans exhibited HBB, however, in

the presence of A. naeslundii, seven strains of C. albicans were classified as HBB

(Table 4.1). Interaction of C. albicans with S. mutans showed that two strains (ALT1

and ALT3) were HBB, while in tri-cultured biofilms, five C. albicans strains (ALT1,

ALT2, ALT3, ALT4 and ALC2) were classified as HBB (Table 4.1).

Analyses of all 32 biofilms for biomass showed that there were seven biofilms

classified as HBB (21.9%), 12 MBB (37.5%) and 13 LBB (40.6%) when the biofilms

were grown in RPMI-1640 (hyphal growth). Biofilms grown in 25% ASM (yeast

form) showed 14 biofilms categorised as HBB (43.8%), ten MBB (31.3%) and eight

LBB (25.0%). There were more biofilms with HBB when grown in 25% ASM (yeast

form) than RPMI-1640 (hyphal form), however, this did not reach statistical

significance (P > 0.05).

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Five RPMI-grown biofilms (hyphal form) had significantly increased biomass

when C. albicans strains were co-cultured with A. naeslundii (ATCC: ALT1, ALT4;

Clinical: ALC1, ALC2 and ALC4) compared with mono-cultured C. albicans biofilm

(P < 0.05). Further, co-culture of C. albicans with S. mutans increased biomass of six

biofilms (ATCC: ALT1, ALT4; Clinical: ALC1, ALC2, ALC3 and ALC4)

significantly (P < 0.05). Five biofilms (ATCC: ALT1, ALT4; Clinical: ALC1, ALC2

and ALC4) increased biomass significantly when C. albicans was co-cultured with

both A. naeslundii and S. mutans when compared with the mono-cultured biofilm of

C. albicans (P < 0.05; Table 4.1).

Two ASM-grown biofilm (ATCC: ALT1 and ALT2; yeast form) had a

significantly increased biomass when C. albicans was co-cultured with A. naeslundii

compared with the mono-cultured C. albicans biofilm (P < 0.05). One biofilm

(ATCC: ALT1) showed a significant increase (P < 0.05) and one (ATCC: ALT2) a

significant decrease (P < 0.05) in biomass when C. albicans was co-cultured with S.

mutans. There was one strain (ATCC: ALT1) that showed a significant increase in

biomass when C. albicans was co-cultured with both A. naeslundii and S. mutans

compared with mono-cultured C. albicans biofilm (P < 0.05; Table 4.1).

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A) B) Figure 4.1 Gram-stained biofilms of C. albicans strain ALC3 observed under light microscope at 200x magnification after 72 h incubation at 37 °C in 24-well plate at 90 rpm. A: ASM-grown C. albicans biofilm; B: RPMI-grown C. albicans biofilm.

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Table 4.1 Biofilm biomass scores of eight strains of RPMI-grown and ASM-grown C. albicans, A. naeslundii (An) and S. mutans (Sm) as measured by OD620nm after 72 h incubation. Data are means from three biological replicates with each replicate consisted of three technical replicates (SD are given in parenthesis). Significant difference (P < 0.05) observed between dual-cultured C. albicans-An (*), C. albicans-Sm (#) or tri-cultured (I) to mono-cultured C. albicans biofilms grown in the same medium.

Strains RPMI-1640 25% ASM Mono An Sm An and Sm Mono An Sm An and Sm

ALT1 2.389 (0.019)

*2.681 (0.016)

#2.541 (0.015)

I2.683 0.042)

2.326 (0.078)

*2.625 (0.010)

#2.572 (0.038)

I2.630 (0.033)

ALT2 2.501 (0.326)

2.658 (0.094)

2.504 (0.063)

2.697 (0.010)

2.533 (0.061)

*2.621 (0.040)

#2.454 (0.089)

I2.610 (0.008)

ALT3 2.408 (0.211)

2.666 (0.064)

2.353 (0.145)

2.684 (0.025)

2.455 (0.212)

2.611 (0.033)

2.569 (0.035)

2.625 (0.011)

ALT4 1.554 (0.170)

*2.492 (0.129)

#2.370 (0.155)

I2.457 (0.064)

2.330 (0.185)

2.455 (0.224)

2.015 (0.161)

2.540 (0.101)

ALC1 1.762 (0.205)

*2.259 (0.267)

#2.401 (0.017)

I2.315 (0.226)

2.253 (0.131)

2.568 (0.117)

1.994 (0.171)

2.452 (0.167)

ALC2 1.594 (0.083)

*2.268 (0.410)

#2.348 (0.128)

I2.061 (0.064)

2.522 (0.090)

2.624 (0.019)

2.284 (0.266)

2.604 (0.040)

ALC3 0.722 (0.270)

1.722 (0.359)

#2.215 (0.205)

1.514 (0.339)

1.937 (0.916)

2.537 (0.133)

1.531 (0.318)

2.275 (0.251)

ALC4 1.445 (0.127)

*1.965 (0.046)

#2.348 (0.128)

I1.952 (0.344)

2.041 (0.551)

2.553 (0.101)

1.676 (0.186)

2.369 (0.220)

An 0.073 (0.002)

0.149 (0.006) 1.711

(0.086) 1.197

(0.066)

Sm 0.066 (0.001)

0.149 (0.006)

0.921 (0.078)

1.197 (0.066)

Low biofilm biomass (LBB) Moderate biofilm biomass (MBB) High biofilm biomass (HBB)

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4.4.3 Effect of microbial interaction and medium on metabolic activity

Biofilm metabolic activity based on the XTT assay was divided into terciles

and categorised based on the following cut-offs: LMA < 0.120, MMA 0.120-0.550,

HMA > 0.550. RPMI-1640 mono-cultured growth resulted in six strains of C.

albicans (ALT1, ALT2, ALT4, ALC1, ALC2 and ALC4) categorised with HMA

(Table 4.2). Seven C. albicans strains (ALT1, ALT2, ALT3, ALT4, ALC1, ALC2

and ALC4) when co-cultured with A. naeslundii in RPMI-1640 had HMA. Only two

strains of C. albicans (ALT1 and ALT2) had HMA when co-cultured with S. mutans

in RPMI-1640. Six C. albicans strains (ALT1, ALT2, ALT3, ALT4, ALC2 and

ALC4) were categorised as having HMA when co-cultured in RPMI-1640 with both

A. naeslundii and S. mutans (Table 4.2).

25% ASM mono-cultured growth resulted in all C. albicans strains being

categorised with LMA (Table 4.2), however, in the presence of A. naeslundii, all

strains had MMA. Interaction of C. albicans with S. mutans showed that all C.

albicans strains remained with LMA whereas, in the presence of both A. naeslundii

and S. mutans, there were three strains having MMA (ALT1, ALT4 and ALC2) and

five strains with LMA (ALT2, ALT3, ALC1, ALC3 and ALC4) (Table 4.2).

Analyses of all 32 biofilms showed that there were 21 biofilms of RPMI-

grown biofilms (hyphal growth) categorised as having HMA (65.6%) and 11 with

MMA (34.4%). In addition, there were 11 ASM-grown biofilms (yeast growth)

categorised as having MMA (34.4%) and 21 with LMA (65.6%). Thus, statistically

significant higher metabolic activity was observed when biofilms were grown in

RPMI-1640 (P < 0.01).

Only C. albicans strains ALT3 when co-cultured with A. naeslundii showed

an increased activity when grown in RPMI-1640 when compared with mono-cultured

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C. albicans. Furthermore, there were four C. albicans strains (ALT4, ALC1, ALC2

and ALC4) that exhibited a decrease in metabolic activity when co-incubated with S.

mutans compared with the mono-cultured biofilm of C. albicans. There was only one

biofilm (ALC1) that showed decreased bioactivity when C. albicans was co-cultured

with both A. naeslundii and S. mutans compared with mono-cultured C. albicans

(Table 4.2).

Three RPMI-grown biofilms (ATCC: ALT3; Clinical: ALC2 and ALC4;

hyphal form) exhibited significant increase activity when C. albicans was co-cultured

with A. naeslundii in comparison with the mono-cultured C. albicans biofilm (P <

0.05). Four biofilms (ATCC: ALT4; Clinical: ALC1, ALC2 and ALC4) showed

significant decrease metabolic activity when C. albicans was co-cultured with S.

mutans. Whereas, one biofilm (Clinical: ALC1) displayed a significant decrease

activity when C. albicans was co-cultured with both A. naeslundii and S. mutans

when compared with mono-cultured C. albicans (P < 0.05; Table 4.2).

Finally, based on metabolic activity per unit biomass in mono-cultured

biofilms, ALT4 and ALC3 were found to be the most active C. albicans strains when

grown in 25% ASM and ALT2 was the least active when grown in the same medium.

Whereas, in RPMI-1640, ALC3 was found to be the most active while ALT3 was the

least (Table 4.3).

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Table 4.2 Static biofilm metabolic activity scores of eight strains of RPMI-grown and ASM-grown C. albicans, A. naeslundii (An) and S. mutans (Sm) as measured by OD450nm-620nm after 72 h incubation. Data are means from three biological replicates with each replicate consisted of three technical replicates (SD are given in parenthesis). Significant difference (P < 0.05) observed between dual-cultured C. albicans-An (*), C. albicans-Sm (#) or tri-cultured (I) to mono-cultured C. albicans biofilms grown in the same medium

Strains RPMI-1640 25% ASM Mono An Sm An and Sm Mono An Sm An and Sm

ALT1 0.780 (0.022)

0.819 (0.021)

0.775 (0.050)

0.782 (0.050)

0.018 (0.000)

*0.398 (0.067)

#0.055 (0.010)

I0.121 (0.022)

ALT2 0.788 (0.012)

0.811 (0.029)

0.813 (0.021)

0.805 (0.020)

0.014 (0.002)

*0.297 (0.083)

0.016 (0.101)

I0.066 (0.018)

ALT3 0.395 (0.074)

*0.610 (0.120)

0.405 (0.028)

I0.617 (0.055)

0.016 (.004)

*0.487 (0.095)

0.027 (0.008)

I0.084 (0.032)

ALT4 0.738 (0.130)

0.665 (0.035)

#0.492 (0.012)

0.595 (0.033)

0.117 (0.028)

*0.549 (0.093)

#0.034 (0.002)

0.171 (0.102)

ALC1 0.641 (0.033)

0.645 (0.058)

#0.395 (0.018)

I0.509 (0.047)

0.024 (0.003)

*0.455 (0.028)

0.021 (0.002)

I0.062 (0.016)

ALC2 0.610 (0.034)

*0.726 (0.003)

#0.507 (0.003)

0.603 (0.034)

0.026 (0.000)

*0.335 (0.036)

0.045 (0.030)

I0.121 (0.053)

ALC3 0.525 (0.030)

0.522 (0.004)

0.394 (0.053)

0.510 (0.022)

0.080 (0.001)

*0.297 (0.045)

#0.027 (0.009)

0.076 (0.023)

ALC4 0.557 (0.024)

*0.638 (0.019)

#0.433 (0.024)

0.630 (0.078)

0.042 (0.000)

*0.344 (0.026)

#0.021 (0.005)

0.057 (0.023)

An 0.050 (0.002)

0.002 (0.001) 0.147

(0.028) 0.011

(0.002)

Sm 0.001 (0.001)

0.002 (0.001)

0.002 (0.002)

0.011 (0.002)

Low metabolic activity (LMA) Moderate metabolic activity (MMA) High metabolic activity (HMA)

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Table 4.3 Mono-culture metabolic activity per biofilm biomass (XTT/CV) scores of 8 strains of RPMI-grown (hyphal form) and ASM-grown (yeast form) C. albicans, A. naeslundii (An) and S. mutans (Sm). Data are means from three biological replicates with each replicate consisted of three technical replicates (SD are given in parenthesis).

Media ALT1 ALT2 ALT3 ALT4 ALC1 ALC2 ALC3 ALC4 An Sm RPMI-1640 0.326

(0.007) 0.319

(0.046) 0.166

(0.044) 0.476

(0.075) 0.366

(0.027) 0.382

(0.009) 0.807

(0.221) 0.388

(0.044) 0.681

(0.035) 0.022

(0.016) 25% ASM 0.008

(0.000) 0.005

(0.001) 0.007

(0.002) 0.051

(0.016) 0.011

(0.002) 0.010

(0.000) 0.051

(0.032) 0.022

(0.006) 0.087

(0.019) 0.003

(0.002)

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4.5 Discussion

To our knowledge, this is the first study to evaluate the effect of microbial

interactions of yeast growth and hyphal growth of C. albicans, A. naeslundii and S.

mutans on the formation of static biofilms in vitro. The results of the present study

clearly demonstrate that both biofilm biomass and metabolic activity are C. albicans

strain and growth medium dependent.

The present study has shown a variation of biofilm biomass and metabolic

activity between strains of C. albicans. Overall, when grown as mono-cultured the

majority of clinical strains had a significantly lower biofilm biomass than the ATCC

reference strains when grown in RPMI (hyphal form). However, a significant

increase of biomass was observed in all clinical strains that did not occur in ATCC

strains (ALT2 and ALT3) when grown in polymicrobial biofilms. Furthermore,

biofilms that were formed by clinical isolates of C. albicans have shown lower

biofilm biomass compared with the reference strains C. albicans (Alnuaimi et al.,

2013). Furthermore, the metabolic activity has been shown to vary among C. albicans

strains; however, the morphology of C. albicans in this previous study was unknown

(Alnuaimi et al., 2013). Strain variability of C. albicans has been shown in the oral

cavity of different individuals (Hellstein et al., 1993; Kleinegger et al., 1996). In

addition, C. albicans strains isolated from HIV-infected patients produce higher levels

of aspartic proteinases (SAPs), compared with strains isolated from uninfected

patients (de Bernardis et al., 1996). SAP is a putative virulence factor that is able to

affect C. albicans biofilm formation in the oral cavity together with phenotypic

switching, morphogenesis and quorum sensing (Morales and Hogan, 2010; Arzmi et

al., 2012). Thus, the results from the present study may indicate a symbiotic

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interaction between clinical C. albicans and oral microorganisms that may lead to the

increase of colonisation in the oral cavity of diseased patients.

The metabolic activity of biofilms was shown to be growth media dependent,

with the majority of ASM-grown C. albicans biofilms having lower metabolic activity

than those grown in RPMI-1640, particularly mono-cultured biofilms (Table 4.2,

Table 4.3). It is postulated that RPMI-1640, which contains limited nutrients, induces

stress in C. albicans, thus promoting hyphal formation. This does not occur when the

yeast is grown in 25% ASM that is rich in nutrients. Interestingly, previous studies

based on the growth rate have shown that C. albicans with low metabolic activity are

more invasive and associated with disease, while conversely those with high activity

are non-invasive (Baillie and Douglas, 1998; Silva et al., 2011; Tobudic et al., 2012).

Furthermore, low metabolic activity has been shown to reduce the antifungal

susceptibility of C. albicans within the biofilm, which could be due to minimal

absorption of antifungal agents such as amphotericin B, thus affecting inactivation

kinetics (Mah and O' Toole, 2001).

The metabolic activity of all C. albicans strains that were grown in 25% ASM

increased in the presence of A. naeslundii in dual-cultured biofilms. However, a

decrease of metabolic activity was observed in tri-cultured biofilms when compared

to the dual-cultured biofilms of C. albicans and A. naeslundii, suggesting that these

microorganisms may be interacting metabolically. It is postulated that in the presence

of A. naeslundii, C. albicans may increase mitochondrial dehydrogenase activity that

in turn, increased the activity of succinate dehydrogenases of A. naeslundii. In

addition, S. mutans has been shown to reduce the metabolic activity in tri-cultured

biofilms compared with the dual-cultured C. albicans-A. naeslundii biofilms,

suggesting that the antagonistic metabolic interaction between A. naeslundii and S.

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mutans, demonstrated in the present study (Table 4.2), may have affected overall

metabolic activity of the consortia. C. albicans and A. naeslundii have been shown to

synthesize mitochondrial and succinate dehydrogenases, respectively, that were

reported to be detectable by XTT (McCluskey et al., 2005; Moffa et al., 2016). Even

though S. mutans has been found to synthesize an NADH-dependent lactate

dehydrogenase; the present study revealed that enzyme activity was not detected with

XTT suggesting that the assay is not suitable for the study of S. mutans metabolic

activity. Furthermore, it is also postulated that the decrease of metabolic activity in

tri-cultured biofilms compared to dual-cultured could be due to the severe nutrient

limitation. The metabolic activity may be higher at earlier time points; however,

several aspects need to be considered such as the strain and morphology of

C.albicans. Even though there are more biofilms when grown as mono-cultured,

however, the competition of nutrient in co-cultured may have induced C. albicans to

produce hyphae, which require more energy. Since the present study was measuring

XTT at on 72 h, therefore, further research to determine the metabolic activity at

different time point including 24 h is highly recommended.

In the present study, the biofilm biomass was shown to vary with microbial

interactions (mono-cultured C. albicans, dual-cultured C. albicans and A. naeslundii,

dual-cultured C. albicans and S. mutans, tri-cultured C. albicans, A. naeslundii and S.

mutans). The majority of RPMI-1640 grown C. albicans (hyphal form) biofilm

biomass were observed to increase in the presence of bacteria compared with mono-

cultured C. albicans. A. naeslundii and S. mutans have been shown to bind to C.

albicans through its mannose-containing surface protein (Rickard et al., 2003; Ledder

et al., 2008; Dutton et al., 2014; Falsetta et al., 2014; Sztajer et al., 2014; Nobile and

Johnson, 2015). This interaction has been reported to induce the formation of

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extracellular polysaccharide, thus promoting the adherence of the late colonisers to

form a complex polymicrobial biofilm potentially enhancing biofilm biomass (Wade,

2013; Cheaib et al., 2015; Nobile and Johnson, 2015; Cavalcanti et al., 2016)..

The present study found that the ATCC strains form excellent mono-cultured

biofilms in both 25% ASM and RPMI-1640 such that addition of A. naeslundii or S.

mutans resulted in no additional biomass in the majority of biofilms. However, the

clinical strains that were poor biofilm formers in RPMI-1640 were observed to

increase biofilm biomass significantly when A. naeslundii or S. mutans was co-

inoculated (Table 4.1). This result indicates that the choice of isolates in the study of

the interaction between oral yeast and oral bacteria in biofilms is critical. The C.

albicans ATCC strains assessed in the present study would appear to have lost either

the ability, or need, to interact with oral bacteria (Harriott and Noverr, 2011), thus

investigations using only ATCC strains of C. albicans are likely to not reflect the true

interactions that are occurring in the oral cavity.

We have demonstrated that C. albicans predominantly in the yeast form when

grown as a biofilm in 25% ASM, whereas, RPMI-grown C. albicans biofilms were

predominated by the hyphal form (Figure 4.1). These results support previous work

that showed the proportion of yeast and hyphal cells of C. albicans present in the

biofilm is dependent upon the nutrient source, where nitrogen-based medium allowed

for more yeast growth and biofilms grown in RPMI-1640 with high salts, amino acids

and D-glucose, showed more hyphal growth (Chandra et al., 2001).

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4.6 Conclusion

Biofilm biomass and metabolic activity have been shown to be both C.

albicans strain and medium dependent. This is likely to have significance in the

development of polymicrobial oral biofilms in vivo.

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CHAPTER 5


ACTINOMYCES NAESLUNDII AND STREPTOCOCCUS MUTANS IN A

FLOW ENVIRONMENT

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5.1 Abstract

C. albicans, A. naeslundii and S. mutans have been shown to exist as

polymicrobial biofilms in the oral cavity. The aim of this study was to determine the

effect of polymicrobial interactions of OSCC-isolated C. albicans (ALC3), A.

naeslundii and S. mutans on biofilm formation in a flow environment. To study

mono-cultured biofilm formation, C. albicans, A. naeslundii and S. mutans were

inoculated in 25% artificial saliva medium (ASM), and standardised to a final density

of 106 cells mL-1, 107 cells mL-1 and 108 cells mL-1 respectively in separate 15 mL

tubes. Cell suspensions (3 mL) were inoculated into a flow-cell system prior to

commencement of a constant medium flow rate of 3 mL h-1 for 24 h at 37 °C. To

study polymicrobial biofilm formation, the same protocol was repeated, except that

the inoculum that was standardised to the same cell density as used in the mono-

cultured biofilm assay, was prepared in the same vial. The biofilms were fixed with

50% ethanol, embedded in 20% gel acrylamide, stained by fluorescent in situ

hybridisation (FISH) using DNA species-specific probes, imaged using confocal

scanning laser microscopy (CSLM) and analysed using COMSTAT software. The

biomass of C. albicans and S. mutans in polymicrobial biofilms exhibited significant

decreases (P < 0.05) compared to mono-cultured biofilms. The roughness coefficient

of polymicrobial biofilms exhibited a significant increase compared to mono-cultured

C. albicans (P < 0.05), however, a significant decrease was observed when compared

to mono-cultured A. naeslundii (P < 0.05). Significant increases of average thickness

and maximum thickness of polymicrobial biofilms were observed when compared to

mono-cultured C. albicans (P < 0.05) and A. naeslundii (P < 0.05), however,

significant decreases of the parameters were observed in polymicrobial biofilms when

compared to mono-cultured S. mutans biofilm (P < 0.05). A significant increase of

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surface colonisation was observed in polymicrobial biofilms when compared to

mono-cultured A. naeslundii (P < 0.05) and S. mutans biofilms (P < 0.05), however, a

significant decrease was observed when compared to mono-cultured C. albicans

biofilm. In conclusion, C. albicans, A. naeslundii and S. mutans formed

polymicrobial biofilms. The inclusion of A. naeslundii in these biofilms resulted in a

decrease in both C. albicans and S. mutans. This may mean that A. naeslundii can be

potentially used as a probiotic to control C. albicans and S. mutans colonisation.

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5.2 Introduction

Oral microorganisms, including C. albicans, A. naeslundii and S. mutans, have

been shown to exist as components of complex polymicrobial biofilms in the oral

cavity (Kolenbrander et al., 2002; El-Azizi et al., 2004; Foster and Kolenbrander,

2004; Kolenbrander et al., 2010). However, slight changes in the microenvironment

such as microbial interactions, nutrient supply and shear forces may affect the

dynamic structure of polymicrobial biofilms (Morales and Hogan, 2010;

Kolenbrander et al., 2010; Diaz et al., 2012; Zhu et al. 2013; Marsh et al., 2016).

Synergies and antagonisms between microorganisms such as the interactions between

C. albicans, A. naeslundii and S. mutans have been previously shown during

planktonic growth (Chapter 3) that have been suggested to affect the dynamic

structure of oral biofilms (Arzmi et al., 2015). Furthermore, the different nutrient

composition of the medium used to grow polymicrobial biofilms, such as 25% ASM

and RPMI-1640, was shown to affect C. albicans morphology, biomass and metabolic

activity in static biofilms (Chapter 4). In addition, mucin containing ASM has been

shown to provide binding sites for the attachment of early colonisers to the substratum

(de Repentigny et al., 2000; Derrien et al., 2010) whereas sucrose containing media

has been reported to induce the synthesis of glucosyltransferases (Gtfs) from S.

mutans that assist in the formation of polymicrobial biofilms (Falsetta et al., 2014).

C. albicans, A. naeslundii and S. mutans are important members of the oral

microbiome (Nobbs and Jenkinson, 2015; Höfs et al., 2016). C. albicans is the most

prevalent opportunistic and pathogenic fungus that can cause oral candidosis (Kim

and Sudbery, 2011). The yeast has also been found to associate with leukoplakic

lesions and is recognised as an independent risk factor for oral carcinoma (Cawson,

1969a). Transition of yeast to hyphae is usually related to the ability of C. albicans to

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colonise oral surfaces. Yeast cells are predominantly found to colonise the surface of

oral substrata, whereas hyphal cells are frequently found during invasive colonisation

of mucosal cells (Finkel and Mitchell, 2011; Banerjee et al., 2013). A. naeslundii has

been categorised among the pioneer colonisers that may constitute up to 27% of

supragingival dental plaque (Nyvad and Kilian, 1987; Li et al., 2004), whereas S.

mutans is a Gram-positive, facultative anaerobic bacterium that utilises a broad

spectrum of sugars and excretes organic acids that leads to the increase of acidity in

plaque inducing dental caries (Takahashi and Nyvad, 2011; Burne et al., 2012). S.

mutans is also known as one of the most important members of the oral microbiome

that supports the structure of mature oral biofilms (Sztajer et al., 2014).

The microbial balance in the oral cavity can be disrupted by various factors

including a high carbohydrate diet, which can lead to C. albicans infection (Williams

et al. 2011). Thus, a balance has to be maintained in order to limit colonisation and

proliferation of opportunistic pathogens or pathobionts in the oral cavity. The idea of

microbial homeostasis led to the discovery of prebiotics, which are nutritional

supplements that beneficially affect the host by improving the microbial balance of

the intestine (Fuller, 1989). Later, probiotics were discovered which have been

defined as live microorganisms that provide health benefits to the host when

administered in adequate amounts (Salminen et al., 1998). Even though probiotics

have been suggested to provide benefits to human health, side effects may be

associated when bacteria are consumed.

According to a 2002 report released by the World Health Organisation

(WHO), there are four types of side effects including systemic infections, deleterious

metabolic activities, excessive immune stimulation in susceptible individuals and

gene transfer (Doron and Snydman, 2015). The use of a probiotic may in some ways

121

be beneficial in changing the flora but this in turn may result in an imbalance of the

normal microbiome and lead to the colonisation of opportunistic microorganisms

(dysbiosis). Even though infection caused by the consumption of probiotics is rare,

septicaemia and endocarditis caused by Lactobicilli spp. has been reported with the

majority of infection cases due to the patient’s normal microbiome (Marteau and

Shanahan, 2003). Thus, the use of any probiotic must be rigorously assessed prior to

use and consequently stringently controlled.

In 1995 synbiotics were discovered; they are a mixture of probiotics and

prebiotics that provide benefits to the host by improving the survival and implantation

of dietary supplements containing live microbes, by selectively stimulating the growth

and/or by activating one or a limited number of health-promoting bacteria (Lilly and

Stillwell, 1965; Kojima et al., 2016).

The aim of the present study was to determine the effect of polymicrobial

interactions of C. albicans, A. naeslundii and S. mutans on biofilm formation in a flow

environment; with the hypotheses that C. albicans, A. naeslundii and S. mutans form

polymicrobial biofilms and that polymicrobial interactions of C. albicans, A.

naeslundii and S. mutans affect colonisation of oral microorganisms.

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C. albicans isolated from an oral cancer patient (ALC3) was sub-cultured on

Sabouraud’s dextrose agar (SDA) (Difco, USA) and incubated at 37 °C aerobically

for 24 h. Stock cultures of A. naeslundii (NCTC 10301) and S. mutans (Ingbritt),

were revived by sub-culturing onto blood agar (Difco, USA) and Todd-Hewitt yeast

extract agar (Difco, USA) respectively. The agar plates were incubated at 37 °C for

48 h (Section 2.1). Following that, biofilms were developed in a flow-cell system for

24 h at 37 °C (Section 2.8; Section 2.9), embedded in gel acrylamide (Section 2.10),

labelled using Fluorescence In Situ Hybridisation (FISH) (Section 2.11), visualised by

CLSM (LSM 510 Meta, Carl Zeiss, Germany; Section 2.12) and analysed using

COMSTAT to determine the roughness coefficient, biofilm biomass, average

thickness and maximum thickness and percentage surface colonisation (Heydorn et

al., 2000; Section 2.12).

The biometric data were statistically analysed using SPSS software version

22.0 by applying ANOVA with a post hoc Tukey test to compare biometric

parameters of C. albicans, A. naeslundii and S. mutans in mono-cultured and

polymicrobial biofilms between replicates. The biometric data were statistically

analysed using SPSS software version 22.0. An independent t-test was applied to

compare the biometric parameters between mono-cultured and polymicrobial biofilms

except for the mono-cultured A. naeslundii, which was analysed using the Mann-

Whitney test due to the wide standard deviation (non-parametric data). Statistical

analyses were considered significant when P < 0.05.

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5.4 Results

5.4.1 Mono-cultured biofilms of C. albicans, A. naeslundii and S. mutans

The present study showed that C. albicans, A. naeslundii and S. mutans were

able to form biofilms in a flow environment when grown in mono-culture after 24 h

incubation (Figure 5.1A, Figure 5.1B, Figure 5.1C). Of all three mono-cultured

biofilms, S. mutans had the largest biomass (27.70 ± 2.83 µm3 µm-2), average

thickness (32.03 ± 1.96 μm) and maximum thickness (35.13 ± 2.87 μm). Conversely,

the mono-cultured A. naeslundii biofilm exhibited the smallest biomass (0.85 ± 0.68

µm3 µm-2) and average thickness (4.26 ± 2.98 μm), being 32-times and 8-times lower

than that of S. mutans respectively (Table 5.1; Table 5.2). In addition, the mono-

cultured C. albicans biofilm had the smallest maximum thickness (15.53 ± 4.47 μm)

being 2-times lower than for S. mutans (Table 5.2).

5.4.2 Polymicrobial biofilms of C. albicans, A. naeslundii and S. mutans

C. albicans, A. naeslundii and S. mutans formed biofilms in a flow

environment when grown as polymicrobial biofilms after 24 h incubation based on the

average values over three biological replicates (Figure 5.1D). The total biomass of

polymicrobial biofilms was 2.017 ± 0.088 µm3 µm-2 with the biomasses of C.

albicans, A. naeslundii and S. mutans that formed polymicrobial biofilms being 0.94 ±

0.24 µm3 µm-2 (46.8%), 0.66 ± 0.81 µm3 µm-2 (32.8%) and 0.41 ± 0.27 µm3 µm-2

(20.4%) respectively (Table 5.1). The roughness coefficient, average thickness,

maximum thickness and percentage colonisation of polymicrobial biofilms was 0.78 ±

0.40 Ra, 8.54 ± 6.62 µm, 22.47 ± 4.47 µm and 13.07%, respectively (Table 5.2). In

addition, the maximum thickness of the polymicrobial biofilms was 22.40 ± 4.50 µm.

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5.4.3 Effect of polymicrobial interaction on C. albicans, A. naeslundii and

S. mutans biofilms

A variation in the biometric parameters of biofilms was observed when

comparisons were made between mono-cultured C. albicans, A. naeslundii and S.

mutans with polymicrobial biofilms (Table 5.2). The present study showed that

mono-cultured S. mutans biofilms exhibited statistically significant larger average

thickness (32.03 ± 1.96 µm), maximum thickness (35.15 ± 2.87 µm) and surface

colonisation (29.94%) compared to the polymicrobial biofilms (8.54 ± 6.62 µm, 22.47

± 4.47 µm and 13.07%, respectively; P < 0.05; Table 5.2).

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Figure 5.1A Representative CLSM image of mono-cultured C. albicans as observed using a 63x objective at 512 x 512 pixels magnification. Biofilm was developed on ASM-coated glass substratum in a flow-cell system for 24 h (3 mL h-1) at 37 °C to form biofilm. The biofilm was embedded in gel acrylamide, labelled using FISH technique and visualised by CLSM (LSM 510 Meta, Carl Zeiss, Germany.

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Figure 5.1B Representative CLSM image of mono-cultured A. naeslundii as observed using a 63x objective at 512 x 512 pixels magnification.

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Figure 5.1C Representative CLSM image of mono-cultured S. mutans as observed using a 63x objective at 512 x 512 pixels magnification.

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Figure 5.1D Representative CLSM image of polymicrobial biofilms as observed using a 63x objective at 512 x 512 pixels magnification (Red: C. albicans; Green: A. naeslundii; Blue: S. mutans).

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Table 5.1 Total biomass (µm3 µm-2) of ASM-grown C. albicans, A. naeslundii and S. mutans after 24 h incubation in a flow-cell (3 mL h-1) at 37 °C in mono-cultured biofilm and polymicrobial biofilms.

Microorganisms Mono-cultured biofilm (µm3 µm-2)

Polymicrobial biofilms (µm3 µm-2) P value

C. albicans 4.43 (1.21)

0.94 (0.24) P < 0.05#

A. naeslundii 0.85 (0.68)

0.66 (0.81) P < 0.05*

S. mutans 27.70 (2.83)

0.41 (0.27) P < 0.05#

Data are means from three separate experiments (SD are given in parenthesis). Data were analysed using independent t-test# and Mann-Whitney* to compare between mono-cultured and polymicrobial biofilms of specific microorganisms. Data were considered as significantly different when P < 0.05.

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Table 5.2 Surface roughness, average and maximum thickness and percentage surface colonisation of ASM-grown C. albicans, A. naeslundii and S. mutans after 24 h incubation in a flow-cell (3 mL h-1) at 37 °C.

C. albicans mono-cultured biofilm

A. naeslundii mono-cultured

biofilm

S. mutans mono-cultured biofilm

Polymicrobial biofilms

Mean P value* Mean P value* Mean P value* Mean Roughness

coefficient (Ra) 0.55

(0.17) P > 0.05 1.24 (0.43) P < 0.05 0.12

(0.05) P < 0.05 0.78 (0.40)

Average thickness (µm)

4.59 (0.73) P < 0.05 4.26

(2.98) P < 0.05 32.03 (1.96) P < 0.05 8.54

(6.62) Maximum

thickness (µm) 8.98

(2.05) P < 0.05 15.53 (4.47) P < 0.05 35.13

(2.87) P < 0.05 22.47 (4.47)

Surface colonisation (%)

45.63 (25.25) P < 0.05 3.66

(2.43) P < 0.05 29.94 (3.74) P < 0.05 13.07

(4.01) Data are means from three separate experiments (SD are given in parenthesis). Data were analysed using independent t-test* to compare between mono-cultured and polymicrobial biofilms of the same biometric (e.g. ‘Roughness coefficient’ of mono-cultured C. albicans biofilm was compared with the ‘roughness coefficient’ of polymicrobial biofilms). Green showed significantly higher and red showed significantly lower in mono-cultured biofilms compared to polymicrobial biofilms. Data were considered as significantly different when P < 0.05.

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5.5 Discussion

To our knowledge, this is the first study to evaluate polymicrobial biofilm

formation by C. albicans, A. naeslundii and S. mutans in a flow-cell environment

using an artificial saliva medium. The present study measured five different

biometric parameters; the roughness coefficient, biofilm biomass, average biofilm

thickness, maximum biofilm thickness and percentage surface colonisation of the

cells in the biofilm.

The present study showed that C. albicans, A. naeslundii and S. mutans

formed biofilms on artificial saliva-coated substratum when grown as a mono-

cultured and polymicrobial biofilm in a flow-cell environment, thus supporting the

hypothesis that C. albicans, A. naeslundii and S. mutans form biofilm in a flow

environment. In this system, A. naeslundii was a very poor biofilm former, whilst S.

mutans produced a robust biofilm when grown as a mono-cultured biofilm. Studies

using gamma-irradiated Stovall flow-cell systems (40 mm long, 4 mm wide and 1 mm

deep) have shown that A. naeslundii and S. mutans form mono-cultured biofilms

when grown in sucrose containing tryptic soy broth and ASM respectively (Dashper

et al., 2013; Blanc et al., 2014; Arai et al., 2015). Furthermore, C. albicans and S.

mutans have been shown to form biofilms both mono-cultured and co-cultured in a

flow environment using saliva supplemented with HIB and PBS in a flow-cell track

(40 mm long, 3 mm wide and 2 mm deep) (Diaz et al., 2012). An in vitro study using

a flow-cell system was shown to simulate the conditions encountered by

microorganisms in the oral cavity such as shear stress rates due to salivary flow

(Sánchez-Vargas et al., 2013). A more robust biofilm formed by S. mutans in the

flow environment is suggested due to the ability of the bacterium to utilise sucrose

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from 25% ASM subsequently forming extracellular polysaccharides (EPS) through

glucosyltransferases (Koo et al., 2010; Bowen and Koo, 2011; Ren et al., 2016). This

did not occur with A. naeslundii in the present study, where poor biofilm forming

ability was observed. It could be that the EPS which are formed in the flow

environment promote the adherence of S. mutans, thus enhancing the development of

a mono-cultured S. mutans biofilm. Further study such as the quantification of EPS is

required to support this hypothesis.

The biomass of the polymicrobial biofilms was significantly reduced

compared to the biomass of C. albicans and S. mutans mono-cultured biofilms.

Furthermore, the percentage surface colonisation in the polymicrobial biofilms was

significantly lower than for mono-cultured C. albicans and S. mutans biofilms, but not

for A. naeslundii. These findings support our hypothesis that polymicrobial

interactions affected microbial colonisation in a flow environment. Even though in

vitro studies have shown that mutualistic interactions between C. albicans and S.

mutans occur through adhesins (non-specific) and lectin-saccharide cell surface

receptors (specific) bindings (McIntire et al., 1982; Rickard et al., 2003; Rosen and

Sela, 2006; Ledder et al., 2008), antagonism between the two species has also been

reported (Thein et al., 2006). C. albicans has been shown to decrease adherence

when co-cultured with S. mutans on acrylic sheets in Gibbons and Nygaard culture

medium (Barbieri et al., 2007). Furthermore, the quorum-sensing molecule Farnesol

that is synthesised by C. albicans during biofilm formation has been reported to

disrupt the membrane of S. mutans, as well as the accumulation of polysaccharide

contents of streptococcal biofilms (Koo et al., 2003; Jabra-Rizk et al., 2006).

Our study has shown that the biomass of both C. albicans and S. mutans in

polymicrobial biofilms was significantly decreased more than 50% compared to the

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mono-cultured biofilms. Furthermore, a negative effect was also observed in the

surface colonisation of C. albicans and S. mutans which exhibited a significant

decrease in polymicrobial biofilms. It would appear that A. naeslundii may have

some potential as a probiotic to inhibit the colonisation of C. albicans in the oral

cavity. Antagonism has been reported between C. albicans and A. naeslundii

(Millsap et al., 1999; Thein et al., 2006). A. naeslundii T14V-J1 has been shown to

suppress the adhesion of C. albicans ATCC 10261 when grown in a flow-cell

chamber (Millsap et al., 2000). The metabolic products of A. naeslundii have been

reported to both inhibit and stimulate the biofilm formation of C. albicans, depending

on the experimental methods employed (Gutiérrez and Benito, 2004; Thein et al.,

2006). In addition, antagonism between S. mutans and A. naeslundii has been widely

reported due to the production of H2O2 and bacteriocins by Streptococcus spp.

(Jakubovics et al., 2008; Avila et al., 2009; Zhu and Kreth, 2012). Although

antagonism between A. naeslundii and S. mutans has been reported, co-aggregation

assays have shown that both species can grow in close proximity (Zhu and Kreth,

2012; Arzmi et al., 2015). It may be that the cell densities of the microorganisms and

C. albicans morphology in polymicrobial biofilms influenced the interaction between

C. albicans, A. naeslundii and S. mutans. A previous study has shown that

polymicrobial biofilms that were generated from an inoculum of 107 cells mL-1 of C.

albicans (hyphal growth) and 108 cells mL-1 of bacteria (Streptococcus oralis and

Actinomyces oris) in a static biofilm system exhibited synergism between

microorganisms (Cavalcanti et al., 2016). In contrast, antagonism between C.

albicans and Actinomyces israelii has also been reported in static polymicrobial

biofilms (Thein et al. 2006).

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In the present study, we observed that all three microorganisms had similar maximum

thickness in polymicrobial biofilms. However, there was significant variation when

grown in mono-culture with S. mutans having the greatest maximum thickness. These

findings support the hypothesis of the present study that polymicrobial interactions

affect colonisation of oral microorganisms. A. naeslundii is an early oral coloniser

that binds to proline-rich proteins of the salivary pellicle (Kolenbrander et al., 2010).

In the presence of high sucrose in 25% ASM, S. mutans has been reported to produce

large amounts of glucosyltransferases that aid the attachment of C. albicans and A.

naeslundii through glucan binding proteins (Koo et al., 2010). Gtfs are associated

with the production of EPS, the prime building blocks of dental plaque (Koo et al.,

2010), and this is most likely the principle contributor to the extracellular component

of the polymicrobial biofilm. Complex polymicrobial biofilms in the oral cavity are

associated with a number of disease states, such as oral candidosis, dental caries and

periodontal diseases (Harriott and Noverr, 2011).

Based on the static biofilm study of eight strains of C. albicans, it has been

shown that the biofilm formation is C. albicans strain-dependent (chapter 4).

However, a similar claim is inappropriate for A. naeslundii and S. mutans since there

were only A. naeslundii (NCTC 10301) and S. mutans (Ingbritt) have been used in the

study. It is suggested that different strain of A. naeslundii and S. mutans may form

different biofilm biometric parameters when grown in flow-cell system. Previous

study has shown that 44 genotypes of S. mutans were producing different range of

biofilm due to the variable amount of glucosyltransferases (Gtfs), particulalrly GtfB

and GtfC expressed by S. mutans (Mattos-Graner et al., 2004). Furthermore, A.

naeslundii genospecies 2 has been the most frequent isolated from adults compared to

genospecies 1 (Paddick et al., 2003). Therefore, it can be postulated that biofilm

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formation may also be A. naeslundii and S. mutans strain-dependent. Further research

is required to support this hypothesis.

5.6 Conclusion

In conclusion, C. albicans, A. naeslundii and S. mutans formed polymicrobial

biofilms in a flow environment. The overall biomass of the polymicrobial biofilm was

low relative to mono-cultured biofilms indicating significant antagonistic interactions

between these species. This was shown to affect the surface roughness, biofilm

thickness and surface colonisation in a flow-cell environment. Furthermore, A.

naeslundii may have some potential as a probiotic to control C. albicans and S.

mutans overgrowth, providing a dynamic balance between C. albicans, A. naeslundii

and S. mutans. Thus, these interactions are likely to play significant roles in the

pathogenicity of oral microorganisms, plaque formation, dysbiosis and oral diseases

(Kolenbrander, 2000; Sbordone and Bortolaia, 2003; Min and Rickard, 2009; Morales

and Hogan, 2010).

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CHAPTER 6

BIOFILM EFFLUENT OF CANDIDA ALBICANS, ACTINOMYCES

NAESLUNDII AND STREPTOCOCCUS MUTANS AFFECT THE ADHESION,

EPITHELIAL MESENCHYMAL TRANSITION AND CYTOKINE

EXPRESSION OF NORMAL AND MALIGNANT ORAL KERATINOCYTES

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6.1 Abstract

Microbial infections, including those caused by C. albicans, S. mutans and A.

naeslundii have been suggested to play a role in carcinogenesis. Malignant tumours

such as carcinomas are characterised by the ability of tumour cells to invade the

underlying connective tissues followed by migration to form metastases at distant

sites. In this context, epithelial to mesenchymal transition (EMT) has been shown to

assist in cell migration through extracellular matrix (ECM), by inducing the formation

of the mesenchymal phenotype of epithelial cells, which is important for metastasis.

This pro-invasive phenotype associates with an altered integrin-ECM adhesion and

inflammation induced by pathogens has been suggested to be involved in determining

characteristics of the tumour microenvironment. In the present study, we assessed the

ability of microbial biofilm effluent obtained from C. albicans (ALC3), A. naeslundii

(AN), S. mutans (SM) and poly-microbial (TRI) biofilms, to induce a pro-invasive

phenotype in oral epithelial cells. To study EMT, OKF6 (normal oral epithelial cell)

and H357 (oral squamous cell carcinoma) cell lines were incubated for 2 h and 24 h at

37 °C, 5% CO2, in test cell growth media (80% serum free medium). The cells were

collected and flow cytometry was undertaken for the detection of vimentin and E-

cadherin. CytoSelect 48-well Cell Adhesion Assay ECM Array kit was used to study

the adhesion of OKF6 and H357 to ECM components. Simultaneously, the

conditioned medium was collected and the presence of cytokines was detected using

Bio-Plex. The present study showed that the incubation of H357 in ALC3 effluent

significantly increased the adhesion of these malignant cells to collagen IV and

laminin I when compared to control non-effluent (NE) (P < 0.05). Furthermore, a

significant decrease of vimentin was observed after 24 h incubation when incubated

with ALC3 compared to NE (P < 0.05). ALC3 effluent was also found to

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significantly increase the expression of IL-10 and GM-CSF from H357 after 2 h

incubation (P < 0.05), and IL-2, IL-4, IL-6, IL-8, IL-10, GM-CSF and TNF-α after 24

h compared to NE (P < 0.05). Finally, an increase of the majority of pro-

inflammatory cytokines in H357 incubated with ALC3 and TRI effluent was observed

after 24 h incubation compared to 2 h. Overall, the majority of H357 incubated in

biofilm effluent increased adhesion to ECM components, and significantly increased

the expression of inflammatory cytokines. For OKF6, the majority of cells showed

significantly decreased adhesion to ECM components (P < 0.05) and exhibited no

change in cytokine expression when compared to NE. Taken together these results

demonstrated that the adhesion of OKF6 and H357 to ECM, EMT and cytokine

expression, are biofilm effluent-dependent. Furthermore, the biofilm effluent affect

on the adhesion, EMT and cytokine expression from OKF6 and H357 showed an

enhanced malignant phenotype when grown in the presence of polymicrobial biofilm

effluent; this may act as a promoter of oral cancer but most likely not as an initiator.

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6.2 Introduction

Cancer is the leading cause of death in developed countries and the second in

the developing countries (Jemal et al., 2011). In 2008, about 12.7 million cancer

cases were reported worldwide, and of these, 263,900 were oral cancer

(Warnakulasuriya, 2009; Jemal et al., 2011). The highest oral cancer rates are found

in South-Central Asia, and Central and Eastern Europe whereas the lowest cases are

in Africa, Central America and Eastern Asia (Jemal et al., 2011). In countries such as

Pakistan, Bangladesh, India and Sri Lanka, oral cancer is the most common cancer

and accounts for up to 30% of all diagnosed cancers (Warnakulasuriya, 2009). In

Malaysia, oral cancer has been the 20th most common cancer in females and 28th for

males with the highest rates being reported for Indian ethnicity followed by Malay

and Chinese (Ashazila et al., 2011).

Oral squamous cell carcinoma (OSCC) accounts for more than 90% of

malignancies originating from the oral mucosa (Casiglia and SB, 2001; Johnson et al.,

2011). It has been reported that the average 5-year survival rate for oral cancer is less

than 50% (Zakrzewska, 1999). These poor figures largely reflect the tumour stage at

presentation as well as the development of loco-regional recurrences and distant

metastases. Hence, the acquisition of a metastatic phenotype is of paramount

importance in determining oral cancer progression and prognosis. The risk factors that

lead to OSCC include tobacco smoking, heavy alcohol consumption, poor oral

hygiene, unhealthy diets, and microbial infections (Hooper et al., 2009;

Chocolatewala et al., 2010; Meurman, 2010; Rajeev et al., 2012; Khajuria and

Metgud, 2015).

Microbial infections by yeasts such as C. albicans, bacteria and viruses have

been widely suggested to have a causal role in oral cancer (Rodriguez et al, 2007;

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Scheper et al., 2008; Hooper et al., 2009; Chocolatewala et al., 2010; Meurman,

2010; Rajeev et al., 2012; Marttila et al., 2013; Khajuria and Metgud, 2015). The

aetiological role of Candida spp. in oral mucosal keratoses progression to carcinoma

has been suggested since 1966 with the majority of non-homogenous leukoplakias

invaded by C. albicans and these have higher malignant transformation potential than

the homogenous leukoplakia (Cawson, 1969a; Cawson, 1969b; Meurman, 2010).

Furthermore, oral yeast carriage has also been found to correlate with the presence of

oral epithelial dysplasia, which supports the role of microbial infections in oral

carcinogenesis (McCullough et al., 2002).

Streptococcus spp. and Actinomyces spp. have been postulated to be involved

in oral carcinogenesis. An in vitro study on S. mutans has shown that this bacterium is

able to synthesise the alcohol dehydrogenase enzyme that converts alcohol to

carcinogenic acetaldehyde, widely reported to be involved in oral carcinogenesis

(Kurkivuori et al., 2007; Hooper et al., 2009). Furthermore, study of the oral

microbiome of patients with OSCC has shown increased numbers of Actinomyces

spp., including A. naeslundii, compared to the individuals with a healthy oral cavity

(Nagy et al., 1998; Pushalkar et al., 2011).

Carcinoma is characterised by the ability of the malignant cell to invade the

underlying connective tissues followed by migration to form metastases at distant

sites (Lyons and Jones, 2007). These processes require the alteration of cell to cell

and cell to extracellular matrix (ECM) interactions that involves adhesion molecules

such as collagen, laminin, fibronectin and fibrinogen (Ahmed et al., 2005; Lyons and

Jones, 2007). These cell adhesion-proteins have been shown to promote the

attachment and migration of cancerous cells to surrounding ECM and are believed to

be involved in tumour cell survival, metastasis and angiogenesis (Fabricius et al.,

141

2011). Integrins, which are in contact with complementary ECM molecules, regulate

normal cell behavior and alterations of integrin-mediated cell adhesion machinery

have been implicated in oral carcinogenesis (Rathinam and Alahari, 2010; Fabricius et

al., 2011).

Epithelial-mesenchymal transition (EMT) is a mechanism of alteration of cell-

to-cell and cell-to-ECM interaction that allows the movement of the epithelial cells to

the surrounding environment (Radisky, 2005). This mechanism has been shown to

assist in cell migration through ECM by inducing the formation of the mesenchymal

phenotype of the epithelial cell. In normal conditions, epithelial cell structure is

maintained by cell-to-cell interactions such as cadherin-based adherent junctions and

desmosomes, whereas mesenchymal cells are mostly without direct contact or defined

cell polarity, but have distinct cell-to-ECM interactions and cytoskeletal structures

(Radisky, 2005). An inappropriate utilisation of EMT may occur in the formation of

OSCC and metastasis of malignant cells (Kang and Massagué, 2004; Yang et al.,

2004; Radisky, 2005). Furthermore, EMT has also been reported to be involved in

the increase of resistance of malignant cells to apoptosis regulator molecules (Maestro

et al., 1999; Vega et al., 2004).

Vimentin and E-cadherin have been widely used as markers for EMT (Hugo et

al., 2007). Vimentin is a protein that belongs to type III intermediate filaments (IF).

IFs are expressed by nearly all eukaryotic cells and are composed of proteins that

provide mechanical strength to the structure of tissues (Cooper, 2000). The

expression of vimentin has been shown to induce the changes in cell shape, motility

and adhesion of epithelial cells to mesenchymal (Mendez et al., 2010). During EMT,

epithelial cells lose adhesion to neighbouring cells and change shape to be elongated

and flat, a common morphology of mesenchymal cells. During this process, vimentin

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is expressed, and this correlates with both mesenchymal shape and enhanced motility

(Mendez et al., 2010). Further, vimentin has been shown to be expressed in vivo

during tumorigenesis and metastasis in prostate cancer and metastatic breast

carcinoma (Lang et al., 2002). Therefore, the increased amount of vimentin from

malignant oral tissue is an indicator of malignancy (Lang et al., 2002; Hugo et al.,

2007; Nijkamp et al., 2011).

Cadherins have been shown to mediate cell-to-cell binding that is critical in

maintaining tissue structure and morphology (Gumbiner, 2005). E-cadherin is a

glycoprotein that establishes homophilic interactions with E-cadherin-adjacent-

molecules expressed by neighbouring cells to produce the core of epithelial adherence

junction (Nagafuchi et al., 1987; Gumbiner, 2005). Functional loss of E-cadherin in

epithelial cells has been considered as a marker for EMT during tumour progression

(Onder et al., 2008; Nijkamp et al., 2011; Yadav et al., 2011). Cells expressing E-

cadherin have been reported to be silenced by a number of different mechanisms

including transcriptional repression (Bolós et al., 2003), histone deacetylation

(Peinado et al., 2004), down-regulation of gene expression through promoter

hypermethylation (Hasegawa et al., 2002) and somatic mutation (Berx et al., 1995).

Inflammation induced by pathogens has been shown to be involved in

carcinogenesis, particularly after the classification of Helicobacter pylori as a class-1

carcinogen in humans by the World Health Organization (WHO) International

Agency for Research on Cancer (IARC) (Peek and Blaser, 2002, Björkholm et al.,

2003, Correa and Houghton, 2007). One of the factors that leads to inflammation is

the increase of pro-inflammatory cytokines due to microbial infection of oral mucosa

(Fantini and Pallone, 2008). Cytokines are signalling molecules that regulate the

differentiation, proliferation and many other important functions of human

143

inflammatory cells. Cytokines are important in host defence and their release from

infected tissues has been shown to activate effector immune cells (leukocytes),

subsequently activating a cascade of specific defence mechanisms towards pathogens.

The cytokines that have been shown to be involved in inflammation include

interleukin (IL)-1α, IL-1β, IL-6, IL-8, IL-18, tumour necrosis factor (TNF)-α, IFN-γ

and GM-CSF (Dongari-Bagtzoglou et al., 1999; Rouabhia et al., 2002, Schaller et al.,

2002, Steele and Fidel, 2002, Dongari-Bagtzoglou and Kashleva, 2003a, Dongari-

Bagtzoglou and Kashleva, 2003b).

Despite considerable clinical and experimental evidence describing the

association between C. albicans and malignant transformation, no study has been

conducted to investigate the effect of C. albicans, S. mutans and A. naeslundii biofilm

effluent on the adhesion of normal and malignant oral keratinocytes. Furthermore, the

paracrine regulation of EMT from C. albicans alone or in the context of polymicrobial

biofilms has never been investigated. In particular, no study has been conducted

regarding the effect of effluent from polymicrobial biofilms formed by the yeast C.

albicans, S. mutans and A. naeslundii on the expression of vimentin and E-cadherin in

the normal oral epithelial cell OKF6 and the OSCC cell line H357. These cell lines

were selected in the present study to determine the role of effluent from biofilms

formed by C. albicans, S. mutans and A. naeslundii to: (1) affect the adhesion of

normal and OSCC cell lines to ECM molecules (fibronectin, collagen I, collagen IV,

laminin I, and fibrinogen), (2) to affect EMT of normal and OSCC cell lines as

indicated by the expression of vimentin and E-cadherin, and (3) to affect cytokine

expression (IL-2, IL-4, IL-6, IL-8, IL-10, TNF-α, GM-CSF and IFN-γ) of normal and

OSCC cell lines. Collectively, we hypothesised that the biofilm effluent affects the

adhesion and EMT of OKF6 and H357, and the expression of pro-inflammatory

144

cytokines from the cell lines, which enhances their malignant phenotype when grown

in the presence of polymicrobial biofilm effluent and may have a role in oral

carcinogenesis.

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OKF6, a normal human oral epithelial cell and H357, cell isolated from the

tongue of patient with OSCC, were grown as described in Section 2.13.

Subsequently, Cell-ECM adhesion was conducted (Section 2.15) to determine the

adhesion of OKF6 and H357 to extracellular proteins of fibronectin, collagen I

collagen IV, laminin and fibrinogen when treated with test cell growth media (Section

2.14) using CytoSelect 48-well Cell Adhesion Assay ECM Array kit (Cell Biolabs,

USA). The epithelial-mesenchymal transition (EMT) assay was conducted to

determine the expression of E-cadherin and vimentin (Section 2.17), whereas the

expression of pro-inflammatory cytokines from OKF6 and H357 which have been

grown in biofilm effluent was assessed with Bio-plex assays (Section 2.18). Test cell

growth medium was prepared using SFM (DMEM/F12 and k-SFM for H357 and

OKF6 respectively), that was diluted in biofilm effluent (Section 2.9) of (1) C.

albicans (ALC3), (2) A. naeslundii (AN), (3) S. mutans (SM) and poly-microbial

(TRI), and non-effluent 25% ASM (NE), to give a final concentration of 80% (v/v)

SFM (Steele and Fidel, 2002). NE is the control to determine the role of biofilm

effluent on cell-ECM adhesion, EMT and the expression of cytokines from OKF6 and

H357.

All data were statistically analysed using SPSS Statistic software version 22.0

using ANOVA post hoc Dunnett’s test to compare the adhesion, EMT and cytokine

expression of cells incubated with biofilm effluent to control (NE), and to compare

between two time points for each assay. ANOVA was chosen due to the data being

normally distributed and the post hoc Dunnett’s test was chosen due to the presence of

the control group (NE). Results were considered as statistically significant when P <

0.05.

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6.4 Results

Part I: Adhesion assay

6.4.1 Adhesion of OKF6 to ECM

Incubation with ALC3 effluent caused a significant decrease in OKF6

adhesion to fibronectin, collagen I, collagen IV and laminin I compared to incubation

with NE (P < 0.05; Table 6.1A). Incubation with AN effluent caused a significant

decrease of adhesion to fibronectin, collagen I and laminin (P < 0.05), whereas

incubation with SM effluent showed decrease of adhesion to collagen IV and

fibrinogen significantly when compared to the incubation in NE (P < 0.05; Table

6.1A). Furthermore, incubation of OKF6 with TRI caused a significant decrease of

adhesion to collagen I and fibrinogen when compared to the incubation in NE (P <

0.05). There was only one situation where OKF6 showed significantly enhanced

adhesion compared to NE, and that was when incubated with S. mutans effluent

where adhesion to fibronectin was increased (P < 0.05; Table 6.1A).

6.4.2 Adhesion of H357 to ECM

Incubation with ALC3 effluent caused a significant decrease of H357 adhesion

to fibronectin and fibrinogen (P < 0.05). However, significant increases were

observed in the adhesion to collagen IV and laminin I, when compared to the

incubation in NE (P < 0.05; Table 6.1A). Incubation of H357 with AN effluent

caused a significant decrease of adhesion to fibronectin. However, a significant

increase of adhesion to fibrinogen was observed compared to the incubation with NE

(P < 0.05; Table 6.1A). The adhesion of H357 to collagen I, collagen IV and laminin

I were increased significantly (P < 0.05) when incubated with SM effluent, whereas a

significant decrease of adhesion to fibronectin and fibrinogen was observed compared

147

to incubation with NE (P < 0.05; Table 6.1A). Incubation with TRI effluent caused a

significant increase of H357 adhesion to fibronectin, collagen I and laminin I (P <

0.05). However, the adhesion of H357 to fibrinogen was observed to decrease

significantly compared to incubation of H357 with NE (P < 0.05; Table 6.1A).

6.4.3 Comparison of adhesion to ECM between OKF6 and H357

The data demonstrated that the majority of biofilm effluent decreased the

adhesion of OKF6 to ECM components, as incubation in all biofilm effluents showed

decreased adhesion of OKF6 to collagen I, collagen IV and fibrinogen when

compared to NE (Table 6.1B; Figure 6.1A). Meanwhile, the majority of microbial

effluent increased the adhesion of H357 to ECM components, while incubation in all

biofilm effluents showed increased adhesion of H357 to laminin I when compared to

NE (Table 6.2; Figure 6.1B). Of note, the large fold changes of increased adhesion of

H357 cells to collagen IV observed after incubation with biofilm effluent from ALC3

(4.58-fold) and SM (2.07-fold). Furthermore, very large enhanced adhesion to

laminin I was observed when H357 was incubated with ALC3 (15.07-fold), SM

(6.54-fold) and TRI (10.69-fold) effluents (Table 6.1B; Figure 6.1B).

Collectively, the results of adhesion assays showed that microbial biofilm

effluent influenced cell-ECM interaction and this was dependent on the pathogen

present in the biofilm and the cell used in the analysis (Table 6.1B; Figure 6.1).

Effluent increased adhesion to ECM components more commonly for the oral cancer

cell line (H357) compared with the normal oral epithelial cell line (OKF6), and this

was most pronounced for adhesion to collagen IV and laminin I.

148

Table 6.1A Adhesion of OKF6 and H357 in 80% serum free medium (SFM) containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents.

ECM molecules

OKF6 H357

NE ALC3 AN SM TRI NE ALC3 AN SM TRI

Fibronectin 0.154

(0.005) 0.080

(0.004) 0.122

(0.009) 0.177

(0.000) 0.149

(0.007) 0.622

(0.043) 0.329

(0.062) 0.422

(0.025) 0.459

(0.030) 0.752

(0.003)

Collagen I 0.165

(0.001) 0.109

(0.009) 0.139

(0.000) 0.157

(0.004) 0.106

(0.011) 0.768

(0.065) 0.645

(0.028) 0.298

(0.035) 0.994

(0.101) 1.155

(0.049)

Collagen IV 0.176

(0.027) 0.133

(0.004) 0.171

(0.013) 0.134

(0.009) 0.156

(0.004) 0.376

(0.137) 1.537

(0.039) 0.189

(0.007) 0.675

(0.111) 0.366

(0.109)

Laminin I 0.041

(0.001) -0.008 (0.009)

-0.008 (0.001)

0.022 (0.018)

0.063 (0.037)

0.042 (0.000)

0.628 (0.077)

0.055 (0.001)

0.273 (0.035)

0.445 (0.093)

Fibrinogen 0.180

(0.011) 0.099

(0.022) 0.146

(0.002) 0.101

(0.027) 0.097

(0.017) 0.214

(0.002) 0.050

(0.002) 0.289

(0.006) 0.096

(0.008) 0.105

(0.003)

Significantly higher compared to NE No-significant difference compared to NE Significantly lower compared to NE

Data is the optical density measured by spectrophotometer at wavelength OD570nm. Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data is considered as significantly different when P < 0.05.

149

Table 6.1B Fold change of OKF6 and H357 adhesion when incubated with C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents compared to non-effluent (NE).

ECM Molecules

OKF6 H357

ALC3 AN SM TRI ALC3 AN SM TRI

Fibronectin 0.52

(0.01) 0.79

(0.08) 1.15

(0.03) 0.97

(0.01) 0.53

(0.06) 0.68

(0.03) 0.74

(0.03) 1.21

(0.00)

Collagen I 0.66

(0.06) 0.84

(0.00) 0.95 (0.0)

0.64 (0.01)

0.84 (0.03)

0.39 (0.04)

1.31 (0.10)

1.51 (0.05)

Collagen IV 0.77

(0.10) 0.99

(0.21) 0.78

(0.01) 0.91

(0.00) 4.58

(0.04) 0.57

(0.01) 2.07

(0.11) 1.04

(0.11)

Laminin I -0.20* (0.22)

-0.20* (0.03)

0.54 (0.02)

1.53 (0.04)

15.07 (0.08)

1.31 (0.00)

6.54 (0.04)

10.69 (0.09)

Fibrinogen 0.55

(0.11) 0.81

(0.05) 0.56

(0.03) 0.54

(0.02) 0.23

(0.00) 1.35

(0.01) 0.45

(0.01) 0.49

(0.00)

Increased compared to NE Decreased compared to NE

Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. * Cells adhered to laminin I were less than those adhered to the control substrata (Bovine serum albumin, BSA).

150

Figure 6.1 Fold change of OKF6 and H357 adhesion when incubated with C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM) and polymicrobial (TRI) biofilm effluents compared to non-effluent (NE). Bars are the SD. The study was conducted in three biological replicates with each replicate consisting of three technical replicates.

-5.00

0.00

5.00

10.00

15.00

20.00

Fold

cha

nge

A) OKF6

ALC3

AN

SM

TRI

-5.00

0.00

5.00

10.00

15.00

20.00

Fold

cha

nge

B) H357

ALC3

AN

SM

TRI

151

Part II: Epithelial-mesenchymal transition (EMT)

6.4.4 Percentage of cells expressing E-cadherin and vimentin

Incubation of OKF6 with ALC3, AN and SM effluent caused a significant

increase of cells expressing vimentin compared to NE after 24 h (P < 0.05; Table

6.2A). Incubation of the cell line with all biofilm effluents caused a significant

increase of cells expressing E-cadherin after 24 h when compared to NE (P < 0.05;

Table 6.2A).

The same set of experiments was subsequently performed on OSCC cells

(H357). Incubation of H357 with all effluents caused a significant decrease of cell

expressing vimentin compared to NE after 24 h incubation (P < 0.05; Table 6.2A). A

significant increase of cells expressing E-cadherin was observed between 2h and 24 h

incubation when H357 was incubated with SM compared to NE (P < 0.05; Table

6.2A).

All biofilm effluents and NE decreased OKF6 cells expressing vimentin after

24 h incubation compared to 2 h (-1.5% to -32.4%; Table 6.2B). An increase of cells

expressing E-cadherin was observed after 24 h when incubated in the majority of

biofilm effluents (ALC3, AN and SM) (12.5% to 23.0%; Table 6.2B).

The incubation of H357 in NE was found to increase cells expressing vimentin

(44.9%) and decrease cells expressing E-cadherin (-12.7%) after 24 h incubation

compared to 2 h (Table 6.2B). Over this time period, the majority of microbial

effluents (ALC3, AN and SM) reduced H357 cells expressing vimentin (-21.5% to -

23.6%) as well as cells expressing E-cadherin (-12.7% to -29.2%; Table 6.2B).

152

Table 6.2A Percentage positive of OKF6 and H357 cells treated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluent at 37 °C, 5% CO2 for 2 h and 24 h.

Hours EMT markers

OKF6 H357


2 Vimentin 62.1 (1.8)

61.1 (0.3)

56.4 (2.2)

60.6 (3.3)

69.2

(2.4) 52.7 (1.4)

56.2 (3.0)

59.0 (1.5)

63.8

(3.8) 60.6 (6.1)

E-cadherin 65.5 (1.3)

61.9 (0.6)

62.3 (1.9)

59.1 (2.3)

65.7 (1.0)

36.5 (1.1)

44.9 (3.9)

57.6

(1.8) 57.9 (2.4)

54.4 (4.6)

24 Vimentin 42.2

(2.4) 52.4 (2.5)

52.6 (3.8)

59.5

(2.4) 46.8

(2.4) 76.4 (1.0)

43.2 (3.1)

45.1 (0.1)

49.9 (5.5)

66.9 (1.2)


71.5

(5.7) 70.0 (2.0)

72.7

(0.3) 64.4 (3.9)

31.8

(0.1) 36.7 (2.9)

42.6

(1.8) 49.2 (3.1)

38.2 (0.6)


Data is percentage positive of cells expressing EMT markers as measured by flow cytometry. Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data was considered as significantly different when P < 0.05.

153

Table 6.2B Percentage difference of positive OKF6 and H357 cells expressing vimentin and E-cadherin between 2 h and 24 h incubated in NE, ALC3, AN, SM, and TRI effluents at 37 °C, 5% CO2 for 2 h and 24 h.

EMT markers

OKF6 (%) H357 (%)


Vimentin -32.0 (4.1)

-14.2 (4.4)

-6.5 (9.1)

-1.5 (9.7)

-32.4 (3.4)

44.9 (3.1)

-23.0 (6.2)

-23.6 (1.0)

-21.5 (7.6)

10.8 (9.0)

E-cadherin -16.6 (3.2)

15.6 (10.3)

12.5 (5.3)

23.0 (5.1)

-2.0 (7.0)

-12.7 (3.4)

-18.1 (9.0)

-26.0 (2.1)

-14.3 (15.6)

-29.2 (7.8)


Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Negative results indicated a decrease of positive cells expressing EMT markers, whereas positive results indicated an increase of positive cells after 24 h incubation. Data was considered as significantly different when P < 0.05.

154

6.4.5 Mean fluorescence intensity (MFI)

Mean fluorescence intensity (MFI) showed that the incubation of OKF6 in SM

effluent caused a significant increase of vimentin compared to NE after 24 h

incubation (P < 0.05; Table 6.3A). In addition, no biofilm effluent was observed to

cause a significant difference of E-cadherin expressed by OKF6 compared to NE (P >

0.05; Table 6.3A).

Incubation of the malignant cell line (H357) with ALC3, AN and SM effluent

was found to decrease the MFI of vimentin significantly after 24 h compared to

incubation with NE (P < 0.05; Table 6.3A). There was no significant difference of E-

cadherin between H357 incubated with biofilm effluent compared to incubation with

NE (P > 0.05; Table 6.3A).

All biofilm effluents and NE decreased expression of vimentin by OKF6 after

24 h incubation compared to 2 h (–16.9% to -53.1%; Table 6.3B). An increase of E-

cadherin was observed after 24 h when incubated in the majority of biofilm effluents

(ALC3, AN and SM; 2.0% to 11.3%; Table 6.3B). Furthermore, H357 when

incubated in NE was found to have increased expression of vimentin (31.7%) and

decreased expression of E-cadherin (-18.1%) after 24 h incubation compared to 2 h

(Table 6.3B). All biofilm effluents were found to reduce the expression of vimentin

by H357 (-30.2% to -58.1%; Table 6.3B). Furthermore, all biofilm effluents and NE

were observed to decrease E-cadherin (-18.1% to -42.2%; Table 6.3B).

Collectively, the data demonstrated that the expression of vimentin and E-

cadherin was regulated in a cell type, time-dependent and biofilm effluent-specific

manner. The expression of vimentin by the normal epithelial cell line, OKF6, was

profoundly decreased after 24 h incubation in NE (32.0%) (Table 6.2B) and MFI

155

(48.2%) (Table 6.3B). Although this decrease was observed when OKF6 was

incubated with mono-cultured biofilm effluent, this decrease was less pronounced

than for the control medium (P < 0.05). Furthermore, there was an increase in both

the number of cells expressing E-cadherin as well as MFI when OKF6 cells were

incubated with mono-cultured biofilm effluent (Table 6.2B and 6.3B).

Interestingly, a paradoxical effect was observed with the EMT of H357 cells,

with the majority of biofilm effluents resulting in an enhanced malignant phenotype

as observed by the decreased expression of E-cadherin, while at the same time having

a diminished malignant phenotype, as observed by the decreased expression of

vimentin.

156

Table 6.3A Mean fluorescence intensity (MFI) of vimentin and E-cadherin of OKF6 and H357 cells treated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h.

Hours EMT markers

OKF6 H357


2 Vimentin 1554.3 (134.1)

1666.3 (86.0)

1330.7 (81.3)

1745.0 (111.9)

2101.7

(188.6) 688.7 (14.5)

831.7 (93.3)

751.0 (71.3)

841.3 (87.4)

933.0

(19.1)


175.3 (5.9)

169.0

(70.0) 166.7

(5.9) 176.3 (4.9)

403.0 (13.1)

488.7 (34.5)

609.0

(39.6) 591.7

(69.6) 613.0

(61.3)

24 Vimentin 805.3

(84.7) 1103.7 (115.6)

1110.3 (227.9)

1370.7 (203.8)

992.3

(216.2) 907.0

(13.2) 346.7 (26.2)

352.7 (23.2)

411.7

(41.0) 651.0

(2.0)

E-cadherin 166.0 (44.2)

178.7 (17.2)

174.7 (5.1)

185.3

(4.2) 161.3

(4.2) 330.0

(4.6) 315.3

(18.0) 356.0

(2.6) 416.0 (43.3)

351.7

(6.0)


Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data was considered as significantly different when P < 0.05.

157

Table 6.3B Percentage difference of mean fluorescence intensity (MFI) of OKF6 and H357 cells expressing vimentin and E-cadherin between 2 h and 24 h incubated in NE, ALC3, AN, SM, and TRI effluents at 37 °C, 5% CO2 for 2 h and 24 h.

EMT markers

OKF6 (%) H357 (%)


Vimentin -48.2 (3.1)

-33.6 (8.4)

-16.9 (14.1)

-21.7 (6.9)

-53.1 (6.3)

31.7 (1.2)

-58.1 (4.3)

-53.0 (1.4)

-50.4 (9.6)

-30.2 (1.6)

E-cadherin -10.1 (22.7)

2.0 (11.5)

3.5 (6.1)

11.3 (3.7)

-8.4 (4.6)

-18.1 (1.6)

-35.3 (4.4)

-41.4 (3.6)

-28.4 (16.5)

-42.2 (6.5)


Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Negative results indicated a decreased percentage of MFI, whereas positive results indicated an increased percentage of MFI after 24 h incubation. Data was considered as significantly different when P < 0.05.

158

Part III: Cytokine assay

6.4.6 Expression of cytokines by OKF6 and H357

IL-2, IL-4, IL-6, IL-8, IL-10, GM-CSF and TNF-α were expressed

constitutively in OKF6 and H357, and were secreted at different concentrations when

grown in different cell types and effluents after 2 h and 24 h. In contrast, IFN-γ was

detected only in OKF6 incubated in NE, SM and TRI effluents for 24 h, and in H357

incubated in AN, SM and TRI for 2 h, and all effluents incubated for 24 h (Table 6.4A

and 6.5B).

159

Table 6.4A Cytokines expressed by OKF6 (pg mL-1) incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h.

Cytokines OKF6 (pg mL-1)

NE ALC3 AN SM TRI

IL-2 0.11

(0.04) 0.16

(0.05) 0.17

(0.04) 0.21

(0.00) 0.18

(0.05)

IL-4 0.04

(0.03) 0.05

(0.02) 0.03

(0.02) 0.05

(0.03) 0.03

(0.01)

IL-6 7.25

(1.77) 3.98

(1.29) 6.05

(3.99) 13.75 (3.43)

9.61 (2.83)

IL-8 50.64 (8.06)

36.27 (3.84)

42.09 (8.70)

71.10 (13.82)

54.13 (14.14)

IL-10 0.68

(0.05) 0.67

(0.04) 0.65

(0.10) 0.68

(0.05) 0.59

(0.04)

GM-CSF 7.09

(0.25) 7.04

(0.10) 6.44

(0.33) 7.25

(0.34) 6.28

(0.34)

IFN-γ LLD LLD LLD LLD LLD

TNF-α 1.27

(0.21) 1.41

(0.12) 1.34

(0.12) 1.41

(0.12) 1.34

(0.12)


Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data was considered as significantly different when P < 0.05. LLD = Less than lowest detectable measure. For IFN-γ, this was 0.4 pg mL-1.

160

Table 6.4B Cytokines expressed by OKF6 (pg mL-1) incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 24 h.

Cytokines OKF6 (pg mL-1)

NE ALC3 AN SM TRI

IL-2 0.33

(0.09) 0.33

(0.05) 0.33

(0.09) 0.33

(0.05) 0.45

(0.02)

IL-4 0.10

(0.03) 0.09

(0.03) 0.09

(0.02) 0.10

(0.03) 0.12

(0.01)

IL-6 36.64

(10.79) 24.69 (4.23)

30.59 (14.99)

42.79 (7.65)

79.85 (10.58)

IL-8 212.05 (13.92)

185.08 (39.66)

177.14 (56.71)

227.01 (100.73)

405.23 (84.00)

IL-10 0.74

(0.01) 0.76

(0.04) 0.69

(0.03) 0.78

(0.02) 0.70

(0.02)

GM-CSF 10.93 (1.78)

10.87 (0.28)

11.76 (1.89)

11.63 (0.65)

15.15 (1.56)

IFN-γ 0.85

(0.78) LLD LLD

1.30 (0.78)

3.36 (0.67)

TNF-α 1.90

(0.22) 1.62

(0.12) 1.69

(0.21) 1.98

(0.25) 2.37

(0.22)


Data was the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data was considered as significantly different when P < 0.05. LLD = Less than lowest detectable measure. For IFN-γ, this was 0.4 pg mL-1.

161

Table 6.4C Cytokines expressed by H357 (pg mL-1) incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h.

Cytokines H357 (pg mL-1)

NE ALC3 AN SM TRI

IL-2 0.16

(0.05) 0.23

(0.04) 0.60

(0.12) 0.69

(0.05) 0.18

(0.05)

IL-4 0.06

(0.02) 0.06

(0.02) 0.11

(0.00) 0.14

(0.01) 0.06

(0.02)

IL-6 12.21 (2.25)

16.74 (5.80)

189.98 (22.27)

234.16 (26.22)

39.36 (0.66)

IL-8 21.89 (4.07)

29.81 (5.55)

59.94 (9.12)

44.31 (0.48)

38.12 (8.67)

IL-10 0.58

(0.02) 0.67

(0.03) 0.70

(0.02) 0.71

(0.02) 0.68

(0.01)

GM-CSF 6.47

(0.24) 7.55

(0.22) 8.45

(0.17) 8.34

(0.66) 7.04

(0.25)

IFN-γ LLD LLD 2.77

(0.35) 5.06

(0.85) 1.30

(0.78)

TNF-α 1.27

(0.21) 1.20

(0.12) 3.06

(0.25) 3.06

(0.34) 1.62

(0.12)


Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data was considered as significantly different when P < 0.05. LLD = Less than lowest detectable measure. For IFN-γ, this was 0.4 pg mL-1.

162

Table 6.4D Cytokines expressed by H357 (pg mL-1) incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 24 h.

Cytokines H357 (pg mL-1)

NE ALC3 AN SM TRI

IL-2 0.21

(0.08) 1.28

(0.16) 1.26

(0.27) 1.91

(0.50) 0.78

(0.10)

IL-4 0.05

(0.02) 0.21

(0.02) 0.23

(0.05) 0.20

(0.04) 0.14

(0.01)

IL-6 39.09

(13.88) 505.02 (29.46)

510.42 (128.99)

947.82 (155.99)

279.80 (3.99)

IL-8 13.59 (2.90)

256.22 (7.96)

281.14 (70.49)

137.61 (32.06)

73.70 (2.40)

IL-10 0.58

(0.04) 0.74

(0.06) 0.73

(0.06) 0.61

(0.07) 0.70

(0.02)

GM-CSF 6.82

(0.59) 11.68 (0.43)

11.68 (1.96)

11.60 (3.63)

10.23 (0.43)

IFN-γ LLD 11.54 (1.74)

10.52 (2.56)

14.30 (5.18)

6.53 (0.31)

TNF-α 1.27

(0.00) 6.95

(0.36) 6.03

(1.55) 7.78

(1.13) 3.14

(0.34)


Data was the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data was considered as significantly different when P < 0.05. LLD = Less than lowest detectable measure. For IFN-γ, this was 0.4 pg mL-1.

163

6.4.6.1 Interleukin 2 (IL-2)

After 2 h, only OKF6 incubated with SM effluent caused a significant increase

(2.24-fold) of IL-2 expression when compared to NE (P < 0.05; Table 6.4A; Figure

6.2A). There was no significant difference of IL-2 expression in OKF6 after 24 h

incubation in all suspension when compared to NE (P > 0.05; Table 6.4B). An

increase of IL-2 was observed when OKF6 was incubated in biofilm effluent (58.7%

to 157.3%) after 24 h incubation compared to 2 h where no significant difference was

observed when compared to NE (P > 0.05; Table 6.5A).

After 2 h, H357 incubated with AN and SM effluents showed a significant

increase of IL-2 (3.89-fold and 4.64-fold respectively), compared to NE (P < 0.05;

Table 6.4C; Figure 6.2A). After 24 h, H357 incubated in ALC3 (6.65-fold), AN (7.11-

fold), SM (10.65-fold) and TRI (4.29-fold) effluents showed significantly increased

expression of IL-2 compared to NE (P < 0.05; Table 6.4D; Figure 6.2A). Increased

expression of IL-2 was observed when H357 was incubated in biofilm effluent

(110.5% to 451.6%) after 24 h incubation compared to 2 h, with the largest two

increases observed when incubated in ALC3 (451.6%) and TRI (342.1%) effluents

compared to NE (P < 0.05; Table 6.5B).

164

Figure 6.2A Fold change of IL-2 expression by OKF6 (left) and H357 (right) cells incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h. Fold changes were the ratio of mean cytokine expression at 2 h and 24 h of biofilm effluents (ALC3, AN, SM and TRI), to NE. Bars are the SD. The study was conducted in three biological replicates with each replicate consisting of three technical replicates.

0.002.004.006.008.00

10.0012.0014.0016.0018.00

ALC3 AN SM TRI

Fold

cha

nge

IL-2

OKF6 2 h

OKF6 24 h

0.002.004.006.008.00

10.0012.0014.0016.0018.00

ALC3 AN SM TRI

Fold

cha

nge

IL-2

H357 2 h

H357 24 h

165


There was no statistically significant change of IL-4 expression in OKF6 after

2 h and 24 h incubation with all biofilm effluent compared to NE (P > 0.05; Table

6.4A; Table 6.4B). Increased expression of IL-4 was observed when OKF6 was

incubated in biofilm effluent (78.6% to 283.3%) after 24 h incubation compared to 2 h

with no significant difference observed when compared to NE (P > 0.05; Table 6.5A).

After 2 h, H357 incubated in AN (1.96-fold) and SM (2.49-fold) effluent

showed significantly increased expression of IL-4 compared to NE (P < 0.05; Table

6.4C; Figure 6.2B). After 24 h, H357 incubated in ALC3 (4.50-fold), AN (4.92-fold),

SM (4.48-fold) and TRI (3.05-fold) effluents showed significant increases in IL-4

compared to NE (P < 0.05; Figure 6.6B). Increased expression of IL-4 was observed

when H357 was incubated in biofilm effluent (50.6% to 282.1%) after 24 h incubation

compared to 2 h with significant increases observed when incubated in ALC3 and

TRI effluent compared to NE (P < 0.05; Table 6.5B).

166

Figure 6.2B Fold change of IL-4 expression by OKF6 (left) and H357 (right) cells incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h. Fold changes are the ratio of mean cytokine expression at 2 h and 24 h of biofilm effluents (ALC3, AN, SM and TRI), to NE. Bars are the SD. The study was conducted in three biological replicates with each replicate consisting of three technical replicates.

0.001.002.003.004.005.006.007.008.00

ALC3 AN SM TRI

Fold

cha

nge

IL-4

OKF6 2 h

OKF6 24 h

0.001.002.003.004.005.006.007.008.00

ALC3 AN SM TRI

Fold

cha

nge

IL-4

H357 2 h

H357 24 h

167


After 2 h, only OKF6 incubated with SM effluent caused significantly

increased (1.99-fold) IL-6 expression when compared to NE (P < 0.05; Table 6.4A;

Figure 6.2C). Meanwhile, after 24 h incubation, OKF6 cells incubated in TRI effluent

was shown to significantly increase expression of IL-6 when compared to NE (2.28-

fold; P < 0.05; Table 6.4B; Figure 6.2C). Increased expression of IL-6 was observed

when OKF6 was incubated in biofilm effluent (3.98% to 13.75%) after 24 h

incubation compared to 2 h with no significant difference observed when compared to

NE (P > 0.05; Table 6.5A).

After 2 h, H357 incubated in AN (15.76-fold), SM (19.87-fold) and TRI (3.30-

fold) effluent showed significantly increased expression of IL-6 compared to NE (P <

0.05; Table 6.4C; Figure 6.2C). After 24 h, H357 grown in ALC3 (13.86-fold), AN

(14.79-fold), SM (25.78-fold) and TRI (7.78-fold) effluents had significantly

increased expression of IL-6 compared to NE (P < 0.05; Table 6.4D; Figure 6.2C).

Increased expression of IL-6 was observed when H357 was incubated in biofilm

effluents (166.2% to 3169.6%) after 24 h incubation, compared to 2 h where

significant increases were observed when incubated in ALC3 (3169.6%) and TRI

(611.1%) effluents compared to NE (P < 0.05; Table 6.5B).

168

Figure 6.2C Fold change of IL-6 expression by OKF6 (left) and H357 (right) cells incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h. Fold changes are the ratio of mean cytokine expression at 2 h and 24 h of biofilm effluents (ALC3, AN, SM and TRI), to NE. Bars are the SD. The study was conducted in three biological replicates with each replicate consisting of three technical replicates.

0.005.00

10.0015.0020.0025.0030.0035.0040.00

ALC3 AN SM TRI

Fold

cha

nge

IL-6

OKF6 2 h

OKF6 24 h

0.005.00

10.0015.0020.0025.0030.0035.0040.00

ALC3 AN SM TRI

Fold

cha

nge

IL-6

H357 2 h

H357 24 h

169


After 2 h, OKF6 incubated with ALC3 effluent caused a significant decrease

(0.72-fold) of IL-8 expression when compared to NE (P < 0.05; Table 6.4A; Figure

6.2D). After 24 h, TRI effluent was shown to significantly increase cells expressing

IL-8 (1.91-fold) compared to cells that were grown in NE (P < 0.05; Table 6.4B;

Figure 6.2D). Increased expression of IL-8 was observed when OKF6 was incubated

in biofilm effluent (224.7% to 662.6%) after 24 h incubation compared to 2 h where

only TRI effluent (662.6%) showed a significant increase when compared to NE (P <

0.05; Table 6.5A).

After 2 h, H357 incubated in AN (2.79-fold) and SM (2.07-fold) effluent

showed significantly increased expression of IL-8 compared to NE (P < 0.05; Table

6.4C; Figure 6.2D). After 24 h, H357 incubated in ALC3 (19.45-fold), AN (21.70-

fold), SM (10.71-fold) and TRI (5.60-fold) effluent was shown to increase IL-8

significantly compared to NE (P < 0.05; Table 6.4D; Figure 6.2D). Increased

expression of IL-8 was observed when H357 was incubated in biofilm effluent

(100.2% to 783.9%) after 24 h incubation compared to 2 h with significant increases

observed when incubated in all biofilm effluents compared to NE (P < 0.05; Table

6.5B).

170

Figure 6.2D Fold change of IL-8 expression by OKF6 (left) and H357 (right) cells incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h. Fold change is the ratio of mean cytokine expression at 2 h and 24 h of biofilm effluents (ALC3, AN, SM and TRI), to NE. Bars are the SD. The study was conducted in three biological replicates with each replicate consisting of three technical replicates.

0.00

5.00

10.00

15.00

20.00

25.00

30.00

35.00

ALC3 AN SM TRI

Fold

cha

nge

IL-8

OKF6 2 h

OKF6 24 h

0.00

5.00

10.00

15.00

20.00

25.00

30.00

35.00

ALC3 AN SM TRI

Fold

cha

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IL-8

H357 2 h

H357 24 h

171


There was no significant difference of OKF6 expressing IL-10 when incubated

in all biofilm effluents compared to NE after 2 h and 24 h (P > 0.05; Table 6.4A;

Table 6.4B). Increased expression of IL-10 was observed when OKF6 was incubated

in biofilm effluent (8.3% to 18.4%) after 24 h incubation compared to 2 h where no

biofilm effluents showed a significant difference when compared to NE (P > 0.05;

Table 6.5A).

ALC3 (1.15-fold), AN (1.20-fold), SM (1.23-fold) and TRI (1.17-fold)

effluents have been shown to increase H357 expressing IL-10 significantly when

compared to NE after 2 h incubation (P < 0.05; Table 6.4C; Figure 6.2E). After 24 h

incubation, H357 in ALC3 (1.28-fold), AN (1.27-fold) and TRI (1.21-fold) effluents

have been shown to increase expression of IL-10 significantly when compared to NE

(P < 0.05; Table 6.4D; Figure 6.2E). Increased expression of IL-10 was observed

when H357 was incubated in ALC3, AN and TRI effluents (3.5% to 10.9%) after 24 h

incubation compared to 2 h and a 15.1% decrease for OKF6 cells grown in SM. No

biofilm effluent showed significant differences when compared to NE (P > 0.05;

Table 6.5B).

172

Figure 6.2E Fold change of IL-10 expression by OKF6 (left) and H357 (right) cells incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h. Fold change is the ratio of mean cytokine expression at 2 h and 24 h of biofilm effluents (ALC3, AN, SM and TRI), to NE. Bars are the SD. The study was conducted in three biological replicates with each replicate consisting of three technical replicates.

0.000.200.400.600.801.001.201.401.60

ALC3 AN SM TRI

Fold

cha

nge

IL-10

OKF6 2h

OKF6 24 h

0.000.200.400.600.801.001.201.401.60

ALC3 AN SM TRI

Fold

cha

nge

IL-10

H357 2h

H357 24 h

173

6.4.6.6 Granulocyte-macrophage colony-stimulating factor (GM-

CSF)

After 2 h incubation, OKF6 grown in TRI effluent (0.89-fold) exhibited

significantly decreased expression of GM-CSF when compared to NE (P < 0.05;

Table 6.4A; Figure 6.2F). After 24 h incubation, OKF6 incubated in TRI effluent

(1.40-fold) exhibited significantly increased expression of GM-CSF compared to NE

(P < 0.05; Table 6.4B; Figure 6.2F). An increase of GM-CSF was observed when

OKF6 was incubated in biofilm effluent (54.5% to 141.4%) after 24 h incubation

compared to 2 h, where TRI effluent (141.1%) showed a significant increase when

compared to NE (P < 0.05; Table 6.5A).

After 2 h incubation, H357 incubated in ALC3 (1.17-fold), AN (1.31-fold) and

SM (1.29-fold) were shown to increase GM-CSF significantly compared to cells

incubated in NE (P < 0.05; Table 6.4C; Figure 6.2F). Incubation of H357 for 24 h in

biofilm effluent of ALC3 (1.72-fold), AN (1.71-fold), SM (1.68-fold) and TRI (1.51-

fold) have been shown to increase expression of GM-CSF significantly compared to

cells that were incubated in NE (P < 0.05; Table 6.4D; Figure 6.2F). Increased

expression of GM-CSF was observed when H357 was incubated in all biofilm

effluents (37.7% to 54.7%) after 24 h incubation compared to 2 h, where ALC3

(54.7%) and TRI (45.4%) effluent showed a significant difference when compared to

NE (P < 0.05; Table 6.5B).

174

Figure 6.2F Fold change of GM-CSF expression by OKF6 (left) and H357 (right) cells incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h. Fold change is the ratio of mean cytokine expression at 2 h and 24 h of biofilm effluents (ALC3, AN, SM and TRI), to NE. Bars are the SD. The study was conducted in three biological replicates with each replicate consisting of three technical replicates.

0.00

0.50

1.00

1.50

2.00

2.50

ALC3 AN SM TRI

Fold

cha

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GM-CSF

OKF6 2h

OKF6 24 h

0.00

0.50

1.00

1.50

2.00

2.50

ALC3 AN SM TRI

Fold

cha

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GM-CSF

H357 2h

H357 24 h

175

6.4.6.7 Interferon gamma (IFN-γ)

IFN-γ expressed by OKF6 cells was not detected after 2 h incubation (Table

6.4A). However, after 24 h incubation only OKF6 cells that were incubated in NE,

SM and TRI effluents were detected expressing IFN-γ (Table 6.4B). Increased

expression of IFN-γ was observed when OKF6 was incubated in biofilm effluent

(0.0% to 739.2%) after 24 h incubation compared to 2 h with only TRI (739.2%)

effluent showing a significant increase when compared to NE (P < 0.05; Table 6.5A).

After 2 h incubation, IFN-γ was only detected when H357 was incubated in

AN, SM and TRI effluents, whereas after 24 h, IFN-γ was only detected when H357

was incubated in ALC3, AN, SM and TRI effluents. Increased expression of IFN-γ

was observed when H357 was incubated in biofilm effluent (192.0% to 2785.8%)

after 24 h incubation compared to 2 h where all biofilm effluents showed significant

increases when compared to NE (P < 0.05; Table 6.5B).

176

6.4.6.8 Tumour necrosis factor alpha (TNF-α)

There was no change of TNF-α expressed by OKF6 after 2 h and 24 h

incubation in all suspensions compared to NE (P > 0.05; Table 6.4A; Table 6.4B).

Increased expression of TNF-α was observed when OKF6 was incubated in all

biofilm effluents (15.8% to 78.7%) after 24 h incubation compared to 2 h with no

biofilm effluent showing significant difference when compared to NE (P > 0.05;

Table 6.5A).

After 2 h, H357 incubated in AN (2.44-fold) and SM (2.49-fold) effluents

exhibited increased expression of TNF-α compared to NE significantly (P < 0.05;

Table 6.4C; Figure 6.2G). In addition, H357 incubated in ALC3 (5.47-fold), AN

(4.75-fold), SM (6.13-fold) and TRI (2.47-fold) effluent was shown to increase

expression of TNF-α compared to NE significantly after 24 h (P < 0.05; Table 6.4D;

Figure 6.2G). Increased expression of TNF-α was observed when H357 was

incubated in biofilm effluent (94.6% to 483.0%) after 24 h incubation compared to 2 h

where all biofilm effluents showed a significant increase when compared to NE (P <

0.05; Table 6.5B).

177

6.4.6.9 Overall

Taken together, these results demonstrate that the expression of IL-2, IL-4, IL-

6, IL-8, IL-10, TNF-α, GM-CSF and IFN-γ were expressed in a cell type, time-

dependent and biofilm effluent-specific manner where the majority of biofilm

effluents induced the expression of cytokines in the malignant cell line (H357) and

not in the normal epithelial cell line (OKF6). Furthermore, the effluent of C. albicans

(ALC3) and polymicrobial biofilms (TRI) was shown to significantly increase

cytokine production by H357 cells after 24 h compared to incubation in artificial

saliva (NE).

178

Figure 6.2G Fold change of TNF-α expression by OKF6 (left) and H357 (right) cells incubated with 80% serum free medium containing 20% of non-effluent ASM (NE), C. albicans (ALC3), A. naeslundii (AN), S. mutans (SM), and polymicrobial (TRI) biofilm effluents at 37 °C, 5% CO2 for 2 h and 24 h. Fold change is the ratio of mean cytokine expression at 2 h and 24 h of biofilm effluents (ALC3, AN, SM and TRI), to NE. Bars are the SD. The study was conducted in three biological replicates with each replicate consisting of three technical replicates.

0.001.002.003.004.005.006.007.008.00

ALC3 AN SM TRI

Fold

cha

nge

TNF-α

OKF6 2h

OKF6 24 h

0.001.002.003.004.005.006.007.008.00

ALC3 AN SM TRI

Fold

cha

nge

TNF-α

H357 2h

H357 24 h

179

Table 6.5A Percentage difference of cytokines expressed by OKF6 incubated in NE, ALC3, AN, SM, and TRI effluents at 37 °C, 5% CO2 between 2 h and 24 h incubation.

Cytokines OKF6 (%)

NE ALC3 AN SM TRI

IL-2 288.0

(299.0) 129.1 (82.9)

112.2 (110.5)

58.7 (22.0)

157.3 (70.7)

IL-4 225.0

(288.3) 78.6 (6.2)

280.0 (113.8)

178.6 (138.9)

283.3 (146.5)

IL-6 415.4 (137.5)

570.8 (279.7)

633.2 (633.5)

218.8 (57.2)

800.9 (380.2)

IL-8 325.6 (67.8)

408.5 (73.7)

332.2 (162.7)

224.7 (145.3)

662.6 (135.6)

IL-10 8.7

(8.1) 12.4 (3.4)

8.3 (17.2)

14.7 (10.6)

18.4 (9.5)

GM-CSF 53.8 (21.3)

54.5 (1.9)

83.5 (35.4)

60.3 (5.4)

141.4 (23.2)

IFN-γ* 112.5

(194.9) 0.0

(0.0) 214.2

(370.9) 225.0

(194.9) 739.2

(167.4)

TNF-α 51.6

(20.4) 15.8

(16.6) 26.0 (8.5)

41.5 (26.4)

78.7 (31.0)


Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data was considered as significantly different when P < 0.05. Positive results indicated increased percentage of cytokines synthesised by OKF6 between 2 h and 24 h incubation. *The measure of LLD (0.4 pg mL-1) was used to assess percentage difference.

180

Table 6.5B Percentage difference of cytokines expressed by H357 incubated in NE, ALC3, AN, SM, and TRI effluents at 37 °C, 5% CO2 between 2 h and 24 h incubation.

Cytokines H357 (%)

NE ALC3 AN SM TRI

IL-2 46.3 (77.9)

451.6 (59.4)

110.5 (8.0)

177.1 (64.9)

342.1 (119.0)

IL-4 -3.6 (68.0)

282.1 (67.6)

106.1 (43.0)

50.6 (39.7)

151.2 (63.9)

IL-6 243.0

(187.4) 3169.6

(1166.9) 166.2 (37.6)

303.2 (21.4)

611.1 (20.8)

IL-8 -35.1 (25.2)

783.9 (196.3)

377.3 (145.5)

210.7 (72.9)

100.2 (45.5)

IL-10 0.0

(6.3) 10.9 (5.6)

5.3 (8.3)

-15.1 (7.5)

3.5 (3.8)

GM-CSF 5.4

(6.9) 54.7 (1.2)

38.0 (20.5)

37.7 (36.3)

45.4 (6.5)

IFN-γ* 0.0

(0.0) 2785.8 (434.5)

276.7 (55.5)

192.0 (118.8)

681.4 (698.3)

TNF-α 1.9 (17.1)

483.0 (62.8)

96.5 (44.0)

154.5 (30.4)

94.6 (27.5)


Data is the mean of three separate experiments (SD are given in parentheses). The study was conducted in three biological replicates with each replicate consisting of three technical replicates. Data was considered as significantly different when P < 0.05. Negative results indicated a decrease of cytokines expressed by H357, whereas positive results indicated an increased percentage between 2 h and 24 h incubation. *The measure of LLD (0.4 pg mL-1) was used to assess percentage difference.

181

6.5 Discussion

Carcinomas are characterised by the ability of malignant cells to invade

underlying connective tissue and to migrate, forming metastasis at distant sites (Lyons

and Jones, 2007). One of the important properties of cells in the progression of oral

cancer is the ability to adhere to extracellular matrix (ECM). To our knowledge, this

is the first study undertaken to elucidate the role of biofilm effluent of C. albicans, A.

naeslundii and S. mutans in the adhesion to ECM molecules, regulation of epithelial-

mesenchymal transition (EMT) and the expression of pro-inflammatory cytokines by

OKF6 (normal epithelial cell line) and H357 (OSCC cell line).

We have shown distinct variation in the ability of OKF6 and H357 to adhere

to ECM components when incubated in biofilm effluent, with H357 exhibiting

increased adhesion to ECM molecules, particularly collagen IV and laminin I. This

was most pronounced when H357 was incubated with C. albicans biofilm effluent.

Further, increased adhesion was also observed when H357 was incubated in

polymicrobial biofilm effluent compared to NE, thus supporting the hypothesis that

the adhesion of cells to ECM molecules is biofilm effluent-dependent and the

polymicrobial biofilm effluent increased the malignant phenotype, suggesting that

biofilm effluent is an oral cancer promoter rather than initiator. The variability of

adhesion of OKF6 and H357 cells to ECM components has been suggested to be due

to the alteration of cells’ integrins following interaction with biofilm effluent (Lyons

and Jones, 2007). Integrins are known as the largest family of cell adhesion

molecules that consist of multiple combinations of α- and β-subunits (Van Waes and

Carey, 1992; Lyons and Jones, 2007). Proteins synthesised by microorganisms such

as proteases from C. albicans (Karkowska-Kuleta et al., 2009) and Bcl-2 family

proteins from bacteria (Khajuria and Metgud, 2015) have been shown to induce

182

alteration of integrins of epithelial cells. Furthermore, the α1-subunit of integrins that

preferentially binds to collagen IV, and the α3-subunit that acts as a receptor for

laminins, have been shown to change during interaction with the proteins of

microorganisms (Lyons and Jones, 2007). The results of the present study support

these previous findings, as biofilm effluent, particularly ALC3, induced enhanced

adhesion of H357 to collagen IV and laminin I and polymicrobial biofilm effluent

enhanced adhesion of the OSCC cells to fibronectin, collagen IV and laminin I. It is

important to note that the oral environment comprises a large range of polymicrobial

organisms, including C. albicans, A. naeslundii and S. mutans that may alter the

integrins of malignant cells. Subsequent triggering of a spectrum of signals involved

in the process of growth and proliferation may promote oral carcinogenesis in a

paracrine fashion (Carter et al., 1990; Ono et al., 1999; Shinohara et al., 1999;

Meurman, 2010).

The EMT assay of the present study revealed a paradoxical effect with both an

enhanced and a diminished malignant phenotype being observed concurrently. These

results support the hypothesis that EMT is biofilm effluent-dependent. EMT is

indicated by the increase of vimentin and the decrease of E-cadherin expression from

epithelial cells (Lang et al., 2002; Hugo et al., 2007; Onder et al., 2008; Nijkamp et

al., 2011; Yadav et al., 2011). The presence of a stimulator, such as a cell wall

extract of the oral bacterium Fusobacterium nucleatum has been shown to increase

the expression of vimentin and decrease the expression of E-cadherin in OSCC cell

lines HN008 and HN5 (Krisanaprakornkit and Iamaroon, 2012). EMT has been

reported to be involved in the increased resistance of malignant cells to apoptosis

regulator molecules (Maestro et al., 1999; Vega et al., 2004) indicating the important

183

role of EMT in metastasis of malignant cells (Kang and Massagué, 2004; Yang et al.,

2004; Radisky, 2005).

The result in the present study of an increase in E-cadherin expression by

OKF6 cells after 24 h incubation with mono-cultured biofilm effluent is likely

indicative of adhesion, colonisation, internalisation and potentially invasion. An

increase of E-cadherin expression has been suggested to be a strategy of colonisation

of C. albicans, A. naeslundii and S. mutans to oral epithelial cells (Delva and

Kowalczyk, 2009). Furthermore, cadherins have been reported to form a route for the

internalisation of bacteria and yeast into the epithelial cells during oral thrush (Phan et

al., 2005; Delva and Kowalcyzyk, 2009). In addition, Als3 protein synthesised by C.

albicans has been thought to mimic cadherin-cadherin binding, thus initiating the

invasion of the yeast to the oral epithelial cells (Phan et al., 2005). The observed

increase in expression of E-cadherin by the H357 cell line at an early time point (2 h)

in the present study may well be indicative of this early colonisation and invasion.

This however did not persist at later (24 h) time points in this cell line. Interestingly,

the normal cell line OKF6 showed an increase in the expression of E-cadherin at this

later stage (24 h), a finding likely to indicate the time required for normal cells to

allow for internalisation or invasion.

The present study has shown significant variability of cytokine expression by

OKF6 and H357 when incubated with the effluent from different biofilms. These

results support the hypotheses that cytokine expression by OKF6 and H357 is biofilm

effluent-dependent and that C. albicans, A. naeslundii, S. mutans and polymicrobial

biofilm effluents increase the malignant phenotype and can act as an oral cancer

promoter. The variability of cytokines expressed by the cells may represent the

homeostatic mechanism of innate immunity at mucosal tissues responding to the

184

presence of biofilm effluent (Steele and Fidel, 2002). Furthermore, oral epithelial

cells increased synthesis of IL-6, IL-8, IL-10 and TNF-α when incubated with pre-

cultured C. albicans medium compared to medium alone (Steele and Fidel, 2002).

Similarly, incubation of endothelial cells with C. albicans conditioned medium has

been shown to increase the expression of IL-6 and IL-8 after 12 h incubation

compared to 8 h (Filler et al., 1996). These results are consistent with our finding,

where H357 was observed to increase the same pro-inflammatory cytokines after 24 h

incubation with ALC3 and polymicrobial biofilm effluents.

The increase of cytokines expressed by H357 is thought to be due to the

presence of proteins glycosylated with N- or O-linked mannosyl residues, β-glucans

and chitins from the C. albicans cell wall, as well as the presence of SAPs (Dongari-

Bagtzoglou and Kashleva, 2003a; Mostefaoui et al., 2004; Schaller et al., 2005).

These proteins have been previously shown to increase the expression of IL-6, IL-8,

IL-10, GM-CSF, TNF-α and IFN-γ from epithelial cells and human mono-nuclear

cells that were incubated in C. albicans conditioned medium compared to NE

(Dongari-Bagtzoglou and Kashleva, 2003a; Mostefaoui et al., 2004; Schaller et al.,

2005; Netea et al., 2006). The increased expression of pro-inflammatory cytokines,

particularly IL-6, IL-8 and GM-CSF are important in inflammation as well as

tumorigenesis of malignant cells (Kitadai et al., 2000). IL-6 has been reported to

have an anti-apoptotic effect on malignant cells (Burgdorf et al., 2009). In addition,

direct autocrine tumour promoting effects of IL-6 have been demonstrated in multiple

myeloma by both increasing proliferation and preventing apoptosis (Thaler et al.,

1994; Frassanito et al., 2001). GM-CSF has been previously shown to be a tumour

cell stimulator (Burgdorf et al., 2009), and IL-8 has been reported to be involved in

carcinogenesis by inducing angiogenesis (Lin and Karin, 2007; Fantini and Pallone,

185

2008). The expression of IL-8 by human carcinoma cells has been shown to directly

correlate with tumour vascularity and disease progression (Kitadai et al., 2000). Thus

the results of the present study clearly demonstrate the role of microbial effluent in

promoting oral carcinogenesis.

The present study has shown that malignant oral epithelial cells (H357) were

observed to increase the expression of many more cytokines than normal epithelial

cells (OKF6) when incubated with biofilm effluent. The tumour microenvironment is

rich with cytokines and other inflammatory mediators that have been shown to

influence the growth of cancer cells (Balkwill and Mantovani, 2001; Balkwill, 2004;

Seruga et al., 2008). Furthermore, high expression of pro-inflammatory cytokines

such as TNF-α in various human cancers, such as breast, prostate, bladder and

leukaemia, has also been reported suggesting an important role of cytokines in oral

cancer progression (Kundu and Surh, 2008). Further, several preclinical studies have

shown a significant increase of TNF-α in gastric lesions and inflamed colonic mucosa

in patients with Helicobacter pylori infection (Noach et al., 1994; Noguchi et al.,

1998). The results of the present study indicate that, via the enhanced expression of

cytokines by malignant oral epithelial cells, oral microbial biofilms and in particular

those containing C. albicans, could potentially act as promoters of oral cancer

progression. Previous study has shown that Candida spp. particularly C. albicans

were isolated from 30% of patients with cancerous lesion and 32% of patients with

precancerous lesion between 2007 and 2009 in Naples, Italy (Galle et al., 2013). It is

suggested that many cancer promoters exist in the oral cavity which C. albicans

infection could be one of them.

186

6.6 Conclusion

Biofilm effluent promoted oral carcinogenesis by increasing the adhesion of

an oral squamous cell carcinoma cell line to extracellular matrix molecules and the

increase of pro-inflammatory cytokine expression. This tumour growth promoting

effect of oral microbial biofilms may be occurring at either the early stages in oral

carcinogenesis or perhaps as an enhancement of later tumour progression.

Nevertheless, the oral microbial biofilm promotion of oral cancer has profound

clinical implications and requires further elucidation of the exact mechanism by

which it occurs, as well as confirmation of its occurrence in vivo.

187

CHAPTER 7

DISCUSSION AND CONCLUSION

188

7.1 Discussion

Cancer has been the leading cause of death in developed countries and second

in the developing countries (Jemal et al., 2011), with oral squamous cell carcinoma

(OSCC) accounting for more than 90% of malignancies originating from the oral

mucosa (Casiglia and SB, 2001; Johnson et al., 2011). The risk factors that lead to

OSCC include heavy alcohol consumption, tobacco smoking, unhealthy diet, poor

oral hygiene and microbial infections (Hooper et al., 2009; Chocolatewala et al.,

2010; Meurman, 2010; Rajeev et al., 2012; Khajuria and Metgud, 2015).

Yeast and bacterial infections have been widely suggested to have a causal

role in oral cancer (Meurman, 2010; Rajeev et al., 2012; Khajuria and Metgud, 2015).

Yeast such as C. albicans carriage has been found to correlate with the presence of

oral epithelial dysplasia (McCullough et al., 2002). Bacteria such as S. mutans have

been shown to synthesise alcohol dehydrogenase. This enzyme is reported to convert

alcohol to carcinogenic acetaldehyde (Kurkivuori et al., 2007; Hooper et al., 2009).

In addition, A. naeslundii has been shown to colonise the oral cavity of cancer

patients more than in healthy individuals (Nagy et al., 1998; Pushalkar et al., 2011).

The promotion of oral carcinogenesis by microorganisms begins from the

interaction of the oral microbiome (Kolenbrander, 2000; Min and Rickard, 2009). To

assess polymicrobial interactions, a co-aggregation study of eight strains of C.

albicans with A. naeslundii and S. mutans was conducted (Section 2.2). The present

study has shown that co-aggregation was C. albicans strain-dependent with the

majority of the yeast grown in RPMI-1640 (hyphal growth). When co-incubated with

S. mutans and A. naeslundii either alone or in combination, variable co-aggregation

resulted (Chapter 3). Variability of co-aggregation was also observed from ASM-

grown C. albicans (yeast growth) strains that were co-incubated with S. mutans and

189

A. naeslundii. It can be postulated that the observed variability in co-aggregation may

be related to that specific strain’s ability to produce both non-specific (adhesins) and

specific (lectin-saccharide) cell surface receptors (Kolenbrander and Williams, 1981;

McIntire et al., 1982; Grimaudo, 1996; Rickard et al., 2003; Rosen and Sela, 2006;

Ledder et al., 2008). The observed variability of co-aggregation in C. albicans may

also be attributable to different strains having different abundances of specific

molecules such as Farnesol, that have been suggested to have a role in polymicrobial

interactions of C. albicans to oral bacteria (Morales and Hogan, 2010). The findings

of the present study supported the hypothesis that auto-aggregation and co-

aggregation are C. albicans strain-dependent, and rejected the null hypothesis that

auto-aggregation and co-aggregation are not C. albicans strain-dependent.

Oral microorganisms are required to develop polymicrobial biofilms on the

oral substrata in order to potentially promote oral carcinogenesis (Chapter 4). To

determine the effect of polymicrobial interaction of C. albicans, A. naeslundii and S.

mutans to biofilm formation, the biofilm biomass and metabolic activity were

assessed using crystal violet and XTT assays, respectively (Section 2.4). The present

study has shown a variation of biofilm biomass and metabolic activity between C.

albicans strains based on the classification proposed by Marcos-Zambrano et al.

(2014). Strain variability of C. albicans is present in the oral cavity of different

individuals (Hellstein et al., 1993; Kleinegger et al., 1996). C. albicans strains

isolated from HIV-infected patients are reported to produce higher levels of aspartic

proteinases (SAPs), that are important in the formation of C. albicans biofilm,

compared to the strains isolated from uninfected patients (Morales and Hogan, 2010;

Arzmi et al., 2012).

190

The biofilm biomass (Chapter 4 and Chapter 5) and metabolic activity were

shown to vary with microbial interactions (Chapter 4) and be morphology-dependent

(Chapter 4). The biofilm biomass in static biofilms of the majority of RPMI-1640

grown C. albicans (hyphal form) was observed to increase in the presence of bacteria

compared with mono-cultured C. albicans. A. naeslundii and S. mutans have been

shown to bind to C. albicans through its mannose-containing surface protein

(Kolenbrander and Williams, 1981; McIntire et al., 1982; Grimaudo et al., 1996;

Rickard et al., 2003; Rosen and Sela, 2006; Ledder et al., 2008). This interaction has

been reported to induce the formation of extracellular polysaccharide, thus promoting

the adherence of the late colonisers to form a complex of polymicrobial biofilm

(Nyvad and Kilian, 1987; Grimaudo et al., 1996; Li et al., 2004). The variability of

metabolic activity in polymicrobial biofilms suggests that these microorganisms may

be interacting metabolically (Chapter 4). It is postulated that in the presence of A.

naeslundii, C. albicans increased mitochondrial dehydrogenase activity that in turn

increased the activity of succinate dehydrogenases of A. naeslundii.

The present study showed that S. mutans decreased the overall metabolic

activity in tri-cultured polymicrobial biofilms compared with the dual-cultured

polymicrobial C. albicans-A. naeslundii biofilms. Furthermore, ASM-grown C.

albicans biofilms were observed to have lower metabolic activity than those grown in

RPMI-1640 (Chapter 4), particularly mono-cultured biofilms. Candida spp. with low

metabolic activity are reported to be more invasive and associated with disease, while

conversely those with high activity are non-invasive (Kuhn et al., 2003; Tobudic et

al., 2012), and this may have a role in promoting oral carcinogenesis. These findings

on static biofilms of C. albicans, A. naeslundii and S. mutans supported our specific

hypotheses that polymicrobial biofilm formation is C. albicans strain- and

191

morphology-dependent, thus rejecting the null hypothesis that polymicrobial biofilm

formation is not C. albicans strain- and morphology-dependent.

The oral cavity has a constant salivary flow with the oral substrata coated with

saliva (Sánchez-Vargas et al., 2013; Marsh et al., 2016). These characteristics of the

oral environment have been shown to limit the colonisation of oral microorganisms

(de Almeida et al., 2008; Marsh et al., 2016) however by-products secreted by the

oral microbiome from the biofilm consortium to the oral cavity may have a role in

promoting oral cancer. Thus, the study of biofilms of C. albicans, A. naeslundii and

S. mutans was conducted in a flow-cell system to determine the effect of the

polymicrobial interaction of C. albicans, A. naeslundii and S. mutans on biofilm

formation in a flow environment (Section 2.8 to Section 2.11). Simultaneously,

effluent from these biofilms was collected for further assessment (Chapter 6). The

present study showed that C. albicans, A. naeslundii and S. mutans were able to form

polymicrobial biofilms on ASM-coated substrata, with the biofilm biomass of C.

albicans in the polymicrobial biofilms significantly decreased compared to the mono-

cultured biofilms (Chapter 5). These results were converse to that observed in static

biofilms (Chapter 4), indicating the important role of salivary flow in the oral cavity.

Furthermore, the biomass assessed by the crystal violet assay (Chapter 4) included

both extracellular polysaccharides and microorganisms, while the biomass assessed

by fluorescence in situ hybridisation (Chapter 5) determined biomass specifically for

the microorganisms within the biofilms. Mutualistic and antagonistic interactions

have been reported between C. albicans with S. mutans (McIntire et al., 1982;

Rickard et al., 2003; Rosen and Sela, 2006; Thein et al., 2006; Ledder et al., 2008),

and C. albicans with A. naeslundii (Millsap et al., 1999; Thein et al., 2006). C.

albicans has been shown to decrease adherence when co-cultured with S. mutans on

192

acrylic sheets in Gibbons and Nygaard culture medium (Barbieri et al., 2007).

Quorum-sensing molecule such as Farnesol synthesised by C. albicans during biofilm

formation has been reported to disrupt the membrane of S. mutans, as well as the

accumulation and polysaccharide contents of biofilms of the streptococci (Koo et al.,

2003; Jabra-Rizk et al., 2006). In addition, the metabolic products of A. naeslundii

have been reported to both inhibit and stimulate the biofilm formation of C. albicans

depending on the experimental methods employed (Gutiérrez and Benito, 2004; Thein

et al., 2006).

The present study has shown that the average thickness and maximum

thickness of polymicrobial biofilms was significantly increased when compared to the

mono-cultured C. albicans but not when compared to S. mutans. The increase of the

thickness is suggested to be due to the increase of extracellular polysaccharide in the

biofilm consortium when surrounded by 25% ASM containing sucrose (Koo et al.,

2010). Extracellular polysaccharides have been shown to provide attachment sites for

C. albicans that is critical for the colonisation of the microorganism to the oral

substrata (Harriott and Noverr, 2011). Therefore, the results of the present study

supported our third specific hypotheses that C. albicans, A. naeslundii and S. mutans

form polymicrobial biofilms, and that polymicrobial interactions affect colonisation

of oral microorganisms in a flow-cell environment. We therefore reject the null

hypotheses that C. albicans, A. naeslundii and S. mutans do not form polymicrobial

biofilms and that polymicrobial interactions do not affect colonisation of oral

microorganisms in a flow-cell environment.

Polymicrobial biofilms have been shown to produce high amounts of

extracellular polysaccharides with various by-products in the oral cavity due to

interaction with microbial colonies and the oral environment (Koo et al., 2003; Jabra-

193

Rizk et al., 2006). However the role of biofilm effluent, from C. albicans, A.

naeslundii and S. mutans grown as mono-cultured and polymicrobial biofilms, on oral

carcinogenesis remains unknown (Chapter 6). To assess the ability of microbial

biofilm effluent in promoting oral carcinogenesis, biofilm effluent obtained from C.

albicans, A. naeslundii, S. mutans and polymicrobial biofilms was assessed on both

normal epithelial cells (OKF6) and oral squamous cell carcinoma epithelial cells

(H357) (Section 2.13). An array of assays was conducted including: 1) an adhesion

assay to assess the ability of the cell lines to adhere to major important extracellular

matrix (ECM) molecules using the CytoSelect 48-well Cell Adhesion Assay ECM

Array kit (Section 2.15); 2) an epithelial to mesenchymal transition (EMT) assay to

assess the phenotypic changes of the cell lines by the detection of vimentin and E-

cadherin expression using flow cytometry (Section 2.17); and 3) the Bio-Plex to

assess the expression of pro-inflammatory cytokines from biofilm effluent-treated

cells (Section 2.18). The present study showed that the OSCC cell line, H357, when

incubated with C. albicans biofilm effluent increased adhesion to ECM molecules,

particularly collagen IV and laminin I (Chapter 6). Furthermore, an increased

adhesion to fibronectin, collagen IV and laminin I was also exhibited when H357 was

incubated in polymicrobial biofilm effluent (TRI). The increased adhesion of H357

cells to ECM components is likely to be due to the alteration of cell integrins while

interacting with the biofilm effluent (Lyons and Jones, 2007). Integrins are known as

the largest family of cell adhesion molecules that consist of multiple combinations of

α- and β-subunits (Van Waes and Carey, 1992; Lyons and Jones, 2007). Proteases of

C. albicans (Karkowska-Kuleta et al., 2009) have been shown to induce alteration of

integrins, particularly the α1-subunit of epithelial cell integrins that preferentially bind

to collagen IV and the α3-subunit that acts as a receptor for laminins (Lyons and

194

Jones, 2007), which are important in the development of oral cancer. The oral

environment comprises more diverse polymicrobial biofilms than that assessed in the

present study, and these may alter integrins of malignant cells more than observed in

the present study, thus promoting oral carcinogenesis in a paracrine fashion (Carter et

al., 1990; Ono et al., 1999; Shinohara et al., 1999; Meurman, 2010).

An increase of E-cadherin expression by OKF6 and H357 cells when

incubated with biofilm effluent is likely indicative of adhesion, colonisation,

internalisation and potentially invasion. An increase of E-cadherin expression has

been suggested to be a strategy of colonisation of C. albicans, A. naeslundii and S.

mutans to oral epithelial cells (Delva and Kowalczyk, 2009). Furthermore, cadherins

have been reported to form a route for the internalisation of pathogens into epithelial

cells during oral thrush (Phan et al., 2005; Delva and Kowalcyzyk, 2009). In

addition, Als3 protein synthesised by C. albicans has been thought to mimic cadherin-

cadherin binding, thus initiating the invasion of yeast into oral epithelial cells (Phan et

al., 2005). These findings indicate the strategy of C. albicans colonisation to the

surface and the subsurface of oral epithelial cells and is likely to promote

carcinogenesis as indicated by the expression of pro-inflammatory cytokines (Section

6.4.3).

We observed an increase of pro-inflammatory cytokine expression by oral

cancer cells, H357 when incubated with C. albicans and polymicrobial effluent. This

may be due to the presence of proteins glycosylated with N- or O-linked mannosyl

residues, β-glucans and chitins from the C. albicans cell wall, as well as the presence

of SAPs (Dongari-Bagtzoglou and Kashleva, 2003a; Mostefaoui et al., 2004; Schaller

et al., 2005; Netea et al., 2006). The increase of pro-inflammatory cytokines,

particularly IL-6, IL-8 and GM-CSF are important in inflammation as well as

195

tumorigenesis of malignant cells (Kitadai et al., 2000). IL-6 has been reported to have

an anti-apoptotic effect on malignant cells (Thaler et al., 1994; Frassanito et al., 2001;

Burgdorf et al., 2009). GM-CSF has been previously shown to be a tumour cell

stimulator (Burgdorf et al., 2009), whereas IL-8 has been reported to be involved in

carcinogenesis by inducing angiogenesis (Lin and Karin, 2007; Fantini and Pallone,

2008). Cytokines are also known to be secreted by the oral epithelial cells in order to

prevent carcinogenesis and as an action to overcome microbial colonisation.

However, research has also shown that the over-secretion of pro-inflammatory

cytokines may induce carcinogenesis. In the present study, we have found a

significant increase of pro-inflammatory cytokines synthesised by the OSCC cell line

compared to the normal epithelial cell line, indicating that biofilm effluent from C.

albicans grown as both mono-cultured and polymicrobial biofilms is promoting oral

cancer, but not inducing cancer (Budhu and Wang, 2006; Fantini and Pallone, 2008).

Thus, these findings on adhesion, EMT and cytokine expression assays

supports our hypothesis that oral epithelial cells have an enhanced malignant

phenotype, promoting oral carcinogenesis, when grown in the presence of

polymicrobial biofilms. Thus the null hypothesis that the presence of polymicrobial

biofilms does not enhance the malignant potential of oral epithelial cells was rejected.

196

7.2 Conclusion and future studies

The findings of the present study supported the overall hypothesis that


oral cancer by promoting carcinogenesis. Moreover, this carcinogenesis promoting

activity of polymicrobial biofilms is likely to be C. albicans strain-specific. This

tumour growth promoting effect of oral microbial biofilms may occur at either the

early stages in oral carcinogenesis or perhaps as an enhancement of later tumour

progression. Nevertheless, the oral microbial biofilm promotion of oral cancer has

profound clinical implications and requires further elucidation of the exact

mechanism by which this occurs, as well as in vivo confirmation of its occurrence.

Future in vivo studies of co-aggregation, biofilm formation of C. albicans, A.

naeslundii and S. mutans, and the role of biofilms in the expression of pro-

inflammatory cytokines are required to assess oral biological factors, such as salivary

flow and immunological components that may influence oral cancer promotion.

These in vivo studies will enhance our understanding of the interaction of

microorganisms in the oral cavity, a process likely to be critical in chronic infection

and potentially oral carcinogenesis. An assessement of the by-products secreted in

biofilm effluent are also required in order to understand what specific proteins lead to

the promotion of oral carcinogenesis. This has the potential for the development of

agents that counteract these proteins and then aid in the prevention of oral cancer.

197

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APPENDICES

EUROPEAN JOURNAL OF INFLAMMATION Vol. 12, no. 2, 227-235 (2014)

EDITORIAL

GAINING MORE INSIGHT INTO THE DETERMINANTS OF CANDIDA SPECIESPATHOGENICITY IN THE ORAL CAVITY

M.H. ARZMP,2, E. ALSHWAIMP, W.H.A. WAN HARUN\ F.ABDUL RAZAK\ F. FARINA5,6,M.J. MCCULLOUGHI and N. CIRILLOI,6

1Melbourne Dental School and Oral Health CRC, The University ofMelbourne, Melbourne, VIC,Australia; 2Department ofBasic Medical Sciences, Kulliyyah ofDentistry, International IslamicUniversity Malaysia, Kuantan, Pahang, Malaysia; 3Department ofRestorative Dental Sciences,College ofDentistry, University ofDammam, KSA; "Department ofOral Biology, Faculty of

Dentistry, University ofMalaya, Kuala Lumpur, Malaysia; 'Facultatea de Medicina si MedicinaDentara Titu Maiorescu, Bucharest, Romania; "Centroper l'Innovazione, la Ricerca, l'Istruzione,

la Salute (IRIS), Italy

Received June 26,2013 -Accepted March 17,2014

Candida infection (candidiasis) is potentially life threatening and can occur in almost all anatomicalsites, including the mouth. Candida species are in fact the most common fungal pathogens isolated fromthe oral cavity and frequently cause superficial infections such as oral candidiasis and denture-associatederythematous stomatitis. Whilst systemic dissemination of Candida from intraoral foci is rare andlargely due to severe deficits of the host immune defenses, the development of localized oral candidiasisis most commonly related to a variety of non-immune determinants such as Candida virulence factorsand permissive oral microenvironment. In particular, phenotypic switching and dental biofilm haveemerged as major determinants for the pathogenicity of Candida and are currently the subject ofintenseresearch. An understanding of the molecular aspects underlying the biological behavior of Candida willbe the key to the development of effective preventive as well as therapeutic measures for invasive andoral candidiasis.

Candida inhabits various parts of the humanbody including the epidermis, vagina, gastro-intes-tinal tract, nails and oral cavity (1). The diseasescaused by Candida became common in the late 19thand 20th centuries and its prevalence is still increas-ing worldwide as a result of multiple factors whichcan facilitate the conversion of its commensal levelto the parasitic level (2). According to Scardina etal. (3), the risk factors that enhance the severity of

a candidal infection can be found widely in patientswith impaired salivary gland, drug abusers, immuno-compromised, high carbohydrate diet, smoking hab-its and Cushing's syndrome. Candidal infection canoccur in almost all human organs. However, it is thesystemic infection that can be much more severe andmay lead to mortality. According to Leroy et al. (4),the mortality rate due to systemic infection of Can-dida is up to 60% and still increasing. The treatment

Key words: oral candidiasis, virulence factors, phenotypic switching, dental biofilm

Mailing address: Prof. Dr. Nicola Cirillo,Melbourne Dental School,The University of Melbourne,720 Swanston Street, Carlton,3053 Victoria, AustraliaTel./Fax: +61 03 9341 1597e-mail: [email protected] 227

1721-727X (2014)Copyright © by BIOLIFE, s.a.s.

This publication andlor article is for individual use only and may not be furtherreproduced without written permission from the copyright holder.

Unauthorized reproduction may result in financial and other penaltiesDISCLOSURE: ALLAUTHORS REPORT NO CONFLICTS OF

INTEREST RELEVANT TO THIS ARTICLE.

228 M.H. ARZMI ET AL.

of candidal infection can be difficult and most of thediagnoses can only be achieved by autopsy. With thecurrent incidence in Europe on the rise, there havebeen reports of a 5-fold increase in candidemia in thelast ten years (5).

Candida has been identified as the commonmember of the oral microflora and estimated to bepresent in approximately 40-60% of the generalpopulation. It can be present either as transient orpermanent colonizer in the oral cavity (6). It is alsorecognised as an opportunistic microorganism thathas the ability to cause oral diseases, such as oralcandidiasis (7).The most common oral condition caused by Can-

dida is oral candidiasis (8). In a most recent study,candidal infection was also associated with oral can-cer, burning mouth syndrome, endodontic diseasesand taste disorder (I). Candida albicans is the maincausative agent of oropharyngeal candidiasis. Re-searchers have, however, found that non-albicansspecies also contributed significantly to the develop-ment of oral candidiasis (9). Cases due to non-albi-cans species are increasing in number and this hasraised great concern to society.

Candida is identified to colonise several typesof host cells including epithelial, endothelial andphagocytic cells. In the oral cavity, Candida pre-fers to colonise several surfaces including the buc-cal and labial mucosa, dorsum or lateral borders oftongue, hard and soft palate regions, tooth surfacesand denture-bearing areas (10). This colonising abil-ity is contributed by factors including the ability oforal Candida to produce specific enzymes such asagglutinin-like proteins and integrin-like proteinsthat lead to the formation ofbiofilm on oral surfaces.In addition, other factors that influence the colonisa-tion of Candida are the reduction of salivary flow,low salivary pH, trauma, carbohydrate-rich diets andepithelial loss (11).Here we distinguish between two categories of

pathogenic determinants: extrinsic determinant, i.e.those provided by the host, which are permissive forgrowth and survival ofCandida; and intrinsic deter-minants, i.e. those related to the characteristics ofCandida species. Oral biofilm, which stricto sensubelongs to the first group, has been considered as anintrinsic determinant in that it relies on the ability ofCandida to interact with the oral microflora.

GROWTH REQUIREMENTOF CANDIDA SPECIES

Availability ofnutrientsCandida is a chemoheterotrophic organism that

requires carbon and nitrogen for growth. Accordingto Madigan and Martinko (12), the mutual interactionofcarbon and nitrogen is important in the metabolismof microorganisms. Carbohydrates are the mostreadily utilised form of carbon in both oxidativeand non-oxidative pathway. Thus, the presence ofcarbohydrates influences the colonisation ofCandidain the oral cavity. Certain carbohydrates, such assucrose and glucose, have been shown to increasethe adhesion potential of Candida albicans towardshard and soft surfaces of the oral cavity. Glucose isan acid promoter that leads to the reduction of pH inthe oral environment and as a consequence, activatesacid proteinases and phospholipases, which enhancethe adherence capability of Candida. In addition,the production of mannoprotein surface layer inan environment where glucose is present has beenshown to assist the adherence capability of Candidaincluding C. krusei in the oral cavity (7).

Candida has a nitrogen content of around 10%of their dry weight (7). The source of nitrogen isusually provided by organic compounds which canbe easily found in the oral environment. Nitrogenis also determined as the main stimulatory factorin yeast extract as it encourages bio-stimulation ofmicrobial growth.

Influence oforalfluidsSaliva provides moisture and helps in

lubricating the oral cavity. Furthermore, it alsoprovides indigenous organic constituents includingantimicrobial factors such as lysozyme, lactoferrin,calprotectin, lactoperoxidase, cystatins, histatins,VEGh and SLPI and chromogranin A which inhibitthe growth of oral pathogens (13). The presence ofcytokines, such as IL-17 and immunoglobulins, insaliva are also beneficial to the oral cavity as theyinhibit the dissemination of oral microorganismsespecially Candida species (14).Saliva also introduces the formation of a thin

film approximately 0.1 mm deep over all externalsurfaces in the oral cavity. The major role of thewhole saliva is to maintain the integrity of teeth by

European Journal of Inflammation 229

clearing off food debris and buffering the potentialdamaging acids produced by oral biofilm or dentalplaque. The chemical composition of secretions fromeach gland is different. Bicarbonate, phosphates andpeptides are examples of buffering agents in thesaliva that give normal saliva a mean pH of 6.75 to7.25 (7).The flow rate of saliva is under the influence of

circadian rhythms where the lowest flow rate hasoften been recorded during sleeping. Low flow rateof saliva reduces the protective function of salivaand increases the colonisation and developmentof microorganisms including Candida. Salivarycomposition is also affected by circadian rhythms, forexample the total concentration of protein in wholesaliva during resting time is estimated at 220 mg/100mL, whereas the total protein in stimulated saliva isestimated at 280 mg/10 mL. The difference in theamount of protein may affect the distribution of thenormal microflora in the oral cavity, as some proteinsare known to serve as receptors in the colonisation ofmicroorganisms to the saliva-coated surfaces of theteeth (7). Proteins and glycoproteins such as mucinin the saliva act as the primary source of nutrients forresident microflora including Candida. In additionto adherence, some proteins are also involved in thehost's defence mechanism by aggregating exogenousmicroorganisms, hence, facilitating their clearancefrom the mouth during swallowing or spitting.In addition to saliva, the gingival crevicular

fluid (GCF) in the oral cavity can also influence thecolonisation of oral Candida species. The flow ofGCF is slow at healthy sites but increases drasticallyat areas with gingivitis by 147% and up to 30-fold atareas with advanced periodontal diseases. GCF alsohas a role in the development of subgingival plaquearound and below the gingival margin. Moreover,it contains higher total protein compared to salivawhich is capable of providing nutrient to severalcommensal microorganisms in the oral cavity (7).Among the host defence components in GCF are IgGand neutrophils which are directed specificallyagainstimportant periodontal microorganisms and inhibit thecolonisationby the action ofopsonisationor activationof complement cascade (15).

Role ofbody temperatureThe optimum growth temperature for Candida

species including C.albicans has been shown to rangefrom 30°Cto 37°C (16). This range oftemperature isalso the optimum temperature of various pathogenicmicroorganisms in the oral cavity. Any alterationin the normal body temperature may influence thecompetitiveness among the normal microflora tosurvive which will then enhance the developmentof opportunistic microorganisms such as Candida.Nonetheless, many experimental assays wereconducted at 37°C and this is generally accepted asthe standard incubation temperature for Candidaspecies (7).

Intrinsic pathogenic determinants of CandidaspeciesThe virulent factors of each different Candida

species are not similar and can be a competitivefactor between each different species. Amongthe important virulent factors of Candida speciesare phenotypic switching, adhesion (both toextracellular matrix and dental biofilm), cell surfacehydrophobicity, and enzyme production.

Phenotypic switchingTwo mechanisms are postulated to be involved

in the ability of Candida to survive and adapt in asuppressed environment. The first is by undergoingmitotic recombination and the second is by carryingout phenotypic switching. A direct consequence ofmitotic recombination is the loss of heterozygositythroughout the entire genome. This deletion ofgenome, however, affects the viability of Candidaespecially in the multiple changing conditions(17). Phenotypic switching, on the other hand is aphenomenon that occurs as a result of changes in thegrowth environment. A severely suppressed growthcondition may lead to high frequency switching incandidal cells (Fig. 1). This adaptation is associatedwith the alteration of gene expression whicheventually may lead to alteration of adhesiveness,susceptibility and the resistance of candidal cellsto phagocytosis and polymorphonucleur (PMN)leukocyte. This mechanism of action does notinvolve deletion of any candidal genome, thus,the heterozygosity of the entire genomic are wellmaintained (7).Phenotypic switching in Candida albicans was

first defined in 1985 as the capacity to undergo


Fig. 1.A, B) Colony morphology ofthe unswitched and switched Candida krusei at lOx magnification using stereoscope.A) Unswitched, (B) switched generation. C, D) Unswitched (C) and switched (D) Candida krusei examined at lOOxmagnification using a light microscope; note Blastoconidia and Pseudohyphae.

spontaneous, reversible transitions between aset number of colony morphologies (18). Thisphenomenon is now recognized as an importanttechnique for the survival of Candida within anenvironment such as the oral cavity. This mechanismenablesCandida to adapt in a suppressed environmentand to develop as dominant in the host. Candida canundergo reversibly high frequency of phenotypicswitching which increases the survivability of thepathogen (17).Phenotypic switching is identified as one of the

important virulent factors in C. albicans (17) C.glabrata (19) and C. krusei (20). The significanceof the switching strategy is in a way similar to thehuman immunity function whereby it is aimed to

counter threats in the host's environment. Therefore,scientists have suggested that phenotypic switchingmechanism does enhance the survivability ofCandida by rapidly changing its phenotype as anadaptive response to the suppressed environment(21).Phenotypic switching may influence the normal

physiological growth ofCandida species such as C.albicans (17). Under the smooth white and wrinklephenotypes, C. albicans has been shown to exhibitfaster growing colonies compared to when it is inthe form of heavy myceliated with ring phenotype.In addition, phenotypic switching is also discoveredto be able to alter the adhesive properties ofCandida. Findings by our group (20) demonstrated


Fig. 2. SEMmicrographs ofCandida krusei observedfor the various growth generations at 2000x magnification. Notepseudohyphae (upper right panel) and attachment on extracellular matrix (lower right panel).

that the adherence ability of second generationswitched C. krusei was increase significantly inflow cell supplemented with unstimulated saliva.Furthermore, this virulence attribute may alsoinduce the formation of tube and pseudohyphae inCandida, which enhances the adherence capacity ofthe candida I strains (19).

Adhesion: key role of the extracellular polymericmatrixThe adherence ability ofCandida is an important

factor in the initiation oforal candidiasis. Adherencecan occur either on the hard tissue surfaces, such asteeth and palatal surface, or on soft surfaces, such asthe buccal and lingual surfaces (22). Characteristics

ofCandida that contribute to the adherence on thesesurfaces include the formation of pseudohyphae andextracellular matrix.A single filament hypha (plural, hyphae) is a long

branching filamentous structure of fungus whichcan be found easily in the developmental phase ofCandida (12). It is classified as the main mode ofvegetative fungal growth and consists ofone or morecells that are surrounded by tubular cell walls madeof chitin. Hyphae usually grow together to formcompact tufts which are known as mycelium. Hyphaeformation is usually referred to the germinationphase of fungi. However, it is also involved in thecolonisation of the target host. Pseudohyphae aredistinguished from true hyphae by their method of


growth, which lacks cytoplasmic connection betweenthe cells. The pseudohyphae of Candida are usuallyfound to possess incomplete budding blastoconidiawhereby cells remain attached to the mother cellsafter division. C. albicans and C. krusei have beenrecognised to develop pseudohyphae which adhereto the monolayer of human epithelial cells and hardsurfaces (20).In many cases, extracellular polymeric

substance (EPS) matrix is also produced by oralmicroorganisms once they are adhered to the oralsurfaces. EPS matrix is a network of non-livingmass which provides support to cells includingCandida (Fig. 2). The presence of EPS matrix,which has a slimy texture, provides a significant roleto support attachment and proliferation of the cells(23). Furthermore, the synthesis of EPS has alsobeen found to increase significantly when exposed toliquid flow (24). This anchorage property assists thecolonisation of Candida to hard tissue surfaces andthus, contributes to the formation ofbiofilm. When ina biofilm, the resistance of candidal species towardsvarious antifungal agents, including amphotericinB, voriconazole and ketoconazole, has been foundto increase up to lOOO-fold compared to planktonicstage (24, 25). Mitchell et al. (26) recently found thatthe presence of matrix B-1,3 glucan in EPS matrixsequesters antifungal drug which then increases theresistance to fluconazole.

Dental biofilmsBiofilm production is considered a potential

virulence factor of some Candida species (27).Dental biofilm is defined as a thin layer comprisingofvarious communities ofmicroorganisms includingbacteria, fungi and yeast that are attached to oralsurfaces and on the surface of prosthesis, includingdental acrylic surfaces and human epithelial cells.Microorganisms in the biofilm are enclosed ina matrix of extracellular polymeric substance(EPS). This biofilm provides protection to themicroorganisms and facilitates the interaction amongeach other with the contribution of enzymes such ascatalase and superoxidase dismutase (11, 28). Thedevelopment of biofilm is dependent on the dietary,salivary and oral environmental factors that interactwith the microorganisms within the community ofthe biofilm.

The formation of biofilm has been shown toreduce the susceptibility of microorganism toantimicrobial agents, which may then lead to theincrease in pathogenicity (11). This phenomenonis suggested to occur due to the restriction of theantimicrobial agents to penetrate the matrix of thebiofilm which then reduces the susceptibility ofthe target microorganism (29). Furthermore, thepresence of transcription factor Efg 1 in C. albicansbiofilm has also been reported to induce the tolerancetoward miconazole, caspofungin and amphotericinB (30).The development of dental biofilm involves

several stages which are the acquired pellicleformation on the oral surface; adhesion, reversibleand irreversible interactions between the pellicle andthe colonising microbes; co-aggregation betweenmicroorganisms; and detachment of microbes fromthe oral surfaces. These sequences of events mayeventually form a structural and functional organisedmicrobial community that, if allowed to accumulate,may enhance the potential of periodontal diseaseand dental caries (28). Specifically, co-aggregationor co-adhesion has been suggested to involveCandida in the late stage of oral biofilm formation.This is a process of microbial adhesion involvingthe late colonisers on to the early colonisers ofdental biofilm. It is a phenomenon of cell-to-cellrecognition of genetically distinct partner cell types(31). The co-aggregation can be facilitated eitherthrough intragenerics such as the interaction betweenS. sanguis and Actinomyces sp. or intergenericssuch as the interaction between Streptococcus sp.or Actinomyces sp. and Prevotella sp. C. krusei hasbeen found to be involved in co-aggreagation with S.mutans, S. sanguis and S. salivarius in the presenceof sucrose. C. albicans has also been reported tohave high coaggregation with S. sanguinis, S. oralisand S.gordonii (32). Protein such as lectin is usuallyinvolved in co-aggregation. This carbohydrate-binding protein attaches to the carbohydrate-bindingprotein receptors ofother cells which then contributeto the increased thickness of the dental biofilm.Once a climax community is achieved in the

biofilm, detachment of some microbes may occurin the final stage of the oral biofilm developmentThe microorganism is released from the matrix ofthe biofilm to the fluid surrounding the biofilm, a


process which has been reported to be facilitated byseveral enzymes such as proteases, fluid shear stress,multivalent cross-linking cations and microbialgrowth status (33). This process of detachmentwill, however, help the microorganism coloniseother surfaces in the oral cavity. An example of amicroorganism involved in the detachment processfrom the oral biofilm is Prevotella loescheii whichproduces proteases that hydrolyse adhesion-associated fimbriae which is important in its co-aggregation with S. oralis (31). Furthermore, thedetachment stage can also be initiated due to thepresence of certain quorum sensing molecules suchas farnesol, which has been found to be relatedto biofilm-self-Iimitation. The level of farnesolincreases proportionally to the number of Candidacells until threshold where the molecule starts tosuppress the yeast-to-mycelium conversion of newlybudding cells. As a result, the adherence is reducedwithin the architecture ofbiofilm, and releasing yeastforms Candida during the dissemination stage (34).

Cell surface hydrophobicityThe virulence factor of C. krusei can also be

observed from the cell surface hydrophobicitycharacteristic. This factor is classified as one ofthe most important adherence mechanisms inthe colonisation of the host surface, as well as indenture-related candidiasis (22). In fact, one of thekey properties contributing to the initial adherence tothe solid surfaces ofacrylic dentures are hydrophobicinteractions, and this feature has salient clinicalimplications for prevention and therapy of denture-related candidiasis. Various experimental approacheshave been used to examine the mechanisms ofhydrophobic interactions between Candida speciesand solid surfaces. The hydrophobic nature of thedenture surface has been cited as a factor in thedevelopment of new bactericidal materials (35).C. krusei is more hydrophobic compared to other

medically important Candida species (22). C. kruseiwas reported to possess the same hydrophobicitylevel as C. glabrata and C. Tropicalis, but is morehydrophobic compared to C. albicans and C.parapsilosis. Super-hydrophilic surfaces have beenreported to accept few bacterial or fungal cells (35)and could be a potent method for the reduction ofthe adherence of relatively hydrophobic fungal cells,

particularly the hyphal form of C. albicans whichcauses denture stomatitis and related infections.

Enzymatic activityHydrolytic enzymes of Candida have been

reported to contribute to its pathogenicity in causingoral diseases such as oral candidiasis. The enzymesinclude aspartyl proteinase, phospholipases, lipases,phosphomonoesterase and hexosaminidase (1).Among these enzymes, aspartyl proteinase hasattracted most interest and is widely considered tobe central in the development of candidal infection.Aspartyl proteinase is a hydrolytic enzyme whichis secreted by the transcription and translation ofsphingolipid activator protein (SAP) gene. Thisenzyme has the ability to invade host and alsocontributes as a defence system of yeast. Examplesof candidal species possessing this enzyme are C.albicans and C. krusei (22).

Another important hydrolytic enzyme isphospholipase which is identified as an enzyme thatinvades the host tissue. This enzyme activity hasbeen observed in many fungal pathogens includingCandida. There are 4 types of phospholipases,namely A, B, C and D. Phospholipase A and Ccan be found in C. albicans; however, there is noevidence that shows the presence ofphospholipase Band D in candidal species (22). Phospholipase A canattack cell membranes and can be easily found on thecell surface especially at the sites of bud formation.Hence, the enzyme activity can be enhanced whenthe hyphae are in direct contact with the host tissue(1).

CONCLUSIONS

Candidiasis is an ubiquitous infectious diseaseand its incidence has been increasing over the lastfew years, not only in immunocompromised patients,thus becoming a public health problem. Knowledgeof factors that affect the virulence of the Candidastrains is essential, and the oral cavity provides anideal environment to study not only the intrinsiccharacteristics of Candida, but also their interactionsin a complex environment such as the oral biofilm.An understanding ofthe molecular aspects underlyingthe biological behavior of Candida will be the key tothe development of effective preventive as well as


therapeuticmeasures for invasive and oral candidiasis.

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FEMS Yeast Research, 15, 2015, fov038

doi: 10.1093/femsyr/fov038Advance Access Publication Date: 8 June 2015Research Article

RESEARCH ARTICLE

Coaggregation of Candida albicans, Actinomycesnaeslundii and Streptococcus mutans is Candida albicansstrain dependentMohd Hafiz Arzmi1,2, Stuart Dashper1, Deanne Catmull1, Nicola Cirillo1,Eric C. Reynolds1 and Michael McCullough1,∗

1Oral Health CRC, Melbourne Dental School, The University of Melbourne, VIC 3053, Australia and 2Kulliyyahof Dentistry, International Islamic University Malaysia, 25200 Kuantan, Pahang, Malaysia∗Corresponding author: Melbourne Dental School, The University of Melbourne, Level 5, 720, Swanston Street, Carlton, VIC 3053, Australia.Tel: +613-9341-1490; Email: [email protected] sentence summary: Coaggregation between Candida albicans, Actinomyces naeslundii and Streptococcus mutans.Editor: Richard Calderone

ABSTRACTMicrobial interactions are necessarily associated with the development of polymicrobial oral biofilms. The objective of thisstudy was to determine the coaggregation of eight strains of Candida albicans with Actinomyces naeslundii and Streptococcusmutans. In autoaggregation assays, C. albicans strains were grown in RPMI-1640 and artificial saliva medium (ASM) whereasbacteria were grown in heart infusion broth. C. albicans, A. naeslundii and S. mutans were suspended to give 106, 107 and108 cells mL−1 respectively, in coaggregation buffer followed by a 1 h incubation. The absorbance difference at 620 nm(!Abs) between 0 h and 1 h was recorded. To study coaggregation, the same protocol was used, except combinations ofmicroorganisms were incubated together. The mean !Abs% of autoaggregation of the majority of RPMI-1640-grownC. albicans was higher than in ASM grown. Coaggregation of C. albicans with A. naeslundii and/or S. mutans was variableamong C. albicans strains. Scanning electron microscopy images showed that A. naeslundii and S. mutans coaggregated withC. albicans in dual- and triculture. In conclusion, the coaggregation of C. albicans, A. naeslundii and S. mutans is C. albicansstrain dependent.

Keywords: aggregation; yeast form; hyphal form

INTRODUCTIONAutoaggregation is defined as the adherence ability of mi-croorganisms belonging to the same species (Boris, Suarezand Barbes 1997), while coaggregation is the ability of genet-ically distinct microorganisms to adhere to each other (Led-der et al. 2008). Both autoaggregation and coaggregation havebeen classified as important mechanisms in the developmentof oral biofilms and postulated to provide protective mecha-nisms to the microbial inhabitants against shear forces that oc-

cur within the oral cavity. Aggregation contributes to the inte-gration of new microbial species into biofilms, facilitating theexchange of genes and metabolic products that in turn sup-ports survival of microorganisms against variable environmen-tal conditions (Gibbon and Nygaard 1970; Bos, Van-der-Mei andBusscher 1996; Kolenbrander 2000; Kolenbrander et al. 2002;Rickard et al. 2003; Al-Ahmad et al. 2007; Ledder et al. 2008).

Furthermore, coaggregation has been shown to improve thecolonization of oral epithelial cells by C. albicans, as prein-cubation of buccal epithelial cells with fimbriated strains of

Received: 9 December 2014; Accepted: 3 June 2015C⃝ FEMS 2015. All rights reserved. For permissions, please e-mail: [email protected]

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2 FEMS Yeast Research, 2015, Vol. 15, No. 5

Escherichia coli or Klebsiella pneumoniae increases the adherenceand subsequent attachment of C. albicans (Bagg and Silverwood1986). Preadherence of Streptococcus sanguinis and S. gordonii tothe hard surfaces of the oral cavity provides adhesion sites forC. albicans, which supports the importance of interkingdom in-teractions in the oral cavity (Jenkinson, Lala, Shepherd 1990;Bamford et al. 2009; Shirtliff, Peters and Jabra-Rizk 2009).

The oral microbiota comprises a wide variety of microorgan-isms such as yeasts (C. albicans) and bacteria (Actinomyces spp.and streptococci). Candida spp. that belong to kingdom fungi, es-pecially C. albicans, have been found to colonize approximately40–50% of healthy oral cavities (Manfredi et al. 2013). The num-ber increases in immunocompromised patients with diseasessuch as AIDS and diabetes (Grimaudo, Nesbitt and Clark 1996;Thein et al. 2009). The human oral microbiome is also comprisedof over 600 prevalent taxa at species level although only half ofthese have been cultured in the laboratory (Dewhirst et al. 2010).Among the important oral bacteria, A. naeslundii is an early oralcolonizer that can constitute up to 27% of supragingival dentalplaque (Nyvad and Kilian 1987; Li et al. 2004). The ability of thisspecies to coaggregate with other oral microorganisms has beenwell recognized (Grimaudo, Nesbitt and Clark 1996; Li et al. 2001).Streptococcus mutans, an acidogenic and aciduric gram-positiveoral bacterium, is widely regarded as a causative agent of dentalcaries (Peters et al. 2012).

The majority of in vitro studies of oral microbial coaggre-gation have assessed dual-species oral bacteria interactions(Cisar, Kolenbrander and McIntire 1979; Handley et al. 1985; Eke,Rotimi and Laughon 1989; Umemoto et al. 1999; Foster andKolenbrander 2004; Shen, Samaranayake and Yip 2005; Rosenand Sela 2006; Ledder et al. 2008), and information of interking-dom interactions is limited. Further, as yet, no study utilizingartificial saliva medium (ASM) for the growth of C. albicans hasbeen undertaken to assess interkingdom coaggregation. This isclinically relevant as C. albicans grows as yeast in ASM and ashyphae in RPMI-1640, and this dimorphism has a role in the vir-ulence of the species (Arzmi et al. 2012, 2014). The yeast formof C. albicans can adhere to the host cell surfaces by the ex-pression of adhesins, which trigger yeast-to-hyphae transition,followed by the expression of invasins by the hyphal form thatmediate the uptake of the fungus by the host cell through endo-cytosis (Molero et al. 1998; Gow et al. 2011; Sudbery 2011; Mayer,Wilson and Hube 2013). In addition, research has also found thatS. salivarius strain K12 preferred to coaggregate to the hyphalregion of C. albicans than the yeast after 3 h incubation in RPMI-1640 at planktonic phase (Ishijima et al. 2012). A similar inter-action was also observed between S. gordonii and C. albicans inwhich more bacteria coaggregated at the hyphal region of theyeast (Bamford et al. 2009).

The aim of the present study was to determine the co-aggregation of C. albicans, A. naeslundii and S. mutans withthe hyphotheses that autoaggregation and coaggregation areC. albicans strain dependent.

MATERIALS AND METHODSGrowth of microorganisms

C. albicansAmerican Type Cell Culture (ATCC) 32354 (ALT1), ATCCMYA-2876 (ALT2), ATCC 90234 (ALT3), ATCC 18804 (ALT4), geno-type A isolated from AIDS patient (ALC1), genotype B isolatedfrom AIDS patient (ALC2), oral cancer isolate 1 (ALC3) and oralcancer isolate 2 (ALC4) were used in this study. C. albicans strains

were subcultured on Sabauraud’s dextrose agar (Difco, USA) andincubated at 37◦C aerobically for 24 h.

To grow bacteria, stock cultures of A. naeslundii (NCTC 10301)and S. mutans (Ingbritt), provided by Oral Health Cooperative Re-search Centre, Melbourne Dental School, The University of Mel-bourne, were revived by subculturing onto blood agar (Difco,USA) and Todd-Hewitt yeast extract agar (Difco, USA), respec-tively. The agar plates were incubated at 37◦C for 48 h.

Aggregation assay

A semiquantitative spectrophotometric assay based on that out-lined by Ledder et al. (2008) and Nagaoka et al. (2008) was usedto analyse the aggregation of the microorganisms. Initially, 24-h cultures of C. albicans grown aerobically in RPMI-1640 or 25%ASM (0.625 g L−1 type II porcine gastric mucin, 0.5 g L−1 bacterio-logical peptone, 0.5 g L−1 tryptone, 0.25 g L−1 yeast extract, 0.088g L−1 NaCl, 0.05 g L−1 KCl, 0.05 g L−1 CaCl2 and 0.25 mg mL−1

haemin, pH 7.0 supplemented with 2.5 mM DTT and 0.5 g L−1

sucrose) to stationary phase were harvested by centrifuging at12 000 g for 5 min and washed twice using coaggregation buffer(0.1 mM CaCl2, 0.1 mM MgCl2, 150 mM NaCl, 3.1 mM NaN3 dis-solved in 1 mM Tris buffer and adjusted to pH 7.0). The super-natant was discarded and the pellet resuspended in coaggrega-tion buffer. A similar protocol was repeated for S. mutans andA. naeslundii except these microorganisms were grown in heartinfusion broth (HIB) to stationary phase.

To determine autoaggregation, C. albicans, A. naeslundii andS. mutans were standardized in coaggregation buffer to give a fi-nal cell density of 106, 107 and 108 cells mL−1, respectively inseparate sterile 2 mL Eppendorf tubes that were equivalent toan optical density of 0.5 at 620 nm wavelength (OD620nm). Eachsuspension was mixed thoroughly using a vortex mixer for 30 sand the OD620nm at time (t) = 0 h was measured. The inoculumwas incubated at room temperature for 1 h to allow aggregationand the OD620nm was recorded. Sterile coaggregation buffer wasused as the blank. Percentage aggregation was calculated usingthe following equation:

%Auto-aggregation = ([OD620nm(t = 0h)

−OD620nm(t = 1h)]/OD620nm(t = 0h)) × 100

Percentage autoaggregation was calculated for classificationof autoaggregation; (1) high (more than 40%), (2) intermediate(30–40%) and (3) low autoaggregation (less than 30%).

A similar protocol was repeated for the study of coaggrega-tion by inoculating C. albicans, A. naeslundii or/and S. mutans (in-terkingdom), and A. naeslundii and S. mutans (intrakingdom) intoa sterile 2 mL Eppendorf tube with the same cell density as inthe autoaggregation. The suspension was mixed thoroughly us-ing a vortex mixer and the OD620nm at t = 0 h was recorded. Thesuspension was incubated at room temperature for 1 h followedby the measurement of optical density at OD620 nm. The OD620nm

at time (t) = 0 h of dual culture and triculture were 1.0 and 1.5,respectively.

Coaggregation was assessed by measuring percentage coag-gregation using the following equation:

%Co-aggregation = ([OD620nm(t = 0h)

−OD620nm(t = 1h)] /OD620nm(t = 0h)) × 100

Arzmi et al. 3

Figure 1. Gram-stained of C. albicans cultures observed under light microscopy at 1000× magnification. Left: C. albicans (ALT4) grown in RPMI-1640 after 24 h incubationat 37◦C; >75% of C. albicans cells were present in hyphal form in this medium. Right: C. albicans (ALT4) grown in ASM after 24 h incubation at 37◦C; 100% of C. albicansdisplaying yeast morphology in this medium.

Scanning electron microscopy (SEM) imaging

The 0 h and 1 h suspensions (100 µL sample) of a selected rep-resentative C. albicans strain, ALT4, A. naeslundii (NCTC 10301)and S. mutans (Ingbritt), prepared as above, were transferred ontocover slips and fixed with 1% osmium tetra-oxide (OsO4) vapour.The specimens were dehydrated thoroughly in a freeze-dryingsystem, sputter coated with palladium gold to a thickness of ap-proximately 20 nm and observed using a scanning electron mi-croscope (XL 30 Series, Philips, Japan).

Statistical analysis

All data were statistically analysed using SPSS software version22.0 using independent t-test and considered statistically signif-icant when P < 0.05.

RESULTSMorphology of C. albicans in RPMI-1640 and ASM

C. albicans was shown to be predominantly in the hyphalform when grown in RPMI-1640 medium after 24 h incubationwhereas the yeast form was the most observed in ASM after thesame period of incubation (Fig. 1).

Autoaggregation

Variation in autoaggregation of RPMI-1640 grown C. albicansstrains (hyphal growth) was observed with a group of fourstrains (ALT3, ALT4, ALC1 and ALC3) exhibiting high autoag-gregation (over 40%), two strains (ALT1 and ALC4) exhibitingintermediate autoaggregation (30–40%) and two strains (ALT2and ALC2) exhibiting low autoaggregation (Table 1; Fig. 2A).The autoaggregation values of A. naeslundii and S. mutanswere also classified as low with 11.4 and 7.4%, respectively(Table 1).

Four strains of ASM-grown C. albicans (ALT2, ALT3, ALC1 andALC4) (yeast growth) exhibiting intermediate autoaggregationwhile the remainder strains (ALT1, ALT4, ALC2 and ALC3) wereclassified as exhibiting low autoaggregation (Table 1; Fig. 2B).

There were four strains of C. albicans that exhibited signifi-cantly more autoaggregation when grown in RPMI-1640 (hyphalgrowth) (ALT1, ALT4, ALC1 and ALC3) compared to ASM (yeastgrowth) (P<0.05). Two strains (ALT2 and ALC2) showed signifi-cantly more autoaggregation when grown in ASM than RPMI-

1640 (P<0.05) and two strains (ALT3 and ALC4) exhibited no dif-ference in autoaggregation regardless of the media type (Fig. 2).

Interkingdom coaggregation

All strains of RPMI-grown C. albicans (hyphal growth) were foundto coaggregate with A. naeslundii ranging from 9.9 ± 0.5% (ALT3)to 26.2 ± 0.4% (ALC3). Coaggregation of RPMI-grown C. albicanswith A. naeslundii and S. mutans was also observed for all strainsof the yeast ranging from 2.2 ± 0.3% (ALT3) to 17.0 ± 0.6% (ALC1).Our study showed that ASM-grown C. albicans strains (yeastform) coaggregated with A. naeslundii ranging from 9.6 ± 0.7%(ALT2) to 23.0 ± 0.1% (ALC3). ASM-grown C. albicans strains wereobserved to coaggregate S. mutans ranging from 9.9± 0.2% (ALT3)to 28.1 ± 0.1% (ALT4) (Table 1). Coaggregation of ASM-grown C.albicans with A. naeslundii and S. mutans were observed in allstrains of the yeast ranging from 12.9 ± 0.4% (ALT2) to 25.8 ±0.5% (ALT1) (Table 1).

SEM analyses

SEM analysis of RPMI-grown C. albicans ALT4 strain exhibitedautoaggregation in coaggregation buffer after 1 h incubation(Fig. 3A). Coaggregation was observed between C. albicans and A.naeslundii (Fig. 3B). In addition, an SEM image also revealed thatS. mutans coaggregated with C. albicans mostly at the hyphal re-gion of the yeast (Fig. 3C). The coaggregation of RPMI-grownALT4C. albicans with A. naeslundii and S. mutans showed that A. naes-lundii and S. mutanswere partially aggregating with C. albicans atthe hyphal region.A. naeslundiiwas also observed to coaggregatewith S. mutans (Fig. 3D).

SEM analysis showed that ASM-grown C. albicans ALT4 strain(yeast growth) had autoaggregation (Fig. 3E) andA. naeslundiiwasfound to coaggregate on the yeast surface after 1 h incubation(Fig. 3F). Coincubation of ALT4 C. albicanswith S. mutans revealedthat there was interkingdom coaggregation between the twomi-croorganisms with clumps of bacteria attached to the yeast sur-face of ALT4 C. albicans (Fig. 3G). In addition, an SEM image ofthe interaction between ASM-grown ALT4 C. albicans with bothbacterial species showed that A. naeslundii and S. mutans coag-gregated on the surface of the yeast. Finally, the image also re-vealed that S. mutans cells were coaggregating with A. naeslundiiafter 1 h incubation (Fig. 3H).

Taken together, the data demonstrate that the autoaggrega-tion and interkingdom coaggregation of C. albicans, A. naeslundiiand S. mutans are C. albicans strain dependent.


Table 1.Mono- and dual-culture aggregation scores of pairs of eight strains of RPMI-grown C. albicans (hyphal form), A. naeslundii and S. mutans.

Percent co-aggregation as measured by OD620 nm change over 1 h (see materials and methods section). Data are means from three separate experiments (SD are given in parenthesis). *Auto-aggregation scores representative of interactionbetween cells from the same culture. A. naeslundii and S. mutans were grown in BHI respectively

DISCUSSIONCoaggregation is a mechanism that induces the development ofa complex architecture of oral biofilms, which assists the attach-ment of secondary colonizers such as S. mutans (Kolenbrander2000; Min and Rickard 2009).

We have shown that interkingdom coaggregation was straindependent. The coaggregation of the majority of RPMI-grown(hyphal growth) C. albicans strains, when grown with S. mutansandA. naeslundii either alone or in combination, resulted in vari-able coaggregation. The observed variability of coaggregation inC. albicans may be attributable to the different abundances ofspecific molecules that are important in adhesion and quorumsensing (e.g. farnesol) from different strains, which have beensuggested to have a role in interkingdom interactions of C. al-bicans and bacteria (Morales and Hogan 2010). Furthermore, thevariability of coaggregation observed in ASM-grown C. albicans(yeast growth) supports our hypothesis that the coaggregationof C. albicans to A. naeslundii and S. mutans is highly dependenton the individual yeast strain.

We have observed variability of coaggregation when ASM-grown C. albicans strains were coincubated with S. mutans andA. naeslundii. This variability suggests that S. mutans might haveinduced the formation of binding sites on the yeast surface thatallow the coaggregation of A. naeslundii to ASM-grown C. albicanswhen cocultured. These results support our hypothesis that co-aggregation is highly dependent on the C. albicans strain. It can-not be related to the production of glucan by S. mutans glucosyl-transferases as no sucrose was present; however, it may be that

specific proteins are induced on the surface of C. albicans due tothe interaction with S. mutans that promotes further interactionwith A. naeslundii (Holmes, Gopal and Jenkinson 1995; Koo et al.2010; Falsetta et al. 2014). Further research is necessary to assessthis hypothetical possibility.

It can be postulated that the observed variability in coaggre-gation may be related to that specific strain’s ability to produceboth non-specific (adhesins) and specific (lectin-saccharide) cellsurface receptors (Kolenbrander and Williams 1981; Mcintire,Crosby and Vatter 1982; Rickard et al. 2003; Rosen and Sela 2006;Ledder et al. 2008). Previous studies have shown that the specificcoaggregation between C. albicans and A. naeslundii is due to thepresence of mannose-containing adhesin protein on the yeastcell surface (Grimaudo, Nesbitt and Clark 1996). This same studyalso showed variation in the coaggregation of A. naeslundii withfour different yeast strains which supports the present study.Furthermore, other research has shown significant strain vari-ation of the cell wall biogenesis in C. albicans, that may have arole in the observed variation in aggregation ability (Ragni et al.2011). Further analysis of the cell wall structure of a range of C.albicans strains is necessary to fully elucidate the mechanism ofthis observed variability.

It has previously been suggested that, due to the limitationof nutrients present in RPMI-1640, growth in this media inducesyeast–hyphae transition leading to predominant hyphal growth(Urban et al. 2006). Our light microscope images confirmed thiswith greater than 75% of C. albicans cells growing in hyphal formin RPMI-1640. No previous study has assessed the formof growthat SEM level when C. albicans is grown in ASM. The present study

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Figure 2. Percentage autoaggregation in RPMI-1640 (A) and ASM (B) grown C. albicans after 1 h incubation in coaggregation buffer. Data were analysed using independentt-test and considered as significantly different when P < 0.05. Asterisk indicates significantly more autoaggregation between the two growth media.

Figure 3. SEM of C. albicans autoaggregation (A and E), interkingdom interaction with A. naeslundii (B and F), S. mutans (C and G) and both bacteria (D and H). C. albicanswas grown in RPMI-1640 (A–D) and ASM (E–H). Magnification is as shown on each image (6500× and 10 000×).

is the first to observe C. albicans cellular morphology in ASM us-ing SEM imaging and we have shown that, similar to the lightmicroscope observations, in this media C. albicans does not growin hyphal form.

Future assessment of coaggregation of C. albicans, A. naes-lundii and S. mutans requires animal studies to assess oralbiological factors, such as salivary flow and immunologicalcomponents that exist in the oral cavity, which may in-fluence aggregation. These in vivo studies of coaggregationare likely to enhance our understanding of the mutual in-

teraction of microorganisms in the oral cavity, a processlikely to be critical in chronic infection and potentially oralcarcinogenesis.

CONCLUSIONIn conclusion, autoaggregation and interkingdom coaggrega-tion of C. albicans have been shown to be strain dependentand this is likely to be important in polymicrobial oral biofilmformation.


FUNDINGThis work was funded by Oral Health Cooperative Research Cen-tre (OHCRC) and the Melbourne Dental School.

Conflict of interest. None declared.

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Medical Mycology, 2016, 0, 1–9doi: 10.1093/mmy/myw042

Advance Access Publication Date: 0 2016Original Article

Original Article

Polymicrobial biofilm formation by Candidaalbicans, Actinomyces naeslundii, andStreptococcus mutans is Candida albicans strainand medium dependentMohd Hafiz Arzmi1,2, Ali D. Alnuaimi1, Stuart Dashper1, Nicola Cirillo1,Eric C. Reynolds1 and Michael McCullough1,∗

1Oral Health CRC, Melbourne Dental School, The University of Melbourne, Victoria, Australia and2Kulliyyah of Dentistry, International Islamic University Malaysia, Kuantan, Pahang, Malaysia∗To whom correspondence should be addressed. Michael McCullough, Level 4, Melbourne Dental School, MelbourneRoyal Dental, 720, Swanston Street, Carlton, 3053 VIC. Tel: +613 9341 1490; E-mail: [email protected]

Received 27 January 2016; Revised 24 April 2016; Accepted 26 April 2016

AbstractOral biofilms comprise of extracellular polysaccharides and polymicrobial microorgan-isms. The objective of this study was to determine the effect of polymicrobial interac-tions of Candida albicans, Actinomyces naeslundii, and Streptococcus mutans on biofilmformation with the hypotheses that biofilm biomass and metabolic activity are bothC. albicans strain and growth medium dependent. To study monospecific biofilms, C. al-bicans, A. naeslundii, and S. mutans were inoculated into artificial saliva medium (ASM)and RPMI-1640 in separate vials, whereas to study polymicrobial biofilm formation, theinoculum containing microorganisms was prepared in the same vial prior inoculation intoa 96-well plate followed by 72 hours incubation. Finally, biofilm biomass and metabolicactivity were measured using crystal violet and XTT assays, respectively. Our resultsshowed variability of monospecies and polymicrobial biofilm biomass between C. al-bicans strains and growth medium. Based on cut-offs, out of 32, seven RPMI-grownbiofilms had high biofilm biomass (HBB), whereas, in ASM-grown biofilms, 14 out of32 were HBB. Of the 32 biofilms grown in RPMI-1640, 21 were high metabolic activity(HMA), whereas in ASM, there was no biofilm had HMA. Significant differences wereobserved between ASM and RPMI-grown biofilms with respect to metabolic activity(P < .01). In conclusion, biofilm biomass and metabolic activity were both C. albicansstrain and growth medium dependent.

Key words: Polymicrobial biofilm, crystal violet assay, XTT assay.

IntroductionThe oral cavity is a habitat for various microorganisms in-cluding yeast and bacteria.1 This oral microbiome provides

a balanced oral environment however perturbation of thishomeostasis may lead to the development of dysbiosis andoral disease.2

C⃝ The Author 2016. Published by Oxford University Press on behalf of The International Society for Human and Animal Mycology.All rights reserved. For permissions, please e-mail: [email protected]

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Candida species, Actinomyces species and streptococciare common inhabitants of the human oral cavity.3,4,5 Can-dida spp. have been found to colonise approximately 50%of healthy human oral cavities.6 Candida albicans is themost frequently isolated Candida spp. from the oral cav-ity, especially in immunocompromised patients with dis-eases such as AIDS and diabetes.7,8 Many actinomycetesand streptococci are normal components of the human oralmicrobiota, with some species associated with dental cariesinitiation and development.4 Actinomyces naeslundii is cat-egorized as an early oral coloniser that can constitute up to27% of supragingival dental plaque.9,10 Streptococcus mu-tans is an acidogenic and aciduric Gram-positive oral bac-terium that is widely regarded as a pathogen that initiatesdental caries in association with other oral bacteria.4,11

Dimorphism is an important virulence factor of C. albi-cans. It is defined as the ability of Candida spp. to changemorphology between yeast and hyphal forms.12,13 C. albi-cans is predominantly in the yeast form during early coloni-sation of the oral cavity, however, subsequent invasion oforal epithelial cells is predominantly by the hyphal form.The yeast form of C. albicans can adhere to host cell sur-faces by the expression of adhesins, which trigger yeast-to-hyphae transition, followed by the expression of invasinsby the hyphal form that mediate the uptake of the fungusby the host cell through induced endocytosis.14–16

The majority of in vitro studies of biofilms havebeen with monospecies and dual-species oral microorgan-isms,17–25 and information from triculture polymicrobialbiofilms remains limited.26–28 As yet, no study utilising ar-tificial saliva medium (ASM) for the growth of C. albicanshas been undertaken to assess polymicrobial biofilms. Thisis clinically relevant as C. albicans grows as yeast in ASMand as hyphae in RPMI-1640.

Crystal violet (CV) and 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-5-[(penylamino)carbonyl]-2H-tetrazolium hy-droxide (XTT) assays are two methods for biofilm quantifi-cation. CV assay measures the microbial biofilm biomasswhere the dye interacts with negatively charged moleculespresent on the surface of the microorganisms and extra-cellular polysaccharide.10 The XTT assay is a colorimetric-based assay of cell metabolic activity using tetrazolium hy-droxide.29 Tetrazolium hydroxide is an active compoundthat is converted to formazan by the activity of dehydro-genases involved in the metabolic pathways of microbialcells.30 Succinate dehydrogenases of prokaryotic cells andmitochondrial dehydrogenases of eukaryotic cells are ex-amples of dehydrogenase activity that can be detected byXTT.29,31

The aims of the present study were to determine the effectof interactions of C. albicans, A. naeslundii and S. mutanson the formation of polymicrobial biofilms and to assess

this interaction when biofilms were grown in ASM for pre-dominantly yeast growth and in RPMI-1640 for predom-inantly hyphal growth. We hypothesized that this polymi-crobial biofilm formation is C. albicans strain- and growthmedium-dependent.

Materials and methods

Growth of microorganisms

C. albicans American Type Cell Culture (ATCC) 32354(ALT1), ATCC MYA-2876 (ALT2), ATCC 90234 (ALT3),ATCC 18804 (ALT4), a genotype A strain isolated fromoral infections in a human immunodeficiency virus (HIV)positive patient (ALC1), a genotype B strain isolated fromoral infections in an HIV positive patient (ALC2), a strainisolated from oral cancer patient number one (ALC3), and astrain isolated from oral cancer patient number two (ALC4)were used in the present study.32,33 C. albicans strains weresubcultured on Sabauraud’s dextrose agar (SDA) (Difco,USA) and incubated at 37◦C aerobically for 24 hours.

Bacteria were grown from stock cultures of A. naeslundii(NCTC 10301) and S. mutans (Ingbritt), provided by theOral Health Cooperative Research Cetre, Melbourne Den-tal School, The University of Melbourne, and were revivedby subculturing onto blood agar (40 g/l blood agar baseand 100 ml/l defibrinated horse blood) and Todd-Hewittyeast extract (THYE) agar (36.4 g/l Todd-Hewitt broth,8 g/l yeast extract and 15 g/l Bacto agar), respectively. Theagar plates were incubated at 37◦C for 24 hours.

Static biofilm formation

A quantitative assay based on that outlined by Ya-mada et al.34 and Alnuaimi et al.32 was used to analyzestatic biofilm formation by the microorganisms. To studymonospecies biofilm, streak diluted cultures of C. albicans,A. naeslundii and S. mutans were grown on SDA, bloodagar and THYE agar respectively, for 24 hours at 37◦C andseveral single colonies were resuspended in RPMI-164032

or 25% ASM (0.625 g/l type II porcine gastric mucin, 0.5g/l bacteriological peptone, 0.5 g/l tryptone, 0.25 g/l yeastextract, 0.088 g/l NaCl, 0.05 g/l KCl, 0.05 g/l CaCl2 and0.25 mg/ml haemin, pH 7.0 supplemented with 2.5 mMDTT and 0.5 g/l sucrose), and standardized to give a finalcell density of 106 cells/ml, 107 cells/ml and 108 cells/ml,respectively, in a separate sterile 2-ml Eppendorf tubes thatequivalent to an absorbance of 0.5 at 620 nm wavelength(OD620nm). The suspensions were mixed thoroughly usinga vortex mixer for 30 seconds. Subsequently, 200 µl ofeach suspension containing 2 × 105 cells (C. albicans), 2 ×106 cells (A. naeslundii) and 2 × 107 cells (S. mutans) of

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initial inoculum was pipetted into each well of sterile 96-well plate (Nunc, Denmark). Finally, the plate was incu-bated in an orbital shaker at 90 rpm for 72 hours at 37◦C(Alyos, Thermo Fisher Scientific, Australia) to mimic the dy-namic of oral environment.32 The medium was replenishedaseptically every 24 hours.

A similar protocol was used to study polymicrobialbiofilm formation by inoculating C. albicans, A. naeslundiior/and S. mutans into a sterile 2-ml Eppendorf tube with asimilar cell density as in the monospecies assay resulting in2 × 105 cells (C. albicans), 2 × 106 cells (A. naeslundii),and 2 × 107 cells (S. mutans) for each combination perwell. The suspension was mixed thoroughly using a vortexmixer and 200 µl of the suspension was pipetted into ster-ile 96-well plate. The plate was incubated aerobically for72 hours at 37◦C in an orbital shaker at 90 rpm and themedium was replenished aseptically every 24 hours.

Gram stain

Gram stain was performed on C. albicans ALC3 strain fol-lowing growth in RPMI-1640 and ASM for 72 hours at37◦C for the determination of morphology. Initially, 1 mLof suspension of RPMI-1640 or ASM-grown C. albicanscontaining 2 × 105 cells was pipetted into each well of12-well plate and incubated at 37◦C in an orbital shakerat 90 rpm. The medium was replenished aseptically every24 hours of incubation. Following incubation, the super-natant was discarded and each well was washed carefullywith phosphate buffered saline (PBS) (Sigma-Aldrich, USA)twice to remove non-adherent cells. Later, Gram stainingwas performed and the sample was observed under a lightmicroscope (CH Series, Olympus, Australia).35

Crystal violet assay

Crystal violet (CV) assay was performed according to theprotocol outlined by Alnuaimi et al.32 Initially, the biofilmin each well of 96-well plate was washed twice with ster-ile PBS to remove nonadherent cells. In sum, 200 µl ofmethanol was added to each well for fixation and incu-bated for 15 minutes at 25◦C. The supernatant was thendiscarded and the plate was air-dried for 45 minutes. And200 µl of 0.1% (w/v) CV solution was added into each welland incubated for a further 20 minutes at 25◦C. The platewas washed gently twice using running distilled water, and200 µl of 33% (v/v) acetic acid was added to de-stain thebiofilm. The plate was incubated for five minutes at roomtemperature. A 100 µl aliquot of this solution was trans-ferred to a new sterile 96-well plate and the absorbancewas measured at OD620 nm using a microtiter plate reader(Victor3, Perkin-Elmer, Australia).

XTT reduction assay

XTT reduction assay was performed according to the pro-tocol provided by the manufacturer (Sigma-Aldrich, USA).Briefly, the biofilm-coated wells were washed twice withsterile PBS to remove non-adherent cells. Subsequently,160 µl of sterile PBS and 40 µl of 4% XTT salt containing1% phenazine methosulphate (Sigma-Aldrich, USA) werepipetted into each well to give a final volume of 200 µl.The plate was incubated at 37◦C for three hours in the dark.Following incubation, 100 µl of the suspension was trans-ferred into a new sterile 96-well plate and the absorbance atOD450 nm and OD620 nm wavelengths were measured usinga microtiter plate reader. Measurement of absorbance atthe reference wavelength of OD620 nm was subtracted fromOD450 nm to remove background absorbance.

Statistical analysis

All biofilms containing C. albicans were divided into tercilesaccording to biofilm biomass and metabolic activity for CVand XTT assays, respectively. This method of dividing apopulation of C. albicans containing biofilms into high,moderate and low has been previously used to assess bothbiofilm biomass and biofilm metabolic activity.36 This divi-sion provided the cut-offs to classify strains as high, mod-erate and low biofilm biomass (HBB, MBB and LBB); andhigh, moderate and low metabolic activity (HMA, MMA,and LMA). Using SPSS software version 22.0, all data werestatistically analyzed by applying chi-square test to com-pare between the categories for each assay and two-tailed t-test to compare between ATCC and clinical strains biofilmbiomass. Comparison between a group of ATCC isolates(ALT1, ALT2, ALT3, and ALT4) and a group of clini-cal isolates (ALC1, ALC2, ALC3, ALC4) of C. albicanswas analyzed using two-tailed t-test. Multiple comparisonsbetween monospecies with polymicrobial biofilms such asbetween monospecies C. albicans ALT1 with dual-speciesC. albicans ALT1-A. naeslundii, C. albicans ALT1-S. mu-tans, and trispecies, were compared using ANOVA post hocTukey test.

Results

Morphology of C. albicans biofilms in RPMI-1640and ASM

C. albicans biofilm growth was predominantly in the hyphalform when grown in RPMI-1640, and in the yeast formwhen grown in ASM after 24 hours incubation as observedby Gram staining (Figure 1).

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Figure 1. Gram-stained biofilms of Candida albicans strain ALC3 observed under light microscope at 200x magnification after 72 hours incubation at37◦C in 24-well plate at 90 rpm. A: Artificial saliva medium (ASM)- grown Candida albicans biofilm; B: RPMI-grown Candida albicans biofilm.

Effect of microbial interaction and growthmedium on biofilm biomass

Biofilm biomass was categorized into terciles using thefollowing CV measurement cut-offs: LBB < 2.280, MBB2.280-2.535, HBB > 2.535. None of monocultured C. al-bicans was categorized as HBB, however, when co-culturedwith A. naeslundii three C. albicans strains (ALT1, ALT2,and ALT3) were categorized as HBB (Table 1). Only ALT1was categorized as HBB when co-cultured with S. mutansin RPMI-1640 whereas in tricultured biofilms, three strains

of C. albicans (ALT1, ALT2, and ALT3) were categorizedHBB (Table 1).

None of ASM monocultured C. albicans exhib-ited HBB, however, in the presence of A. naeslundii,seven strains of C. albicans were classified as HBB(Table 1). Interaction of C. albicans with S. mutans showedthat two strains (ALT1 and ALT3) were HBB, whilein tricultured biofilms, five C. albicans strains (ALT1,ALT2, ALT3, ALT4, and ALC2) were classified as HBB(Table 1).

Table 1. Static biofilm biomass scores of 8 strains of RPMI-grown (hyphal form) and artificial saliva medium (ASM)-grown

(yeast form) Candida albicans, Actinomyces naeslundii (An), and Streptococcus mutans (Sm).

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Table 2. Static biofilm metabolic activity scores of RPMI-grown (hyphal form) and artificial saliva medium (ASM)-grown (yeast

form) Candida albicans, Actinomyces naeslundii (An) and Streptococcus mutans (Sm).

Analyses of all 32 biofilms for biomass showed thatthere were seven biofilms classified as HBB (21.9%), 12MBB (37.5%), and 13 LBB (40.6%) when the biofilms weregrown in RPMI-1640 (hyphal growth). Biofilms grown inASM (yeast form) showed 14 biofilms categorized as HBB(43.8%), ten MBB (31.3%), and eight LBB (25.0%). Therewere more biofilms with HBB when grown in ASM (yeastform) than RPMI-1640 (hyphal form), however, this didnot reach statistical significance (P > .05).

Five RPMI-grown biofilms (hyphal form) had signifi-cantly increased biomass when C. albicans strains were co-cultured with A. naeslundii (ATCC: ALT1, ALT4; Clinical:ALC1, ALC2 and ALC4) compared with monocultured C.albicans biofilm (P < .05). Further, co-culture of C. al-bicans with S. mutans increased biomass of six biofilms(ATCC: ALT1, ALT4; Clinical: ALC1, ALC2, ALC3 andALC4) significantly (P < .05). Five biofilms (ATCC:ALT1, ALT4; Clinical: ALC1, ALC2, and ALC4) increasedbiomass significantly when C. albicans was co-culturedwith both A. naeslundii and S. mutans when comparedwith the monocultured biofilm of C. albicans (P < .05;Table 1).

Two ASM-grown biofilm (ATCC: ALT1 and ALT2;yeast form) had a significantly increased biomass when C.albicans was co-cultured with A. naeslundii compared withthe monocultured C. albicans biofilm (P < .05). One biofilm

(ATCC: ALT1) showed a significant increase (P < .05)and one (ATCC: ALT2) a significant decrease (P < .05)in biomass when C. albicans was co-cultured with S. mu-tans. There was one strain (ATCC: ALT1) that showeda significant increase in biomass when C. albicans wasco-cultured with both A. naeslundii and S. mutans com-pared with monocultured C. albicans biofilm (P < .05;Table 1).

Effect of microbial interaction and growthmedium on metabolic activity

Biofilm metabolic activity based on the XTT assay was di-vided into terciles and categorized based on the follow-ing cut-offs: LMA < 0.120, MMA 0.120-0.550, HMA >

0.550. RPMI-1640 monocultured growth resulted in sixstrains of C. albicans (ALT1, ALT2, ALT4, ALC1, ALC2,and ALC4) categorized with HMA (Table 2). Seven C. al-bicans strains (ALT1, ALT2, ALT3, ALT4, ALC1, ALC2,and ALC4) when co-cultured with A. naeslundii in RPMI-1640 had HMA. Only two strains of C. albicans (ALT1and ALT2) had HMA when co-cultured with S. mutans inRPMI-1640. Six C. albicans strains (ALT1, ALT2, ALT3,ALT4, ALC2, and ALC4) were categorized as having HMAwhen co-cultured in RPMI-1640 with both A. naeslundiiand S. mutans (Table 2).

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Table 3. Monospecies biofilm metabolic activity over biomass (XTT assay/crystal violet assay) scores of eight strains of RPMI-

grown (hyphal form) and artificial saliva medium (ASM) - grown (yeast form) Candida albicans (ALT1 to ALT4 for ATCC strains

and ALC1 to ALC4 for clinical strains), Actinomyces naeslundii (An) and Streptococcus mutans (Sm).

Media ALT1 ALT2 ALT3 ALT4 ALC1 ALC2 ALC3 ALC4 An Sm

RPMI-1640 0.326 0.319 0.166 0.476 0.366 0.382 0.807 0.388 0.681 0.022(0.007) (0.046) (0.044) (0.075) (0.027) (0.009) (0.221) (0.044) (0.035) (0.016)

ASM 0.008 0.005 0.007 0.051 0.011 0.010 0.051 0.022 0.087 0.003(0.000) (0.001) (0.002) (0.016) (0.002) (0.000) (0.032) (0.006) (0.019) (0.002)

Data are means from three separate experiments (SD are given in parentheses).

ASM monocultured growth resulted in all C. albicansstrains being categorized with LMA (Table 2), however, inthe presence of A. naeslundii, all strains had MMA. Inter-action of C. albicans with S. mutans showed that all C.albicans strains remained with LMA whereas, in the pres-ence of both A. naeslundii and S. mutans, there were threestrains having MMA (ALT1, ALT4, and ALC2) and fivestrains with LMA (ALT2, ALT3, ALC1, ALC3, and ALC4)(Table 2).

Analyses of all 32 biofilms showed that there were21 biofilms of RPMI-grown biofilms (hyphal growth) cat-egorized as having HMA (65.6%) and 11 with MMA(34.4%). In addition, there were 11 ASM-grown biofilms(yeast growth) categorized as having MMA (34.4%) and21 with LMA (65.6%). Thus, statistically significant highermetabolic activity was observed when biofilms were grownin RPMI-1640 (P < .01).

Only C. albicans strains ALT3 when co-cultured withA. naeslundii showed an increased activity when grownin RPMI-1640 when compared with monospecies C. al-bicans. Furthermore, there were four C. albicans strains(ALT4, ALC1, ALC2, and ALC4) that exhibited a decreasein metabolic activity when co-incubated with S. mutanscompared with the monocultured biofilm of C. albicans.There was only one biofilm (ALC1) that showed decreasedbioactivity when C. albicans was co-cultured with both A.naeslundii and S. mutans compared with monocultured C.albicans (Table 2).

Three RPMI-grown biofilms (ATCC: ALT3; Clinical:ALC2 and ALC4; hyphal form) exhibited significant in-creased activity when C. albicans was co-cultured with A.naeslundii in comparison with the monocultured C. albi-cans biofilm (P < .05). Four biofilms (ATCC: ALT4; Clini-cal: ALC1, ALC2, and ALC4) showed significant decreasedmetabolic activity when C. albicans was co-cultured with S.mutans. Whereas, one biofilm (Clinical: ALC1) displayeda significant decreased activity when C. albicans was co-cultured with both A. naeslundii and S. mutans when com-pared with monocultured C. albicans (P < .05; Table 2).

Finally, based on metabolic activity per unit biomass inmonospecies biofilms, ALT4 and ALC3 were found to be

the most active C. albicans strains when grown in ASM andALT2 was the least active when grown in the same medium.Whereas, in RPMI-1640, ALC3 was found to be the mostactive while ALT3 was the least (Table 3).

DiscussionTo our knowledge, this is the first study to evaluate theeffect of microbial interactions of yeast growth and hyphalgrowth of C. albicans, A. naeslundii, and S. mutans onthe formation of static biofilms in vitro. The results of thepresent study clearly demonstrate that both biofilm biomassand metabolic activity are C. albicans strain and growthmedium dependent.

The present study has shown a variation of biofilmbiomass and metabolic activity between strains of C. al-bicans. Overall, when grown as monospecies the majorityof clinical strains had a significantly lower biofilm biomassthan the ATCC reference strains. However, a significant in-crease of biomass was observed in all clinical strains that didnot occur in ATCC strains (ALT2 and ALT3) when grownin polymicrobial biofilms. Previous research also showedthat biofilms formed by clinical isolates of C. albicans ex-hibited lower biofilm biomass compared with the referencestrains C. albicans.32 Furthermore, the metabolic activityhas been shown to vary among C. albicans strains; how-ever, the morphology of C. albicans in this previous studywas unknown.32 Strain variability of C. albicans has beenshown in the oral cavity of different individuals.37,38 Pre-vious research has shown that C. albicans strains isolatedfrom HIV-infected patients produce higher levels of asparticproteinases (SAPs), compared with strains isolated from un-infected patients.39 SAP is a putative virulence factor thatis able to affect C. albicans biofilm formation in the oralcavity together with phenotypic switching, morphogenesisand quorum sensing.1,12 Thus, the results from the presentstudy may indicate a symbiotic interaction between clinicalC. albicans and oral microorganisms that may lead to the in-crease of colonisation in the oral cavity of diseased patients.

The metabolic activity of biofilms was shown tobe growth media dependent, with the majority of

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ASM-grown C. albicans biofilms having lower metabolicactivity than those grown in RPMI-1640, particularlymonospecies biofilms (Table 2, Table 3). It is postulatedthat RPMI-1640, which contains limited nutrients, inducesstress in C. albicans, thus promoting hyphal formation. Thisdoes not occur when the yeast is grown in ASM that is richin nutrients. Interestingly, previous studies have shown thatCandida spp. with low metabolic activity are more invasiveand associated with disease, while conversely those withhigh activity are non-invasive.31,40,41 Furthermore, lowmetabolic activity has been shown to reduce the antifungalsusceptibility of C. albicans within the biofilm, which couldbe due to minimal absorption of antifungal agents such asamphotericin B, thus affecting inactivation kinetics.42

The metabolic activity of all C. albicans strains that weregrown in ASM increased in the presence of A. naeslundii indual-cultured biofilms. However, a decrease of metabolicactivity was observed in trispecies biofilms when comparedto the dual-cultured biofilms of C. albicans and A. naes-lundii, suggesting that these microorganisms may be inter-acting metabolically. It is postulated that in the presenceof A. naeslundii, C. albicans may increase mitochondrialdehydrogenase activity that in turn, increased the activityof succinate dehydrogenases of A. naeslundii. In addition,S. mutans has been shown to reduce the metabolic activ-ity in trispecies biofilms compared with the dual-culturedC. albicans-A. naeslundii biofilms, suggesting that the an-tagonistic metabolic interaction between A. naeslundii andS. mutans, demonstrated in the present study (Table 2),may have affected overall metabolic activity of the con-sortia. C. albicans and A. naeslundii have been shown tosynthesize mitochondrial and succinate dehydrogenases, re-spectively, that were reported to be detectable by XTT.29,31

Even though S. mutans has been found to synthesize anNADH-dependent lactate dehydrogenase; the present studyrevealed that enzyme activity was not detected with XTTsuggesting that the assay is not suitable for the study of S.mutans metabolic activity.

In the present study, the biofilm biomass was shown tovary with microbial interactions (monocultured C. albicans,dual-cultured C. albicans and A. naeslundii, dual-culturedC. albicans and S. mutans, tricultured C. albicans, A. naes-lundii, and S. mutans). The majority of RPMI-1640 grownC. albicans (hyphal form) biofilm biomass was observed toincrease in the presence of bacteria compared with mono-cultured C. albicans. Previous research has shown that A.naeslundii and S. mutans bind to C. albicans through itsmannose-containing surface protein.7,23,24,43,44,45 This in-teraction has been reported to induce the formation of ex-tracellular polysaccharide, thus promoting the adherenceof the late colonisers to form a complex polymicrobialbiofilm potentially enhancing biofilm biomass.4,7,9,10 Pre-

vious studies have also demonstrated that oral biofilms arecomposed of various microorganisms1,2 indicating the im-portant role of polymicrobial interactions in plaque biofilmdevelopment, dysbiosis, and oral disease.

The present study found that the ATCC strains formexcellent monocultured biofilms in both ASM and RPMI-1640 such that addition of A. naeslundii or S. mutans re-sulted in no additional biomass in the majority of biofilms.However, the clinical strains that were poor biofilm for-mers in RPMI-1640 were observed to increase biofilmbiomass significantly when A. naeslundii or S. mutans wasco-inoculated (Table 1). This result indicates that the choiceof isolates in the study of the interaction between oral yeastand oral bacteria in biofilms is critical. The C. albicansATCC strains assessed in the present study would appearto have lost either the ability, or need, to interact with oralbacteria;46 thus, investigations using only ATCC strains ofC. albicans are likely to not reflect the true interactions thatare occurring in the oral cavity.

We have demonstrated that C. albicans predominantly inthe yeast form when grown as a biofilm in ASM, whereasRPMI-grown C. albicans biofilms were predominated bythe hyphal form (Fig. 1). These results support previouswork that showed the proportion of yeast and hyphal cellsof C. albicans present in the biofilm is dependent upon thenutrient source, where nitrogen-based medium allowed formore yeast growth and biofilms grown in RPMI-1640 withhigh salts, amino acids and D-glucose, showed more hyphalgrowth.47

ConclusionBiofilm biomass and metabolic activity have been shown tobe both C. albicans strain and growth medium dependent.This is likely to have significance in the development ofpolymicrobial oral biofilms in vivo.

AcknowledgmentsThis work was funded by the Oral Health Cooperative ResearchCentre (OHCRC) and Melbourne Dental School, The University ofMelbourne.

Declaration of interestThe authors report no conflicts of interest. The authors alone areresponsible for the content and the writing of the paper.

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