impact of quorum sensing on the virulence of vibrio crassostreae … · 2016-12-10 · faculty of...

FACULTY OF BIOSCIENCE ENGINEERING

GHENT UNIVERSITY

Academic Year 2015-2016

Impact of quorum sensing on the virulence of Vibrio

crassostreae and Vibrio tasmaniensis in vitro and in vivo

in blue mussel larvae

SHIKDER SAIFUL ISLAM

Promoter : Dr. ir. Tom Defoirdt

Supervisor : Mieke Eggermont

Master’s dissertation submitted in partial fulfillment of the requirements for

the degree of

MASTER OF SCIENCE IN AQUACULTURE

i

Copyright

The author and promoter give permission to put this thesis to disposal for consultation and to copy

parts of it for personal use. Any other use falls under the limitations of copyright, in particular the

obligation to explicitly mention the source when citing parts out of this thesis.

Ghent

August, 2016

Promoter

Dr. ir. Tom Defoirdt

Supervisor

Mieke Eggermont

Author

Shikder Saiful Islam

ii

Acknowledgements

To begin with, I wish to express my heartfelt gratitude to Almighty Allah, Whose never-ending

blessings enabled me to successfully finish this research work and prepare this master disserta-

tion.

I wish to express my deepest admiration and sincere appreciation to Dr. ir. Tom Defoirdt for his

enduring direction, passionate supervision, praiseworthy suggestions, positive condemnation and

immeasurable help during my thesis write up.

I would like to express my greatest appreciation to Mieke Eggermont, to whom I be indebted my

warm respect. My thoughtful pleasure and indeptness goes to her for support during the hard

times and honest concern helped me to proceed and make come to an end line.

I wish to thank Prof. Dr. ir. Peter Bossier for his continuous suggestions, worthy inspirations and

regular monitoring during the thesis work and notably last two years to complete my MSc Aqua-

culture programme from Ghent University.

I thank Dr. Frédérique Le Roux (Station Biologique de Roscoff, Roscoff, France) for generously

providing us with strains LGP32 and J2-9 and their quorum sensing mutants.

My thanks go to Dr. Nancy Nevejan and Tom Baelemans for helping to supply of my experi-

mental animals.

I wish to state my respect to the professors and staff members connected with the Master of Sci-

ence in Aquaculture.

I would like to highly acknowledge Dr. ir. Peter De Schryver, Brigitte Van Moffaert, Geert

Vandewiele, Christ Mahieu, Anita De Haese and other staffs of Laboratory of Aquaculture and

Artemia Reference Center for their unforgettable helps and sincere appreciation.

I am also grateful to my friends at Ghent University, who always appreciated me during my hard

time and depression.

I would also like to thank Flemish Interuniversity Council for their support to promote my stud-

ies and the Laboratory of Aquaculture and Artemia Reference Center, Ghent University for

providing a helpful atmosphere for my studies.

I gratefully acknowledge VLIR-UOS for the financial support of the full research and study ex-

penses.

Finally I would like to thank my beloved mother, to the departed soul of my father and sister, and

other family members for their boundless love, hysterically inspiration and mental support during

my stay here in Belgium and all the way through of my life.

Date: August 19, 2016 Shikder Saiful Islam

Ghent, Belgium

iii

Table of Contents

Copyright .................................................................................................................................................. i

Acknowledgements ................................................................................................................................. ii

List of Figures ........................................................................................................................................ vi

List of tables ......................................................................................................................................... viii

List of abbriviations and units ................................................................................................................ ix

Dedication .............................................................................................................................................. xi

Abstract ................................................................................................................................................. xii

1 Introduction ........................................................................................................................................ 1

2 Review of literature ............................................................................................................................ 3

2.1 Overview of aquaculture ..................................................................................................... 3

2.2 The impact of bacterial diseases on aquaculture ................................................................... 5

2.2.1 Environmental stress and disease outbreaks in aquaculture ............................................5

2.2.2 Disease caused by pathogenic vibrios...........................................................................6

2.2.3 Problems related to the use of antibiotics in aquaculture ................................................6

2.3 The blue mussel (Mytilus edulis) ......................................................................................... 7

2.3.1 Taxonomy ..................................................................................................................7

2.3.2 Anatomy ....................................................................................................................8

2.3.2.1 External anatomy ............................................................................................... 8

2.3.2.2 Internal anatomy ................................................................................................ 8

2.3.2.3 Digestive system ............................................................................................... 9

2.3.2.4 Cardiovascular and respiratory system ................................................................ 9

2.3.2.5 Immune system ............................................................................................... 10

2.3.3 Life cycle .................................................................................................................11

2.3.3.1 Reproduction ................................................................................................... 11

2.3.3.2 Larval development ......................................................................................... 11

2.3.4 Blue mussel cultivation .............................................................................................12

2.3.5 Diseases in bivalve and blue mussel cultivation ..........................................................13

2.3.5.1 Diversity of pathogens ..................................................................................... 13

2.3.5.2 Larval diseases ................................................................................................ 14

2.3.5.3 Diseases in juvenile and adult bivalves .............................................................. 15

2.4 Virulence factors of vibrios ............................................................................................... 16

2.4.1 Motility and chemotaxis ............................................................................................16

2.4.2 Production of extracellular polysaccharides and biofilm formation ...............................17

2.4.3 Production of lytic enzymes ......................................................................................18

2.4.3.1 Hemolysin ...................................................................................................... 18

2.4.3.2 Proteases ......................................................................................................... 18

2.4.3.3 Lipase ............................................................................................................. 19

iv

2.4.3.4 Phospholipase ................................................................................................. 19

2.4.3.5 Chitinase ......................................................................................................... 19

2.4.4 Iron acquisition and siderophores ...............................................................................20

2.5 Quorum sensing, bacterial cell-to-cell communication ........................................................ 20

2.5.1 General overview .....................................................................................................20

2.5.2 Multichannel quorum sensing systems in vibrios ........................................................21

2.5.3 Impact of quorum sensing on the virulence of vibrios..................................................24

2.6 Antivirulence therapy: inhibition of virulence factor production as a new strategy to

control bacterial disease .................................................................................................... 24

2.6.1 Quorum sensing disruption ........................................................................................24

2.6.1.1 Inhibition of signal molecules biosynthesis ........................................................ 24

2.6.1.2 Application of quorum sensing antagonists ........................................................ 25

2.6.1.3 Enzymatic inactivation and biodegradation of quorum sensing signal molecules .. 25

2.6.2 Antivirulence compounds targeting other regulatory mechanisms ................................25

2.6.3 Antivirulence compounds targeting one specific virulence factor .................................26

3 Materials and methods ...................................................................................................................... 27

3.1 Preparation of culture media ............................................................................................. 27

3.1.1 Luria-Bertani broth (LB35).........................................................................................27

3.1.2 Luria-Bertani Agar (LB35 agar) ..................................................................................27

3.1.3 Thiosulphate-citrate-bile salts-sucrose (TCBS) ...........................................................27

3.2 Preparation of antibiotic stock solutions............................................................................. 27

3.3 Bacterial strains ............................................................................................................... 27

3.4 Storage and culture of bacterial strains .............................................................................. 28

3.5 Bacterial strain verification ............................................................................................... 28

3.5.1 DNA Extraction .......................................................................................................28

3.5.2 ERIC-PCR ...............................................................................................................28

3.5.3 Gel Electrophoresis ..................................................................................................29

3.5.4 Assessment of Virulence factors ................................................................................29

3.5.5 Swimming motility ...................................................................................................29

3.5.6 Lytic enzymes ..........................................................................................................30

3.5.6.1 Hemolysin ...................................................................................................... 30

3.5.6.2 Caseinase ........................................................................................................ 30

3.5.6.3 Gelatinase ....................................................................................................... 30

3.5.6.4 Lipase ............................................................................................................. 30

3.5.6.5 Phospholipase ................................................................................................. 30

3.5.7 Biofilm formation .....................................................................................................31

3.5.8 Exopolysaccharide production ...................................................................................31

3.6 Larval challenge test ........................................................................................................ 31

3.6.1 Blue mussel D-larvae ................................................................................................31

v

3.6.2 Experimental design .................................................................................................32

3.6.2.1 Challenge test 1 ............................................................................................... 33

3.6.2.2 Challenge test 2 ............................................................................................... 33

3.6.2.3 Challenge test 3 ............................................................................................... 34

3.7 Statistical analysis ............................................................................................................ 34

4 Results .............................................................................................................................................. 35

4.1 Molecular characterization of wild types and selected quorum sensing mutants .................... 35

4.2 Virulence factor production by wild types and quorum sensing mutants ............................... 35

4.2.1 Swimming motility ...................................................................................................36

4.2.2 Production of lytic enzymes ......................................................................................36

4.2.2.1 Hemolysin ...................................................................................................... 36

4.2.2.2 Proteases ......................................................................................................... 37

4.2.2.3 Phospholipase ................................................................................................. 38

4.2.2.4 Lipase ............................................................................................................. 38

4.2.3 Biofilm formation .....................................................................................................38

4.2.4 Exopolysaccharide production ...................................................................................40

4.3 Blue mussel larval challenge tests ..................................................................................... 41

4.3.1 Challenge test 1: mussel larvae challenged with pathogenic isolates Photobacterium

sp. ME5 and Vibrio sp. ME6 .....................................................................................41

4.3.2 Challenge test 2: mussel larvae challenged with V. crassostreae J2-9 and V.

tasmaniensis LGP32 and their QS mutants .................................................................43

4.3.3 Challenge test 3: impact of the quorum sensing inhibitor cinnamaldehyde on the

survival of mussel larvae challenged with V. crassostreae J2-9, V. tasmaniensis

LGP32, Photobacterium sp. ME5 and Vibrio sp. ME6 ................................................47

5 Discussion......................................................................................................................................... 49

6 Conclusions and recommendations .................................................................................................. 54

7 References ........................................................................................................................................ 55

vi

List of Figures

Figure 2.1: Global trends of capture fisheries and aquaculture invention in the last 65 years ......... 3

Figure 2.2: Statistics of the bivalve molluscs production in the last sixty years in the marine

environment ................................................................................................................. 5

Figure 2.3: Morphological variation of the three different Mytilus species. .................................... 7

Figure 2.4: a) Inner anatomy of Mytilus edulis, in the upper margin a visible white posterior

adductor muscle (Wikipedia). b) General internal anatomy of the blue mussel, Mytilus

edulis. ........................................................................................................................... 8

Figure 2.5: a) Digestive system and respiratory structure of the blue mussel, Mytilus edulis and b)

shape of the crystalline style ........................................................................................ 9

Figure 2.6: Open circulatory system of bivalve molluscs. ............................................................. 10

Figure 2.7: a) Mussel haemocytes encapsulating the disease agent, Marteilia sydneyi; b) Massive

haemocyte infiltration in the mantle connective tissues of the hard clam, Mercenaria

mercenaria ................................................................................................................ 11

Figure 2.8: a) Spawning of Mytilus edulis, releasing milky white sperms and particulate eggs b)

Life cycle of the blue mussel, Mytilus edulis. ............................................................ 12

Figure 2.9: Life cycle, grow-out culture, harvesting, grading and marketing of blue mussel M.

edulis .......................................................................................................................... 13

Figure 2.10: AHL-mediated quorum sensing.. .............................................................................. 21

Figure 2.11: Chemical structure of quorum sensing signal molecules. ......................................... 23

Figure 2.12: Quorum sensing system in Vibrio campbellii.. .......................................................... 23

Figure 3.1: a) Challenge of mussel larvae with pathogenic vibrios Photobacterium sp. ME5 and

Vibrio sp. ME6, b) control for challenge test 1 and 2, c) larvae after 1 day from

control of challenge test 1 .......................................................................................... 33

Figure 3.2: a) Challenge of mussel larvae with experimental wild strain LGP32 and its quorum

sensing mutants, b) wild strain J2-9 and its quorum sensing mutants, c) larvae after 2

days challenge with quorum sensing mutants LGP32ΔluxM. .................................... 33

Figure 3.3: a) Challenge of mussel larvae with pathogenic vibrios ME5 and ME6 containing

cinnamaldehyde, b) challenge with experimental wild strain LGP32 and J2-9

containing cinnamaldehyde, c) larvae after 5 days challenge with ME6 at the

cinnamaldehyde concentration 1 µM. ........................................................................ 34

Figure 4.1: ERIC-PCR band patterns of V. crassostreae J2-9 and V. tasmaniensis LGP32 wild

types and their selected QS mutants, and of Photobacterium sp. ME5 and Vibrio sp.

ME6.. .......................................................................................................................... 35

Figure 4.2: Production of biofilms on polystyrene 96-well plates by wild type and quorum sensing

mutants of Vibrio crassostreae J2-9.. ......................................................................... 39


mutants of Vibrio tasmaniensis LGP32. ..................................................................... 40

vii

Figure 4.4: Production of exopolysaccharides by wild type V. crassostreae J2-9 and its QS

mutants. ...................................................................................................................... 40

Figure 4.5: Production of exopolysaccharides by wild type V. tasmaniensis LGP32 and its QS

mutants. ...................................................................................................................... 41

Figure 4.6: Survival of blue mussel larvae during five days of challenge with the mussel

pathogenic isolates Photobacterium sp. ME5 and Vibrio sp. ME6. .......................... 43

Figure 4.7: Bacterial density in the blue mussel larval rearing water during the five days of

challenge with Photobacterium sp. ME5 and Vibrio sp. ME6 .................................. 43

Figure 4.8: Survival of blue mussel larvae during five days of challenge with V. crassostreae J2-9

wild type and three of selected QS mutants (ΔluxM, ΔluxS and ΔcqsA) ................... 45

Figure 4.9: Survival of blue mussel larvae during five days of challenge with V. tasmaniensis

LGP32 wild type and three selected QS mutants (ΔluxM, ΔluxR and ΔluxS). .......... 46

Figure 4.10: Survival of blue mussel larvae challenged with V. crassostreae J2-9, V. tasmaniensis

LGP32, Photobacterium sp. ME5 and Vibrio sp. ME6, with and without

cinnamaldehyde. ........................................................................................................ 48

viii

List of tables

Table 2.1: Statistics of fisheries and aquaculture production and its utilization from the year 2009

till 2014 ................................................................................................................................. 4

Table 2.2: Commercially important aquaculture and mariculture fish/ shellfish species groups

with their global production in 2012. .................................................................................... 4

Table 2.3: Quorum sensing system of some pathogenic bacteria and their influence on virulence

factors. ................................................................................................................................. 24

Table 3.1: Overview of Vibrio crassostreae and Vibrio tasmaniensis strains used in this study. ........ 28

Table 3.2: PCR mastermix used for DNA amplification. ..................................................................... 29

Table 3.3: In vivo challenge test 1 of blue mussel larvae with bacterial strains ME5 and ME6. ......... 33

Table 3.4: In vivo challenge test 2 of blue mussel larvae with V. crassostreae, V. tasmaniensis and

a selection of quorum sensing deletion mutants. ................................................................ 34

Table 4.1: Swimming motility halo diameter (mm) (mean ± standard deviation of three replicates)

of wild types and QS mutants of Vibrio tasmaniensis LGP32 and Vibrio crassostreae

J2-9 after 24 hours of incubation on soft LB35 agar. ........................................................... 36

Table 4.2: Haemolytic activity (mean ± standard deviation of three replicates) of wild types and

QS mutants of Vibrio tasmaniensis and Vibrio crassostreae after 4 days of incubation

on LB35 agar with defibrinated sheep blood........................................................................ 36

Table 4.3: Caseinase activity (mean ± standard deviation of three replicates) of wild types and QS

mutants of Vibrio tasmaniensis and Vibrio crassostreae after 4 days of incubation on

LB35 agar with skim milk powder. ...................................................................................... 37

Table 4.4: Gelatinase activity (mean ± standard deviation of three replicates) of wild types and QS


LB35 agar with 0.5 % gelatin. .............................................................................................. 37

Table 4.5: Phospholipase activity (mean ± standard deviation of three replicates) of wild types

and QS mutants of Vibrio tasmaniensis and Vibrio crassostreae after 4 days of

incubation on LB35 agar with 1 % egg yolk emulsion. ....................................................... 38

Table 4.6: Lipase activity (mean ± standard deviation in three replicates) of wild types and QS


LB35 agar with 1 % tween80. .............................................................................................. 38

ix

List of abbriviations and units

Ab Antibiotics

AHL Acylated homoserine lactones

AI Autoinducer

AMPs Antimicrobial peptides

bp Base pair

BSA Bovine serum albumin

CAI Cholerae autoinducer

c-di-GMP Cyclic di-GMP (cyclic dimeric guanosine 3’,5’-monophosphate)

CFU Colony forming unit

DMSO Dimethylsulfoxyde

DNA De-oxyribonucleic acid

eDNA Extracellular de-oxyribonucleic acid

rDNA Ribosomal de-oxyribonucleic acid

EmpA Extracellular metalloproteses

EPS Extracellular polysaccharide

ERIC Enterobacterial Repetitive Intergenic Consensus

FAO Food and Agricultural Organization

FASW Filtered autoclaved seawater

FOC Fisheries and Oceans Canada

FSW Filtered seawater

Fur Ferric uptake regulator

HAI Harveyi autoinducer

HHL N-hexanoyl-L-homoserine lactone

Holo-ACP Holo-(acyl-carrier-protein)

HSP70 Heat shock protein 70

J2-9 Wild type Vibrio crassostreae

LB Luria-Bertani broth

LGP32 Wild type Vibrio tasmaniensis

ME5 Photobacterium sp.

ME6 Vibrio sp.

NADPH Nicotinamide Adenine Dinucleotide Phosphate Hydrogen

NC Negative control

NO Nitrogen oxides

OD Optical density

OMPs Outer membrane proteins

OMVs Outer membrane vesicles

P Statistical p-value

PCR Polymerase chain reaction

PHB poly-β-hydroxybutyrate

PLA Phospolipase A

x

PLC Phospolipase C

PRRs Pathogen recognition receptors

QS Quorum sensing

Rif Rifampicin

RNA Ribinucleic acid

mRNA Messanger ribonucleic acid

sRNAs Five small regulatory ribonucleic acids

ROS Reactive oxygen species

SPSS Statistical Package for the Social Sciences

T2SS Type II secretion system

TAE Tris-acetate-EDTA

TCBS Thiosulphate-citrate-bile salts-sucrose

WHO World Health Organization

µL Microliter

mL Milliliter

ºC Degree Celsius

gL-1

Gram per liter

L Liter

L mL-1

Larvae per milliliter

mgL-1

Milligram per liter

nm Nanometer

mm Millimeter

mM Millimolar

µM Micromolar

v/v Volume/volume

w/v Weight/volume

xi

Dedication

To my beloved mother and departed soul of my fa-

ther and sister, for your endless love and inherent

inspiration

And

To my beloved family members, for all care,

patience and encouragement

xii

Abstract

Vibrio crassostreae and Vibrio tasmaniensis are two important pathogens in mollusc aquaculture.

In this study, the impact of quorum sensing on the production of virulence factors, including

swimming motility, hemolysisn, caseinase, gelatinase, phospholipase, lipase, biofilm formation

and exopolysaccharide production was investigated. In addition, the link between quorum sensing

and virulence of V. crassostreae and V. tasmaniensis towards blue mussel (Mytilus edulis) larvae

was also determined. The non-toxic quorum sensing inhibitor cinnamaldehyde was used to check

whether it can protect the mussel larvae from the wild type V. crassostreae, V. tasmaniensis and

the blue mussel pathogenic isolates Photobacteriumm sp. ME5 and Vibrio sp. ME6.

The virulence factor plate assay revealed that two of the V. tasmaniensis mutants, ΔluxM and

ΔluxS, showed higher swimming motility than the wild type, and that for V. crassostreae, the

ΔcqsA mutant is less motile than the wild type. In addition, the luxR deletion mutant of V.

tasmaniensis has a higher biofilm production than the wild type. However, this difference was

relatively small and was not confirmed by a higher exopolysaccharide production. These results

can be explained similar to other vibrios in which some of virulence factors are regulated by

quorum sensing (negatively or positively), and others are independent of quorum sensing.

The in vivo challenge test results showed that both V. tasmaniensis LGP32 and V. crassostreae

J2-9 are pathogenic to blue mussel larvae (Mytilus edulis). In case of V. crassostreae, the luxS

deletion mutant was more virulent than the wild type, whereas the virulence of the other mutants

was not different from that of the wild type. In case of V. tasmaniensis, there were no differences

in virulence between the wild type and the quorum sensing mutants.

Finally, the quorum sensing inhibitor cinnamaldehyde did not protect mussel larvae from these

pathogens at the concentrations tested in this thesis (1 and 10 µM).

In conclusion, inhibition of the three channel quorum sensing system will not be effective to pro-

tect blue mussel larvae from V. crassostreae and V. tasmaniensis.

Keywords: virulence factors, quorum sensing, Vibrio crassostreae, Vibrio tasmaniensis, Mytilus

edulis

Chapter one Introduction

1

1 Introduction

Aquaculture is one of the continuously fastest-growing food-producing industries, with a produc-

tion of 93.4 million tonnes in 2014 (FAO, 2016). Despite of all splendid achievements, this indus-

try still fights with different constraints, among which the most important one are disease out-

breaks, which cause serious economic losses (i.e. tens of billions dollars in last two decades)

(FAO, 2016; Liao and Chao, 2009; Lima et al., 2013). Mussels are the third largest group of mol-

luscs in respect of production, and blue mussel (Mytilus edulis) culture is significantly contrib-

uting to the aquaculture industry (FAO, 2014a). Although hatchery technology is available and

although it has advantages over natural spat collection, still the mussel cultivation totally depends

on natural spat (Eggermont et al., 2014). The major problem is that hatchery production is not

cost effective due to unexpected mass mortalities in dense larval cultures, which might be caused

by opportunistic heterotrophic bacteria (Eggermont et al., 2014). Most of the bacteria that have

been isolated from blue mussels are Vibrio species (Garrido-Maestu et al., 2016). Splendidus

clade vibrios have been described several times in relation to the mollusc mortality outbreaks. V.

crassostreae and V. tasmaniensis are two important members of Splendidus clade vibrios that can

be pathogenic to molluscs (Travers et al., 2015). Both of these bacteria can cooperate in

polymicrobial infections; meaning that interactions of virulent and avirulent strains can facilitate

the disease caused by V. tasmaniensis (Gay et al., 2004b) and V. crasostreae (Lemire et al.,

2015). For example, injections of 106 or 4 × 10

4 V. crasostreae (J2-9) per animal (i.e. oyster) in-

duced 90 % and 0 % mortality respectively; whereas, in combination with the avirulent strain J2-

20, injection of 4 × 104 virulent strain J2-9 induced 70 % mortality (Lemire et al., 2015).

Defoirdt (2014) argued that the pathogenicity mechanisms of bacteria are controlled by several

virulence factors, including lytic enzymes, flagellar motility, extracellular polysaccharides and

biofilm production, iron acquisition and siderophore formation and secretion systems. Thus far,

little has been published on the virulence determinants of V. crassostreae. A major virulence fac-

tor of V. crassostreae J2-9 is an outer membrane protein, which was identified by Lemire et al.

(2015). The pathogenesis of V. tasmaniensis LGP32 is more well-defined, which causes extensive

lesions inside the translucent part of the muscle of Crassostrea gigas (Gay et al., 2004b). In addi-

tion, V. tasmaniensis LGP32 pathogenesis is mediated by the OmpU (outer membrane protein)

porin, which helps to internalize the bacteria into haemocytes (Duperthuy et al., 2010, 2011;

Travers et al., 2015). The virulence of V. tasmaniensis LGP32 is also controlled by secreted tox-

ins, i.e. metalloprotease (Vsm) and hydrolases from outer membrane vesicles (OMVs) (Binesse et

al., 2008; Le Roux et al., 2007). Apart from this, other virulence factors have not been character-

ized for V. crassostreae and V. tasmaniensis.

The production of virulence factors is controlled by several regulatory mechanisms, such as quor-

um sensing (QS) (Defoirdt, 2014). Quorum sensing regulates gene expression in response to small

signal molecules (autoinducers) (Defoirdt, 2014; Ng and Bassler, 2009). Like many other vibrios,

V. crassostreae contains a three channel quorum sensing system including the synthase/receptor

pairs LuxM/LuxN, LuxS/LuxP-Q and LuxCqsA/LuxCqsS (Lemire et al., 2015). Similarly, the

quorum sensing system of V. tasmaniensis was also found to have a homologue of the syn-

Chapter one Introduction

2

thase/receptor pairs LuxM/LuxN, LuxR/LuxI, LuxS/LuxP-Q (Tait et al., 2010). However, the

interaction between virulence and quorum sensing has not been reported yet in V. crassostreae

and V. tasmaniensis.

In this thesis we aimed at determining the impact of quorum sensing on the virulence of Vibrio

crassostreae and Vibrio tasmaniensis in order to evaluate whether quorum sensing inhibition

would be effective to control vibriosis caused by these pathogens in molluscs. In the first part of

this thesis, we investigated the impact of quorum sensing on several putative virulence factors of

V. crassostreae and V. tasmaniensis in vitro. In the second part, blue mussel larvae were chal-

lenged with V. crassostreae and V. tasmaniensis wild types and quorum sensing mutants in order

to determine their virulence in vivo. In this part, cinnamaldehyde was also used as a second strate-

gy to determine the impact of quorum sensing on the virulence of V. crassostreae and V.

tasmaniensis towards blue mussel larvae.

Chapter two Review of literature

3

2 Review of literature

2.1 Overview of aquaculture

The total world population is increasing day by day with the present number of 7.0 billion humans

in the year 2011, which is 0.1 billion more than that of the previous year (FAO, 2012). This in-

crease of population has been fairly constant at a rate of 1.6 % in the last decade (FAO, 2014a)

which leads to a significant increase of the demand for food as well as high quality proteins. In-

deed, with the increasing population and protein demand, the fish food consumption rate has

grown from 117.3 million tons in the year 2007 to 136.2 million tons in the year 2012 (FAO,

2014a). The growing population occupied most of the surface area of the globe which resulted in

reduction of lands dedicated for agriculture. In the past, the major protein source was livestock

and poultry; in the future the land based production alone cannot meet the growing protein re-

quirement. So, it is important to promote promising sectors for future protein supply for the large

population of the world. Hence, the fisheries and aquaculture sector is a potential source to meet

the high protein demand of the world and therefore, will play an increasing and essential role in

the livelihoods of millions of people.

Figure 2.1: Global trends of capture fisheries and aquaculture invention in the last 65 years (FAO,

2016).

Capture fisheries has reached its maximum potential in the last decades (90.8 million tons in

2007, remained stable 93.4 million tons in 2014 (FAO, 2016)). Sustaining and increasing fish

supplies will thus not be possible. Captured fisheries and aquaculture together supplies a total

production of 167.2 million tonnes, of which 73.8 million tons (44.14 %) is supplied by aquacul-

ture production (FAO, 2016). Aquaculture contributes even 48.9 % of the food fish consumed by

people all over the world (FAO, 2014a).

According to FAO (2014a), in 2010 16.7 % of the global animal protein and 6.5 % of all con-

sumed proteins have been supplied from fish. Seafood (fish, shellfish, macro/ micro algae) is con-

sidered as a good source of proteins, fatty acids, nutrients and minerals. Some of these nutrients


4

have role in fecundity, brain development and in preventing heart disease (due to presence of ω-3

fatty acid) (Stanton, 2011).

Table 2.1: Statistics of fisheries and aquaculture production and its utilization from the year 2009 till

2014 (FAO, 2016).

2009 2010 2011 2012 2013 2014

Million tons

PRODUCTION

Capture

Inland 10.5 11.3 11.1 11.6 11.7 11.9

Marine 79.6 77.8 82.6 79.7 81.0 81.5

Total capture 90.1 89.1 93.7 91.3 92.7 93.4

Aquaculture

Inland 34.3 36.8 38.7 41.9 44.8 47.1

Marine 21.4 22.3 23.3 24.7 25.5 26.7

Total Aquaculture 55.7 59.0 62.0 66.6 70.3 73.8

TOTAL WORLD FISHERIES 145.8 148.1 155.7 158.0 162.9 167.2

UTILIZATION

Human consumption 123.7 128.2 131.2 136.2 141.5 146.3

Non-food uses 22.1 19.9 24.5 21.7 21.4 20.9

Population (billions) 6.8 6.9 7.0 7.1 7.2 7.3

Per capita food fish supply (kg) 18.1 18.5 18.7 19.2 19.7 20.1

Of the 66.6 million tonnes of farmed fish and shellfish that was produced in 2012, 66.3 % (44.2

million tonnes) were finfish species, 9.7 % (6.4 million tonnes) were crustaceans, and 22.8 %

(15.2 million tonnes) were molluscs (FAO, 2014a; Table 2.2).

Table 2.2: Commercially important aquaculture and mariculture fish/ shellfish species groups with

their global production in 2012 (FAO, 2014a).

Inland aquaculture Mariculture Quantity subtotal Value subtotal

(Million tonnes) (Million tonnes) (Million

tonnes)

(Percentage

by volume)

(US$

million)

(Percentage

by value)

Finfish 38.599 5.552 44.151 66.3 87499 63.5

Crustaceans 2.530 3.917 6.447 9.7 30864 22.4

Mollusks 0.287 14.884 15.171 22.8 15857 11.5

Other species 0.530 0.335 0.865 1.3 3512 2.5

Total 41.946 24.687 66.633 100 137732 100

Mussels are considered as the third largest cultured mollusc species and its aquaculture activity is

significantly increasing (Figure 2.2). Spain, New-Zealand and China contribute for about 95 % of

the worlds’ mussel production. The blue mussel Mytilus edulis contributed for 184,429 tonnes to

the global mussel production of 250,029 tonnes in 2012 (FAO, 2014a). Due to climatic change

and occurrences of algal blooms, capture production of blue mussel has been gradually decreasing

in the past decades (Smaal, 2002); therefore, the demand for farmed blue mussel progressively

increases.


5

Figure 2.2: Statistics of the bivalve molluscs production in the last sixty years in the marine

environment (FAO, 2014a).

2.2 The impact of bacterial diseases on aquaculture

With the development of aquaculture during the last decades, the industry has encountered serious

constraints; including competition for resources and technology, unpopularity of super-intensive

aquaculture, lack of a proper biotechnological approach and environmental considerations, how-

ever notably the most dangerous is disease outbreak (Liao and Chao, 2009; Lima et al., 2013).

The intensification of aquaculture leads to (re)emergence of diseases caused by various pathogens

(viruses, bacteria, parasites) with new diseases or existing pathogens infecting new animals every

year (Murray and Peeler, 2005). Consequently, disease emergence has become a serious concern

in both the captive and the cultured fisheries sector due to the continuous environmental pres-

sures, the direct impact of human activities and the risk of pathogen spread from aquaculture

(Naish et al., 2007). During the past century, the spread of pathogens arose in a similar trend with

the rise of the intensive aquaculture. Naish et al. (2007) stated that the increased emergence of

various new diseases in fish and shellfish aquaculture might be due to the increased global

movement of aquatic animals and their products and to anthropogenic stress (biotic and abiotic

source) towards the aquatic ecosystem.

2.2.1 Environmental stress and disease outbreaks in aquaculture

Susceptibility and occurrence of disease in animals mainly depends on the environment in which

the animal is cultured (Jirtle and Skinner, 2007). Understanding this relationship is important for

the disease control in aquaculture. Diseases caused by pathogenic or opportunistic bacteria have

been problematic in the intensive culture of molluscs, fish and shrimp leading to losses up to 100

% (Muroga, 2001; Soto-Rodriguez et al., 2003). Obligate pathogens and exotic diseases are not

the only factors responsible for disease outbreaks in aquaculture, environmental stress can also

interfere with the normal physiology of the animals; hence, increasing the opportunity for disease

outbreak. Snieszko (1974) stated that when susceptible animals are exposed to pathogens in

stressful environmental conditions, then the risk of disease outbreak is higher. Reducing stress is a

key factor in preventing disease outbreaks in aquaculture.


6

2.2.2 Disease caused by pathogenic vibrios

Among the groups of pathogenic microorganisms that cause serious losses in aquaculture, bacte-

ria are taking the lead with devastating economic effects. The most pathogenic bacteria causing

disease in aquaculture belong to the family of Vibrionaceae, causing vibriosis (McHenery and

Birkbeck, 1986), which results in high mortalities of fish and shellfish cultures (Austin and

Zhang, 2006).

Vibrios are Gram-negative, rod shaped, motile, mesophilic, chemo-organotrophic and facultative

anaerobe bacteria, best known as cause for food borne diseases (Farmer, 1992; Thompson et al.,

2005). By molecular technique of 16S rDNA gene sequence analysis, vibrios are considered to

belong to the Gammaproteobacteria. They are abundantly found in aquatic habitats, often in asso-

ciation with eukaryotes. Vibrio association ranges from mutualistic like Vibrio fischeri-bobtail

squid (Ruby, 1996) to pathogenic like V. cholerae in humans (Wachsmuth et al., 1994). Probiotic

strains of Vibrio for fish and shellfish also have been documented (Verschuere et al., 2000). Dif-

ferent Vibrio strains have been identified in bivalves and have been reported causing bacterial

diseases (Travers et al., 2015).

2.2.3 Problems related to the use of antibiotics in aquaculture

Antibiotics are natural or synthetic drugs that are used to kill (bactericidal activity) or inhibit (bac-

teriostatic activity) bacterial growth. They are used for the treatment of infectious diseases in hu-

man, animals and plants (Burridge et al., 2010; Serrano, 2005). Although most of the antibiotics

have a rather low toxicity, there are significant environmental concerns with the widespread use

of antibiotics (Burridge et al., 2010). Most of antibiotics consist of stable chemical compounds,

that are not dissociated in the body of animals and that remain active for a long time after excre-

tion as stool or urine as well as passing to the environment with uningested food particles

(Aarestrup, 2006; Sørum, 2006). The persistence of antibiotic residues in the environment due to

a massive use/ misuse/abuse in an attempt to control infectious diseases increased the selection of

antibiotic resistant strains. This leads to the widespread resistance to conventional antibiotics and

increasing reports of non treatable infectious diseases in humans and animals (Defoirdt et al.,

2010; Defoirdt et al., 2011).

Bacteria can become resistant through spontaneous mutations, and horizontal gene transfer which

can be stimulated between different species and genera including marine bacteria, human and fish

pathogens (Baquero et al., 2008; Silbergeld et al., 2008). Those bacteria carry resistance genes

and can grow rapidly since their competitors are removed. Hence, rapid evolution of antibiotic-

resistant strains of pathogens has been described in aquaculture system together with their spread

to strains of importance in human disease (Defoirdt et al., 2007a; Defoirdt et al., 2010). Antibiot-

ics may also affect the biological diversity of phytoplankton and zooplankton, including algal

toxicity and growth inhibition by different antibiotics (Ferreira et al., 2007; Lützhøft et al., 1999);

these changes may affect animals and human health when contaminated shellfish and fin fishes

are consumed (Defoirdt et al., 2007a). Antibiotics are also harmful for fish culture because of

their impact on non-target species (Burridge et al., 2010). Often non-target fauna like fish, crusta-


7

ceans and molluscs are a pathway of bringing antibiotic residues into the human population (Al-

derman and Hastings, 1998; Cabello, 2006).

In addition the presence of residual antibiotics in aquaculture products can cause problems by

altering the human gut microbiota; generating risks of allergy and toxicity (Cabello, 2006).

Hence, commercialized aquaculture products cause problems in both producing and importing

countries (Defoirdt et al., 2007a). To prevent these global problems related to antibiotics, the use

of antibiotics should be limited and a focus should be put on the development of new control

strategies of pathogenic bacteria to ensure a good productivity and sustainable aquaculture indus-

try (Defoirdt, 2013).

2.3 The blue mussel (Mytilus edulis)

2.3.1 Taxonomy

Kingdom: Animalia

Phylum: Mollusca

Class: Bivalvia

Subclass: Pteriomorphia

Order: Mytiloida

Family: Mytilidae

Subfamily: Mytilinae

Genus: Mytilus

Species: M. edulis (Linnaeus, 1758)

The taxonomy of the blue mussel (Mytilus) has been a controversial and unclear issue for decades.

Several authors recognized more than 30 designations, because of their extreme phenotypic shell

plasticity characteristic of the genus (Beaumont et al., 2007; Gardner and Thompson, 2009).

However, based on biochemical and genetic variations by using molecular markers (Borsa et al.,

2012; Skurikhina et al., 2001) three smooth-shell mussels species have been identified, namely M.

edulis, M. galloprovincialis and M. trossulus (Figure 2.3). Whereas, M. edulis and M.

galloprovincialis are more closely related (Hilbish et al., 2000). When their distribution overlaps

they can hybridize and produce fertile hybrids (Beaumont et al., 2007; Gardner and Thompson,

2009).

Figure 2.3: Morphological variation of the three different Mytilus species.


8

2.3.2 Anatomy

2.3.2.1 External anatomy

Two hinged valves are joined together by a ligament on the outside to form the external shell of

the mussel. The shells are closed when necessary by strong internal muscles. The shell shape of

the blue mussel is an elongated triangle; the longest dimension is ca. 7-10 cm and the colour is

dark blue or black and has a sculpture of fine concentric lines (Newell, 1989) (Figure 2.3). The

shell has an inner prismatic layer, a chalky white crystal like middle layer and a pigmented skin

like outer periostracum layer. The layers are formed by an inner iridescent layer of nacre com-

posed of calcium carbonate, which is continuously secreted by the mantle (Wikipedia; Carter,

1980).

2.3.2.2 Internal anatomy

The internal anatomy of mussels consists of distinctive characteristic organs (Figure 2.4). A dark-

ly pigmented foot is found in the center of the visceral mass which is used as an anchor (Newell,

1989; Wikipedia). The extended mantle, which is intact at the margin of both valves of the shell

except at the inhalant and exhalent siphon (Wilbur and Saleuddin, 1983). The gills are situated at

the ventral side of the mussel and have a respiratory function. The gills also filter out the food

particles and guide them towards the mouth region (Newell, 1989). The labial palps surrounding

the mouth are involved in the selection of food particles. An oesophagus leads the food to the

anterior part of the stomach (Ward and Shumway, 2004).

An inert and chitinous gastric shield is present in the stomach, as is a cristalline style that rotates

(Morton, 1983). Post-ingestive selection happens in the stomach; digestible organic and indigesti-

ble inorganic particles are selected, subjected for further processing in the mid-gut and directed

outside through the anus and finally the exhalent siphon (Gosling, 2004; Ward and Shumway,

2004). The stomach is completely surrounded by the digestive gland.

Figure 2.4: a) Inner anatomy of Mytilus edulis, in the upper margin a visible white posterior adductor

muscle (Wikipedia). b) General internal anatomy of the blue mussel, Mytilus edulis (FOC, 2003).

a b


9

2.3.2.3 Digestive system

Mytilus edulis, is an active filter feeder which mainly ingests organic matter, phytoplankton, bac-

teria, toxins, parasite larvae and chemical pollutants, acting as scavenger (Bayne, 1983). It is often

used as a biomarker in environment monitoring programs (Brenner et al., 2014). Water enters into

the mantle and branchial chamber through inhalant siphon by the activity of cilia of gill filaments

(Newell and Jordan, 1983) and exits the exhalant siphon by passing through the supra-branchial

chamber where particles larger than 5 µm are retained efficiently (Møhlenberg and Riisgård,

1978) (Figure 2.5a). Materials with lower organic contents (Kiørboe et al., 1980) and larger par-

ticles are rejected by the gills and ejected as pseudofaeces through the inhalant siphon and carried

away by the current activity of the exhalant siphon (Beninger et al., 1999; Jørgensen, 1981). The

ingested particles in the stomach are processed by a mechanical and enzymatic attack of continu-

ously revolving crystalline style, and are subsequently directed to the ducts of the digestive diver-

ticula following endocytosis into the digestive cells, subject to intracellular digestion (Bayne,

1976; Newell, 1979) (Figure 2.5b). Assimilated food particles are absorbed by the blood stream

through diffusion. However, particles of lower nutritional value and excess particles form intesti-

nal faeces by passing through digestive diverticula are removed from the stomach into the mid-gut

(Newell, 1979; Riisgård et al., 2011). Glandular faeces are ejected into the duct following the

stomach and mid-gut. In the mid-gut both the glandular and intestinal faeces are coated by mucus

faecal ribbon and passes through the anus to outside (Newell, 1979; Riisgård et al., 2011).

Figure 2.5: a) Digestive system and respiratory structure of the blue mussel, Mytilus edulis and b)

shape of the crystalline style (Xu, 2002).

2.3.2.4 Cardiovascular and respiratory system

Like other bivalves, mussels have an open circulatory system, which means that after passing

through the arteries the haemolymph bathes all the organs and joins again in veins that empty into

sinuses before coming back to the heart (Pruzzo et al., 2005; Gosling, 2004) (Figure 2.6). The

mussels heart consists of two auricles and a single ventricle. The blood enters the auricles through

a big vein, passes through the auriculo-ventricular valves to the ventricle and leaves the ventricle

through the anterior aorta which branches into several arteries providing the viscera and mantle

with haemolymph. Both gills and mantle have a rich haemolymph supply which makes them a

suitable site for respiration. Haemolymph flows via the afferent gill vein to the gills, circulates

within the hollow tubular filaments. During this passage through the gills, haemolymph gets rich

a b


10

with oxygen through diffusion. Oxygenated hemolymph flows through the efferent gill veins via

the kidney or directly to the heart. The oxygenation efficiency is low (1-13 %), although depend-

ing on the heart rate it can increase up to 30 % (Bayne et al., 1976).

Figure 2.6: Open circulatory system of bivalve molluscs.

2.3.2.5 Immune system

Similar to other invertebrates the immune system of bivalves is an innate immune system, a rela-

tively simple system in comparison to the adaptive immune system of vertebrates (Galloway and

Depledge, 2001; Novoa and Figueras, 2012). An innate immune system lacks the immune

memory and cannot produce antibodies at the initial contact of pathogens (Novoa and Figueras,

2012; Ottaviani, 2011). The shell and mucus of bivalves act as a primary defense barrier; in addi-

tion, the mucus contains bactericidal enzymes (Gliński and Jarosz, 1996; Ratcliffe et al., 1985).

The innate immune system contains cellular (haemocytes) and humoral components (Galloway

and Depledge, 2001; Novoa and Figueras, 2012).

Haemocytes are cellular response components which can recognize non-self particles; ingest,

destroy and expel them by phagocytosis as well as by cytotoxic reaction (Carballal et al., 1997).

During these reactions they release lysosomal enzymes and anti-microbial peptides. Respiratory

burst by production of oxygen radicals and the action of NADPH-oxidase, is also described in

bivalves. Molecular reactive oxygen species are reduced into reactive oxygen intermediates, in-

cluding superoxide anion, hydrogen peroxide, and other intermediated compounds with high bac-

tericidal activity (Novoa and Figueras, 2012; Pruzzo et al., 2005). Mussel haemocytes are also

able to produce nitrogen radicals (Ottaviani et al., 1993). When the pathogens are bigger than the

haemocytes, they are removed by recruiting fibroblast deposit mucopolysaccharide residues and

fibrous material to form a glycoprotein associated reticulum (Ratcliffe et al., 1985) (Figure 2.7).

Humoral components of the innate immune response are present in the plasma. They include an-

timicrobial peptides (AMPs), protease inhibitors, lysozyme and pathogen recognition receptors

(PRRs) (Buchmann, 2014; Novoa and Figueras, 2012; Song et al., 2010).


11

Figure 2.7: a) Mussel haemocytes encapsulating the disease agent, Marteilia sydneyi; b) Massive

haemocyte infiltration in the mantle connective tissues of the hard clam, Mercenaria mercenaria

(Allam and Raftos, 2015).

2.3.3 Life cycle

2.3.3.1 Reproduction

Blue mussels are diecious, whereas in rare instances some individuals with both sexes (hermaph-

rodites) may occur in the population (FOC, 2003; Seed, 1976). Mussels generally attain sexual

maturity after one year. However, certain environmental conditions (e.g. longer exposed time to

air) may result in slower growth and a longer time to reach reproductive maturity, which can even

be delayed until the second year (Newell, 1989). Seasonal changes of environmental factors; in-

cluding food availability and water temperature, can also influence reproductive maturity (Gray et

al., 1997; Newell et al., 1982). The gonad development of mussels begins in autumn and contin-

ues through the winter in European areas. In spring and summer, spawning takes place (Gabbott

and Bayne, 1973; Pieters et al., 1980). The fecundity of the blue mussel depends on environmen-

tal conditions (temperature, food supply, tidal exposure etc.), size and the site of reproduction and

can vary from year to year (Bayne et al., 1983). Spawning is synchronus which ensures that eggs

and sperms are present in the water column concurrently (Newell, 1989). Males release sperms

first which stimulates the females to release the eggs (Newell and Thompson, 1984) (Figure

2.8a). Spermatozoa swim freely in the water column and fertilize the eggs by attaching with an

acrosome filament that enables them to penetrate the eggs (Bayne, 1976; Newell, 1989).

2.3.3.2 Larval development

The larval development of blue mussels takes between 15 and 35 days, depending on environ-

mental conditions (Newell, 1989), of which water temperature and salinity are considered to be

the limiting factors (Bayne, 1976, 1983). Within half an hour after fertilization, the zygote starts

to develop into a two-celled stage, proceeding into the multi-celled blastula and gastrula stages.

Within 24 hours after fertilization the ciliated embryo differentiates to form the trochophore larva,

which is motile but non-feeding, and nutrients are supplied from the yolk (Newell, 1989). The

larvae then develop into the veliger stage called D larvae which have a functional mouth and ele-

mentary system. A group of cilia at the anterior side is later enlarged to form the velum, which is

used to filter particles, mostly nano-plankton (Helm et al., 2004; Newell, 1989). The veliger lar-


12

vae continue to develop into a pediveliger larvae; which is characterized by the development of a

photosensitive eye spot and an elongated foot with a byssal gland. They gain little ability to swim

horizontally, controlled by environmental stimuli (especially the salinity gradient change in estu-

aries) (Newell, 1989; Wood and Hargis, 1971).

Figure 2.8: a) Spawning of Mytilus edulis, releasing milky white sperms and particulate eggs b) Life

cycle of the blue mussel, Mytilus edulis.

After full development of the pediveliger, the foot extends and makes contact with a filamentous

substrate, crawls over it and attaches loosely (Bayne, 1976). When they find a suitable substrate

the larvae metamorphosed into the juvenile form also termed plantigrade (Newell, 1989), leading

to the loss of the velum (Balseiro et al., 2013); grown up to 1.5 mm shell the plantigrade larvae

release themselves from the filamentous substrate and is carried passively by current (Bayne,

1976) or by active crawling (Ytrøy, 2008) to the bottom. They secret byssus threads for settlement

either on the substrate or directly on to the mussel shells (Bayne, 1976; Tyler-Walters, 2008)

(Figure 2.8b).

2.3.4 Blue mussel cultivation

Mussel farming has expanded and developed as an aquaculture industry in many parts of the

world, including Europe, Asia, Australia and North America. Historically, two important mussel

species M. edulis and M. galloprovincialis have been cultured in Europe. European countries con-

tribute for half of the total mussel production of the world; in 2005 the global production of the

blue mussel was 391,210 tonnes, of which 365,764 tonnes (93.45 %) was produced in Europe

(FAO, 2008). Spain is by far the largest (300000 tonnes annually) mussel producer (Beaumont et

al., 2007), more than the combined total of the Netherlands, France, Italy, Ireland and UK (FAO,

2006). Traditionally, wild seeds of mussels are collected (around 20 mm shell length) and cul-

tured to a marketable size of 4–7 cm in 1–3 years. There are three main culture methods, bottom

culture, bouchots culture and suspended rafts or long lines culture (Beaumont et al., 2007) (Fig-

ure 2.9).

There is little interest in developing hatchery culture of mussels in Europe. However, in New

Zealand one commercial hatchery on greenshell mussel (Perna canaliculus) continuously pro-

vides spats to the rope culture farmers (Beaumont et al., 2007; Galley et al., 2010). Although, it is

not cost effective yet (Helm et al., 2004); hatchery technology to produce mussels has several

a b


13

advantages over wild caught spat, such as natural stock conservation, growth of a particular strain,

genetic enhancement, development of disease resistance populations and a reliable supply of spats

(FAO, 2014b). Nevertheless, the major bottleneck is the unsuspected mass mortality of mussels in

dense larval cultures (Eggermont et al., 2014) probably of microbial origin as described in many

other bivalve species (Beaz-Hidalgo et al., 2010; Genard et al., 2013).

Figure 2.9: Life cycle, grow-out culture, harvesting, grading and marketing of blue mussel M. edulis

(FAO, 2014b).

2.3.5 Diseases in bivalve and blue mussel cultivation

Mass mortalities caused by microbial diseases are recognized as the most significant constraints

of the shellfish aquaculture industry (Defoirdt, 2013; Paillard et al., 2004). Disease outbreak can

occur in any life-stage or production stage (Paillard et al., 2004; Travers et al., 2015).

2.3.5.1 Diversity of pathogens

A variety of pathogens can cause disease in bivalves, including bacteria, viruses, parasites (i.e.

protozoans, trematodes), and fungi (Paillard et al., 2004; Thieltges, 2006; Travers et al., 2015).

Parasite-related mortalities of bivalves are mainly linked to protozoans, including Perkinsus,

Haplosporidium, Marteilia and Bonamia (Paillard et al., 2004; Robledo et al., 2014). Other para-

sitic agents that have been reported are: Turbonilla sp. (Boglio and Lucas, 1997), Polydora ciliate

(Buschbaum et al., 2007), trematodes (Renicola roscovita) (Mouritsen, 2002; Thieltges, 2006)

and polychaete (Stefaniak et al., 2005). Parasites previously found in mussels (Mytilus

galloprovincialis) are protists (Marteilia refringrns), trematodes (Proctoeces maculates),

microsporidians (Steinhausia mytilovum), flatworms (Urastoma cyprinae), copepods (Mytilicola

intestinahs), ciliates (Ancistrum mytili) (Bignell et al., 2011; Villalba et al., 1997), turbellarians

(Urastoma cyprinae) (Robledo et al., 1994), rickettsiae and ascetosporans (Marteiliu) (Bignell et


14

al., 2011; Figueras et al., 1991). Pea crab (Pinnotheres novaezelandiae) is a common parasite in

the green-lipped mussel (Perna canaliculus) (Trottier et al., 2012). A trematod parasite

(Prosorhynchus squamatus) was also identified in blue mussel Mytilus edulis (Coustau et al.,

1993).

Viruses can be bio-accumulated in the tissues of bivalves after filter feeding; some can cause mor-

talities (Renault and Novoa, 2004). Important viruses that have been found in bivalves are: irido-

like virus, harpesviruses, retroviruses, papovaviridae, togaviridae, reoviridae, birnaviridae and

picornaviridae (Bower, 2001; Farley, 1978; Rasmussen, 1986; Renault and Novoa, 2004). Viruses

have been found in mussels as well: rotavirus, hepatitis A virus (Abad et al., 1997; Enriquez et

al., 1992) and picornalike virus (Rasmussen, 1986).

Bacterial pathogens can seriously affect bivalve aquaculture. Bacterial pathogens are commonly

found associated with the larval stages and can cause high mortalities in hatcheries as well as in

natural beds (Paillard et al., 2004; Travers et al., 2015). The most important disease causing

agents belong to the genus Vibrio; including Splendidus clade vibrios, e.g. V. Crassostreae, V.

tasmaniensis, V. cyclitrophicus, V. celticus and V. Splendidus (Le Roux et al., 2015; Travers et

al., 2015; Vanhove et al., 2015), Harveyi clade vibrios, e.g. V. harveyi, V. campbellii, V.

parahaemolyticus, V. alginolyticus, V. rotiferianus, V. natriegens, V. Communis, V. azureus, V.

owensii, and V. Sagamiensis and V. Mytili (Ruwandeepika et al., 2012; Saulnier et al., 2010;

Thompson et al., 2004; Travers et al., 2015) or the species V. aestuarianus, V. tubiashii, V.

coralliilyticus, V. Tapetis (Travers et al., 2015) and rickettsia (Crosson et al., 2014). Several spe-

cies of vibrios have been isolated from blue mussels, including V. alginolyticus and V.

anguillarum, which have been reported to cause toxicity to haemocytes in blue mussels (Lane and

Birkbeck, 1999; Nottage and Birkbeck, 1990). Romero et al. (2014) noted that V. splendidus and

V. aestuarianus isolates are only moderately pathogenic for mussels (Mytilus galloprovincialis)

injected intramuscular and almost none in bath challenge tests, and little mortality occurred only

at high bacterial concentration and in adverse environmental conditions.

2.3.5.2 Larval diseases

Larvae of bivalves have to pass several stages of metamorphosis. All life stages are vulnerable to

disease (Beaz-Hidalgo et al., 2010). The most common disease of larvae is bacillary necrosis. It

was first identified in clam larvae (Guillard, 1959) and was later on also detected in Crassostrea

virginica, Ostrea edulis, Mercenaria mercenaria, Argopecten irradians and Teredo navalis

(Tubiash et al., 1965). The bacteria which are associated with bacillary necrosis disease are V.

alginolyticus, V. tubiashii, V. anguillarum ((Tubiash et al., 1965; Tubiash and Otto, 1986), V.

pectinicida (Lambert et al., 1999) and V. splendidus (Gómez-León et al., 2005). The disease ex-

posing period of bacillary necrosis is swift and dramatic, within 4-5 hours after exposure. Typical

symptoms are reduced motility and quiescent lying with either an extended velum or a rudimen-

tary foot. Swarming of bacteria around the larvae, originating from the discrete foci on the mar-

gins of the larvae is the most common pathognomonic sign of bacillary necrosis (Tubiash et al.,

1965).


15

Bivalve larval diseases caused by vibrios occur in three patterns, termed as Pathogenesis I, II and

III (Elston and Leibovitz (1980). In pathogenesis I, all the larval stages are affected and the most

pronounced symptoms are colonization of the mantle, reduced motility and invasion of the viscer-

al cavity. During pathogenesis II, the early stage of veliger larvae are affected with abnormal

swimming, velum disruption and extension. However, the larvae remain active before the bacteria

invade the organs of the digestive tract. At that point, signs of visceral atrophy can be seen. In

pathogenesis III, the late veliger or pediveliger larval stage is affected and the larvae become im-

mobile. In addition, progression and extension of the visceral atrophy occurres and lesions in the

organs of digestive tract can be seen. Another prominent/major sign of vibriosis in hatcheries is

sedimentation of immotile larvae called “spotting” (Beaz-Hidalgo et al., 2010).

Many bivalves (e.g. oysters, clams and scallops) have been found with symptoms of bacillary

necrosis, however it was not observed for the larvae of Mytilus edulis yet (Luna-González et al.,

2002), in contrast to M. galloprovincialis (Anguiano-Beltrán et al., 2004). V. splendidus and V.

coralliilyticus/neptunius-like isolates have been reported to be able to cause mortalities in green-

shell mussel larvae (Perna canaliculus) (Kesarcodi-Watson et al., 2009).

2.3.5.3 Diseases in juvenile and adult bivalves

Juvenile and adult bivalves can be infected with different types of bacterial diseases, including

summer mortality (Beaz-Hidalgo et al., 2010), brown ring disease (Beaz-Hidalgo et al., 2010),

nocardiosis (Travers et al., 2015) and aka juvenile oyster disease (Travers et al., 2015).

Summer mortality merely affects oysters. It is caused by several Vibrio species, including V.

splendidus (Gay et al., 2004a, b; Lacoste et al., 2001), V. aestuarianus (Garnier et al., 2008), V.

harveyi (Allain et al., 2009). Bacteria are not the only causative agent of summer mortality in

oysters, it occurs due to a combination of a complex interaction of opportunistic infectious agents

together with the physiological and/or genetic condition of the host and certain environmental

factors (Labreuche et al., 2006; Paillard et al., 2004; Pruzzo et al., 2005).

Brown ring disease is caused by V. tapetis and affects adult clams (R. philippinarum and R.

decussatus) (Paillard, 2004; Paillard et al., 1994) and oysters (C. virginica) (Allam et al., 2001,

2006). This disease causes an alteration of the calcification process and the appearance of

conchiolin consisted brown deposits between the edge of the pallial line and the shell (Borrego et

al., 1996). Brown ring disease happens more frequently at 15 ºC and at a salinity of 20 ppt (during

spring season) (Paillard et al., 1994, 2004; Reid et al., 2003).

Nocardiosis disease is caused by N. crassostreae and occurs in the Pacific oyster C. gigas and the

European flat oyster Ostrea edulis (Travers et al., 2015; Elston et al., 1987). In some locations

infection and mortality occur due to this disease during the summer and fall months when temper-

ature is generally over 20 ºC (Bower et al., 2005; Friedman et al., 1991, 1998).

Aka juvenile oyster disease is caused by Roseovarius crassostreae (Boettcher et al., 1999) and

damnages oyster haemocytes, subsequently causing mortality (Gómez-León et al., 2008). The

symptoms of this disease are mantle retraction, epithelial degradation, lesions and infiltration of


16

haemocytes into the pallial cavity (Bricelj et al., 1992; Ford and Borrero, 2001; Maloy et al.,

2007).

Most of the diseases mentioned above were described for oysters, clams and scallops. A few de-

scriptions of Vibrio-interactions were associated with Mytilus-species, however only in vitro ef-

fects (e.g. ROS and NO production, AMP expression, HSP70 gene expression etc.) (Cellura et al.,

2007; Ciacci et al., 2010; Nottage and Birkbeck, 1990). No specific bacterial pathogens for M.

edulis, causing mass mortality have been described so far. However, the addition of organic mat-

ter in the mussel rearing water can enhance the growth of heterotrophic bacteria causing mass

mortality of the adult mussel M. edulis (Eggermont et al., 2014).

2.4 Virulence factors of vibrios

Virulence factors are gene products that allow the pathogens to infect and damage the host, in-

cluding products involved in motility and adhesion of the pathogens to the host, protection from

host defence mechanism and host tissue degradation, iron acquisition and toxins (Defoirdt, 2014).

Virulence factors control the infectious cycles of pathogenic bacteria by promoting the pathogens

entry, its growth and reproduction in the host’s body and exit from the host’s body (Defoirdt,

2014; Donnenberg, 2000). There are several factors which enable pathogens to confer virulence

activities which include swimming motility and chemotaxis, extracellular polysaccharide produc-

tion, biofilm formation and production of lytic enzymes (e.g. hemolysin, caseinase, gelatinase,

lipase and phospholipase) and these factors will be discussed further.

2.4.1 Motility and chemotaxis

Colonization and adherence to the host’s surface are crucial mechanisms to infect the host. This is

achieved by flagella that empower the bacteria to overcome negative electrostatic forces and serve

as an initial interaction of the bacteria to the host (Defoirdt, 2014; Haiko and Westerlund-

Wikström, 2013). Flagella perform these activities by acting as a helical propeller by a specialized

rotating mechanism (McCarter, 2004). The rotating mechanism is powered by an electrochemical

H+ or Na

+ gradient (Schuster and Khan, 1994). Some pathogenic bacteria (e.g. V. harveyi) have a

dual flagellar function; in liquid medium they swim with a single polar flagellum and in the more

viscous medium they swarm with many lateral flagella (Gode-Potratz and McCarter, 2011;

McCarter, 2004; Yang and Defoirdt, 2015). Flagellar motility has been investigated as an im-

portant virulence factor during the initial infection and is not needed later on when infection is

established. It has also been noted that non-flagellated isogenic mutants have lower virulence

rather than the wild strains (Josenhans and Suerbaum, 2002).

Bacterial movement towards either favourable environment or away from unfavourable environ-

ment is known as chemotaxis. This movement is controlled by a response to a concentration gra-

dient of small phosphorylated response regulators, which are responsible for switching the

flagellar motor rotation on or off (Wadhams and Armitage, 2004). Particular trans-membrane

receptors detect the chemotactic signals by responding to methyl accepting chemotaxis proteins

which are known as specific chemical stimuli (Butler and Camilli, 2005). Afterwards, a histidine

kinase, a regulator in response, a coupling protein and sensory adaptation mediating enzymes


17

actively and passively relate the input signal to the flagellar motor (Boin et al., 2004). Chemotaxis

can be an important virulence factor as it might controls bacterial movement towards target host

(Defoirdt, 2014). O'Toole et al. (1996) reported that chemotaxis in V. anguillarum is controlled by

a gene cheR located in the upstream of a transposon insertion flgB and they also observed that

cheR mutant showed non-chemotactic movement as well as lower virulence.

2.4.2 Production of extracellular polysaccharides and biofilm formation

Many pathogenic bacteria have been found to produce extracellular polysaccharides (EPS) which

have important functions in the pathogenesis (Coplin and Cook, 1990; Hayward, 1991). It has

been reported that bacteria secrete EPS around their cell as a capsule or sometimes like loose

slime (Costerton et al., 1981). The major components of EPS are organic fractions of carbohy-

drates, glycoproteins, proteins, extracellular DNA, glycolipids and humic substances (Flemming

et al., 2007; Sakuragi and Kolter, 2007). Capsular polysaccharide creates a dense and high molec-

ular weight layer around the bacterial cells (Chen et al., 2010). This type of polysaccharide cap-

sule facilitates the attachment to host cells and protects the cells from the action of the host’s de-

fense (Batoni et al., 2016; Chen et al., 2010).

Another group of EPS, the exopolysaccharides, form loose slime that surrounds the bacterial cell

and forms a biofilm matrix. Exopolysaccharide consists of linear or branched long molecules,

including polysaccharides, proteins, lipids and nucleic acids (Defoirdt, 2014; Flemming and

Wingender, 2010). Exopolysaccharide forms the immediate environment of the microorganisms

in a biofilm, supporting their growth by providing nutrient access and protecting the cells from

antimicrobials, phagocytes predation, and even from drying (Donlan and Costerton, 2002;

Flemming and Wingender, 2001, 2010). In addition, exopolysaccharide provides the biofilm’s

mechanical stability, forms a cohesive by mediating their adhesion to surfaces, establishes an

interconnecting three-dimensional polymer network, which forms immobilizing biofilm cells

(Flemming and Wingender, 2010; Wang et al., 2013). The biofilm matrix facilitates the bacteria

to form the most successful living form on earth by providing an external digestive and physio-

logical system (Flemming and Wingender, 2001, 2010). EPS molecules in the photosynthetic

communities can act as light transmitters which supply light energy (photons) to the deeper organ-

isms in the microbial mat. All these functions enable microorganisms to behave together like mul-

ticellular organisms (Flemming and Wingender, 2001).

It has been described that specific genes are responsible for the biofilm formation of vibrios (e.g.

exopolysaccharide biosynthesis, flagella, pilli) together with some regulatory processes (e.g.

quorum sensing, c-di-GMP signalling) (Yildiz and Visick, 2009). For example, in V. cholerae, the

VpsR gene is responsible for switching from smooth to rugose phenotype resulting in the produc-

tion of EPS (Rashid et al., 2004). On the other hand, repression of the hapR gene in the same

bacteria results in increased biofilm formation and makes the bacteria more virulent (Hammer and

Bassler, 2003; Zhu et al., 2002). Quorum sensing controls the extracellular DNA (eDNA) produc-

tion by P. aeruginosa, which organizes its biofilm formation (Allesen-Holm et al., 2006; Gloag et

al., 2013).


18

2.4.3 Production of lytic enzymes

The pathogenesis of many bacteria is controlled by different lytic enzymes (Finlay and Falkow,

1997), which cause damage to the host tissues and allowing the bacteria to spread in the host and

to get nutrients from the body (Defoirdt, 2014). The best described lytic enzymes produced by

bacteria are hemolysins, proteases (e.g. caseinases, gelatinases) lipases, phospholipases etc.

2.4.3.1 Hemolysin

The extracellular protein, hemolysin, produced by pathogenic Gram-negative bacteria of the

Vibrionaceae family, is an important virulence factor towards both fish and shellfish (Hirono and

Aoki, 1991; Sun et al., 2007). Hemolysins cause pathogenesis by lysing blood cells, either by

producing phospholipase enzymes to break the cell membrane (Sun et al., 2007) or by acting as a

porin in the plasma membrane (Defoirdt, 2014; López-Hernández et al., 2015). Besides haemolyt-

ic activity, haemolysins also have other activities like enterotoxin; cytotoxin and cardiotoxin

(Baffone et al., 2005; Hiyoshi et al., 2010). Hemolysins of pathogenic bacteria are controlled by

genes such as vhhA and vhhB genes of V. harveyi VIB 645, tlh gene of V. parahaemolyticus,

VvhA gene of V. vulnificus (Hasegawa et al., 2008; Kim et al., 2003; Sun et al., 2007).

2.4.3.2 Proteases

Proteases are lytic enzymes, which are capable to degrade collagen, fibronectin and gelatin related

proteins; thereby considering an important virulence factor of pathogenic bacteria (Defoirdt,

2014; Li et al., 2015). The lytic enzymes related to protease activity, include metalloproteases,

serine proteases, cysteine proteases, collagenases, caseinases and gelatinases (Defoirdt, 2014). It

has been revealed that with the presence of extracellular metalloproteases (EmpA) and mucinase,

the pathogenic vibrios is able to degrade the mucus layer of the host; hence colonize into the tis-

sues, subsequently causing systemic infections in different organs (Frans et al., 2011). Han et al.

(2011) reported that loss of EmpA from V. anguillarum resulted in reduced virulence to their

hosts.

Caseinases are protease related lytic enzymes, which are able to degrade caseine. They can be

screened by skim milk containing agar (Defoirdt, 2014; Natrah et al., 2011a). Several species of

vibrios (e.g. V. harveyi, V. anguillarum, V. alginolyticus, V. parahaemolyticus) have been found

to show caseinase activity, which has been proposed as a virulence factor for fish and shellfish

(Bunpa et al., 2016; Costa et al., 2013). Caseinolytic activity has been recognized as a lethal tox-

icity of bacteria and detected as an active lytic enzyme in V. parahaemolyticus isolated from oys-

ters, Crassostrea rhizophorae (Costa et al., 2013).

Gelatinases are proteolytic enzymes produced by pathogenic bacteria, which have a hydrolysing

capacity towards gelatin, haemoglobin, casein, collagen as well as other peptides (Vergis et al.,

2002). Most of the vibrios isolated from sea water and some human pathogenic bacteria (e.g. En-

terococcus faecalis, Aeromonas) have been found to show gelatinase activity (Bunpa et al., 2016;

Costa et al., 2013). Vanmaele et al. (2015) investigated that pathogenic Harveyi clade vibrios

actively produce gelatinase in vitro. Natrah et al. (2011a) reported that in V. harveyi gelatinase


19

activity is positively regulated by quorum sensing; therefore, an important virulence factor during

infection.

2.4.3.3 Lipase

Lipases are hydrolases that merely act on carboxyl ester bonds of triacylglycerols; usually at the

lipid-water interface and work as biocatalysts (Defoirdt, 2014; Teo et al., 2003). Lipases are con-

sidered as important virulence factors, which hydrolyse triacylglycerols, degrading gastric mucus

or causing mucosal damage by disrupting phospholipid rich layer (Gribbon et al., 1993; Smoot,

1997; Teo et al., 2003). Harveyi clade vibrios were found to produce lipase enzymes as a viru-

lence factor (Vanmaele et al., 2015; Natrah et al., 2011a). Liuxy et al. (1996) reported that lipase

activity of V. harveyi is controlled by 1650 bp vst gene, which encode 549 amino acids (Teo et al.,

2003). In gram positive bacteria, Bacillus cereus lipase activity has been reported as a virulence

factor which is regulated by a toxin repressor, rot (Rutherford and Bassler, 2012).

2.4.3.4 Phospholipase

Phospholipase causes hydrolysis of the host cell membrane phospholipids; and are therefor re-

sponsible for tissue destruction. In addition, phospholipases also modulate signalling pathways by

producing a lipid second messenger, which has a noticeable effect on the host. The most studied,

phospolipase C (PLC), is responsible for the direct disruption of nucleated and erythrocyte cell

membrane. However, PLA, produced by vibrios, also has pathogenic effects on the hosts by

cleaving fatty acids at the sn1 (PLA1) and sn2 (PLA2) positions of phosphotidylcholine (Koo et

al., 2007; Lee et al., 2002). Kang et al. (1998) identified phospholipase gene phl in V. mimicus

which is placed upstream towards the opposite direction in the transcription of hemolysin gene

vmhA. Phospholipase lec gene of V. cholerae lies upstream of the hemolysin gene hlyA also to-

wards the opposite direction (Fiore et al., 1997). In P. aeruginosa an extracellular phospholipase

C gene plcB was found to control the virulence (Barker et al., 2004; Cao et al., 2001). It has been

detected that T2SS is a defensive mechanism of pathogenic vibrios through which they secret

degradative enzymes like phospholipase (Dubert et al., 2016). Li et al. (2013) investigated that

plp gene of V. anguillarum able to lyse fish erythrocytes by encoding a phospholipase (PLA2) and

mutation of this gene does not affect its virulence on rainbow trout. Sun et al. (2007) studied that

in V. harveyi, Ser153 is a specific residue, which controls phospholipase B enzymatic activity and

hemolysin function in turbot. Natrah et al. (2011a) worked with five phospholipase genes of V.

Harveyi (e.g. pl-1, pl-2, pl-3, omplA-1 and omplA-2) and its dependency on quorum sensing to

express their virulence. Gram-positive bacteria (e.g. Bacillus) have the virulence to produce phos-

pholipase to cause diseases in human (Bottone, 2010).

2.4.3.5 Chitinase

Chitinases are enzymes which can degrade chitin (e.g. crustacean chitinous exoskeletons) by hy-

drolysing glycosidic bonds and help the pathogens to invade into host tissues (García-Fraga et al.,

2015; Defoirdt, 2014). Chitinase activity of bacteria has been known by the genera of Aeromonus,

Serratia, Vibrio, Streptomyces and Bacillus (Saima and Roohi, 2013). Those bacteria cause char-


20

acteristic lesions by degrading carapace of crabs (Wang, 2011), exoskeleton of zooplanktons and

shellfishes (e.g. shrimp) (Strom and Paranjpye, 2000; Setia and Suharjono, 2015).

2.4.4 Iron acquisition and siderophores

Iron is an important nutrient for several biological processes of animals, including DNA replica-

tion, growth of animals, defence against oxidative stress, methanogenesis etc. (Sandy and Butler,

2009). Pathogenic bacteria require iron within their hosts for their replication and to cause disease

(Skaar, 2010). However, the bioavailability of iron for bacteria is limited inside the hosts, since it

remains sequestered intracellular within ferritin (an iron storage protein) or complexed with heme

(Defoirdt, 2014). In addition, aerobic condition and the neutral pH of serum make extracellular

iron insoluble. Therefore, invading pathogens face difficulties to access the iron. Transferring, a

serum-iron binding protein enhances this difficulty (Defoirdt, 2014; Skaar, 2010; Sandy and But-

ler, 2009). To avoid this difficulty, many pathogens have evolved some iron up taking mecha-

nisms within the vertebrates; including siderophores-mediated iron uptake by secreting low mo-

lecular weight iron binding compounds (Defoirdt, 2014; Sandy and Butler, 2009). This system

enables bacteria to compete with iron sequestration by the hosts using a receptor at the bacterial

surface binds the siderophores-iron complex, internalizes into the cell and released into the cyto-

plasm (Defoirdt, 2014; Skaar, 2010). In addition, they also evolved with a sensing system, Fur

(iron-dependent repressor ferric uptake regulator) which binds to the promoter region of iron-

regulated genes and regulates the expression of the genes in the presence of iron (Defoirdt, 2014;

Finlay and Falkow, 1997; Skaar, 2010). Eventually, siderophores have been determined as viru-

lence factors in some bacteria (e.g. Edwardsiella tarda, V. harveyi, V. parahaemolyticus, V.

anguillarum and V. cholerae) (Amaro et al., 1990; Andrus et al., 1983; Chakraborty et al., 2011;

Owens et al., 1996; Pybus et al., 1994).

2.5 Quorum sensing, bacterial cell-to-cell communication

2.5.1 General overview

Quorum sensing is a bacterial cell-to-cell communication which is developed by the mechanism

of gene regulation in which bacteria regulate the expression of certain genes by producing or in

response to small signal molecules (e.g. autonducers) (Defoirdt, 2014; Ng and Bassler, 2009).

Bacteria synthesize autoinducers (AIs) intracellularly and release it actively or passively to the

outside environment, and the molecules are accumulated extracellularly with the increased bacte-

rial population density. When signal molecules concentration increases to the threshold level, the

AIs are recognized by specific receptors and a signal transduction is triggered, resulting in the

expression or repression of different target genes which brings a collective behavioural adaptation

(Atkinson and Williams, 2009; Ng and Bassler, 2009).

This mechanism was first discovered by Nealson et al. (1970) in marine bacterium V. fischeri.

Afterwards, several gram negative bacteria (e.g. Vibrio spp., Aeromonas spp.) have been found to

produce quorum sensing signalling molecules during infection of their hosts (Bi et al., 2007;

Defoirdt, 2014; Janda and Abbott, 2010). Acylated homoserine lactones (AHLs) have been noted


21

as the most common AI molecules used by Gram-negative bacteria (Defoirdt, 2014; Galloway et

al., 2011).

It has been reported that AHLs share conserved homoserine lactone rings (at β and γ positions),

that are N-acylated with 4 to 14 carbons at the R-position (derived from fatty acid biosynthesis)

which can have a substitution (oxo or hydroxyl) at C3 position (Chhabra et al., 2005; Fuqua and

Greenberg, 2002) (Figure 2.11 a, b, c, d). AHLs signals are produced by LuxI enzyme homo-

logues, that are diffused freely to the outside of the cells and signals are detected by LuxR homo-

logue(s) (Natrah et al., 2011b).

In LuxI/LuxR system, LuxR is highly specific to the signal molecules produced by LuxI, which

after detecting signal molecules leads to the conformational changes of regulatory protein that

allows it to bind to the promoter region of the target genes (Figure 2.10). Due to the specificity of

the LuxI/LuxR enzymes, it has been reported as an intra-species communication system and cer-

tain bacteria might have different AHLs for multiple LuxI/LuxR communication (Natrah et al.,

2011b; Waters and Bassler, 2005).

Figure 2.10: AHL-mediated quorum sensing. The LuxI protein synthase AHL signal molecules.

Signal molecules diffuse outside of cells through plasma membrane; this concentration increases

subsequently with the population density of bacteria. While concentration reached to a threshold

level, AHL binds to a response regulator LuxR, which then regulates the activation or inactivation of

target genes (adapted from the PhD thesis of Defoirdt, 2007).

2.5.2 Multichannel quorum sensing systems in vibrios

In vibrios a distinct multi-channel quorum sensing system has been found, in which they use dif-

ferent types of signal molecules (Milton, 2006) and in some species together with AHL system

(e.g. Vibrio anguillarum) (Defoirdt, 2014). The multi-channel quorum sensing system has been

intensively studied in Vibrio campbellii (formerly called Vibrio harveyi) (Defoirdt, 2014; Lin et

al., 2010) and Vibrio anguillarum (Milton, 2006). Three types of quorum sensing signal mole-

cules, HAI-1 (harveyi autoinducer 1), AI-2 (autoinducer 2) and CAI-1 (cholerae autoinducer 1)

have been found to be produced, detected and responded by Vibrio campbellii (Ng and Bassler,

2009).


22

HAI-1 (a classical AHL signal molecule) is synthesized by LuxM protein, which has so far only

been found in V. campbellii and closely related V. parahaemolyticus (Bassler et al., 1997; Cao

and Meighen, 1989; Qian, 2015) (Figure 2.11 a, b). AI-2 (a furanosyl borate diester) is synthe-

sized by LuxS enzyme (Chen et al., 2002) and the third autoinducer, the CAI-1 ((Z)-3-

aminoundec-2-en-4-one) molecule is synthesized by CqsA enzyme (Ng and Bassler, 2009) (Fig-

ure 2.11 d, e).

In V. campbellii, the AIs are detected by a membrane bound two-component receptor proteins

(autophophorylating histidine sensor kinase protein), which feed a shared phosphorylation/

dephosphorylation signal transduction cascade (Qian, 2015) (Figure 2.12) that controls activa-

tion/ deactivation of LuxRVh (the transcription regulator protein) (Defoirdt, 2014; Ng and Bassler,

2009).

The HAI-1 signal molecule is detected by LuxN, CqsS detects CAI-1 and AI-2 is responded by

LuxQ which is supported by LuxP (a periplasmic binding protein) (Ng and Bassler, 2009). It has

been detected that the quorum sensing circuit of V. campbellii is coordinated by two master regu-

lators, LuxRVh and AphA, in which LuxRVh is considered as the principal quorum sensing regula-

tor. Rutherford et al. (2011) reported that this master regulator (LuxRVh) is able to control gene

expression in both high and low cell densities; whereas, AphA only controls gene expression at

low cell densities (in the absence of the AIs). The phosphorylation status of LuxO determines the

concentration of sRNAs (five small regulatory RNAs) that controls the concentration of LuxRVh

and AphA (Ng and Bassler, 2009).

The net results of kinase and phosphatase proteins together with three AIs (depends on the con-

centration) determine the phosphorylation status of LuxO. AphA is inversely related to LuxRVh.

One of the master regulators individually controls some genes, other genes are controlled by both

of them; hence, together they control hundreds of genes (Defoirdt, 2014). sRNAs can also bind to

mRNA to regulate the genes in another way which enhances translation; therefore, by this way

genes are negatively regulated by quorum sensing (Defoirdt, 2014; Ng and Bassler, 2009). How-

ever, quorum sensing gene regulations strongly depend on the appropriate environmental condi-

tion as well as type of hosts to express particular quorum sensing regulated genes (Defoirdt, 2014;

Pande et al., 2013).


23

Figure 2.11: Chemical structure of quorum sensing signal molecules a) N-(3-hydroxybutanoyl) -L-

homoserine lactone, HAI-1 produced by Vibrio campbellii, b) AHL, homoserine lactone autoinducers

produced by different Gram-negative bacteria, c) N-butanoyl-L-homoserine lactone, signal molecule

produced by Aeromonas hydrophila and Aeromonas salmonicida, d) N-tetradecanoyl-L-homoserine

lactone, an antagonistic long chain AHL molecule towards quorum sensing in Aeromonas hydrophila

and Aeromonas salmonicida, e) Autoinducer-2 (AI-2) of Vibrio harveyi, furanosyl borate diester 3A-

methyl-5,6-dihydro-furo [2,3-D][1,3,2] dioxaborole-2,2,6,6 A tetraol, f) Cholerae autoinducer 1 (CAI-

1) (e.g. Vibrio campbellii), (Z)-3aminoundec-2-en-4-one (adapted from Defoirdt, 2014; Ng and Bassler,

2009).

Figure 2.12: Quorum sensing system in Vibrio campbellii. The LuxM enzyme synthesizes the

autoinducer HAI-1 which is detected by LuxN (two-component receptor protein) at the cell surface.

Subsequently, LuxS and CqsA enzymes produce the autoinducers AI-2 and CAI-1, which are

detected by LuxQ and CqsS two-component receptor proteins, respectively. Periplasmic protein

LuxP helps LuxQ to detect the AI-2. The receptors being autophosphorylated (transfer phosphate to

LuxO via LuxU) in the absence of autoinducers (left). Phosphorylated LuxO, together with σ54

trigger

production of sRNAs (five small regulatory RNAs), which inhibit translation of LuxRVh (master

regulator) and promote the translation of another master regulator AphA. In contrast, when

autoinducers present at high concentrations (right), the receptor proteins are converted to

phosphatases from kinases, which instigate LuxO to dephosphorylate and make it inactive.

Therefore, it blocks sRNAs formation and AphA translation; but promote translation of LuxRVh.

Either individually or in couple AphA and LuxRVh are transcriptional regulators, which are

responsible for transcription of different target genes (Adapted from Defoirdt, 2014).


24

2.5.3 Impact of quorum sensing on the virulence of vibrios

Quorum sensing regulates a wide variety of bacterial phenotypes and gene products which are

responsible for virulence of aquaculture pathogens (Natrah et al., 2011a, b; Qian, 2015). Defoirdt

(2014) reported that the list of quorum sensing bacteria which are pathogenic to plants, animals

and humans is still increasing; many of them are aquaculture pathogens. A list of quorum sensing

bacteria and their relation to virulence are presented in Table 2.3.

Table 2.3: Quorum sensing system of some pathogenic bacteria and their influence on virulence

factors.

Bacteria Signals Quorum sensing regulated virulence and viru-

lence factors

Reference

Vibrio

anguillarum

HHL, AI-2,

CAI-1

Biofilm formation, empA metalloprotease, serine

and glycin protease, melanin pigment,

haemagglutinin protease, haemolytic activity,

extracellular polysaccharides, lipase, mortality of

rainbow trout

Croxatto et al., 2002; Frans et

al., 2011; Rasch et al., 2004;

Qian, 2015; Milton, 2006

Vibrio cholerae AI-2, CAI-1 Toxin-coregulated pilus, exotoxin, cholera toxin,

biofilm development, protease production,

hemolysin

Atkinson and Williams, 2009;

Ng and Bassler, 2009; Tsou

and Zhu, 2010

Vibrio

campbellii

HAI-1, AI-

2, CAI-1

Bioluminescence, Toxin T1, metalloprotease,

siderophore, chitinase A, phospholipase, type III

secretion, extracellular polysaccharide, biofilm

formation, lethality to artemia and rotifers

Defoirdt, 2014; Manefield et

al., 2000; Mok et al., 2003;

Ng and Bassler, 2009; Natrah

et al., 2012; Ruwandeepika et

al., 2012

Vibrio

alginolyticus

AI-2 Biosynthesis from flagella, protease and polysac-

charide, biofilm formation

Qian, 2015; Rui et al., 2008

HAI-1, 3-hydroxybutanoyl-L-homoserine lactone; AI-2, Autoinducer 2 (furanosyl borate diester 3A-

methyl-5,6-dihydrofuro (2,3-D) (1,3,2) diox-aborole-2,2,6,6A-tetraol); CAI-1, (Z)-3-aminoundec-2-en-4-

one; HHL, N-hexanoyl-L-homoserine lactone

2.6 Antivirulence therapy: inhibition of virulence factor production as

a new strategy to control bacterial disease

Antivirulence therapy is a process to combat infection by inactivating the disease causing mecha-

nisms of pathogenic bacteria without using antibiotics and without killing them (Clatworthy et al.,

2007; Defoirdt, 2014). It is a way of inhibiting bacteria from producing virulence factor(s) by

interfering with the regulation of virulence factors, for instance, disarming the pathogens without

harming commensal bacteria (Defoirdt, 2014). Disruption of quorum sensing is considered as a

new anti-infective strategy to control pathogens from producing virulence factors (Qian, 2015;

Finch et al., 1998).

2.6.1 Quorum sensing disruption

2.6.1.1 Inhibition of signal molecules biosynthesis

To date, it has been reported that blocking quorum sensing signal molecules (e.g. AHL) might be

a promising process to inhibit virulence of bacteria. Parsek et al. (1999) found that analogues of

S-adenosylmethionine (homoserine lactone moiety), holo-ACP and sinefungin are able to block

AHLs production in vitro. Recently, Christensen et al. (2013) identified three inhibitors, which

are able to inhibit signal molecules biosynthesis; however, those inhibitors have not been tested in

vivo.


25

2.6.1.2 Application of quorum sensing antagonists

It has been noted that receptor proteins (e.g. LuxR) have preferential binding on the AHLs pro-

duced by cognate proteins (e.g. LuxI). Therefore, those AHLs are strongly different from cognate

AHL, less active, even inhibitory (Fuqua et al., 2001). Natarah et al. (2012) found that the long

chain AHL (N-tetradecanoyl-L-homoserine lactone) protected the burbot larvae (Lota lota) from

Aeromonus species rather than the short chain AHL (N-butanoyl-L-homoserine lactone). In addi-

tion, several synthetic AHL analogues (e.g. N-sulfonyl homoserine lactones) have also been re-

ported as quorum sensing inhibitors (Janssens et al., 2008).

Halogenated furanones produced either naturally by marine algae or synthetic derivatives, are

intensively studied quorum sensing disrupting molecules which have been found to work in AHLs

as well as multichannel quorum sensing system (Janssens et al., 2008). These molecules interact

with LuxR receptor proteins and also interfere with DNA binding activity of LuxRVh master regu-

lator in the multichannel quorum sensing system (Defoirdt et al., 2007b). However, these mole-

cules are toxic to higher organisms and brominated thiofenones and sulphur analogues have been

reported to function in the similar way by interfering with DNA binding activity (Defoirdt, 2014;

Benneche et al., 2011).

Cinnamaldehyde, a safe, non-toxic flavouring substance has been also reported to perform like an

analougue of brominated furanones (Brackman et al., 2008; Defoirdt, 2014). Recently, it has been

studied that it can reduce the mortality of Macrobrachium rossenbergii and Artemia larvae

against V. campbellii (Brackman et al., 2008; Pande et al., 2013), as well as burbot larvae from

Aeromonas hydrophila and Aeromonas salmonicida (Natarah et al., 2012).

Besides those molecules, quorum sensing inhibiting molecules have been found to be produced by

macro-algae (e.g. families; Rhodomelaceae, Caulerpaceae and Galaxauraceae) (Skindersoe et al.,

2008), micro-algae (Chlamydomonas mutablis, Chlamydomonas reinhardtii, Chlorella fusca,

Chlorella vulgaris) (Teplitski et al., 2004) and some marine bacteria (Symploca hydnoides,

Lyngbya majuscule, Lyngbya majuscule, Halobacillus salinus) (Dobretsov et al., 2010; Teasdale

et al., 2009).

2.6.1.3 Enzymatic inactivation and biodegradation of quorum sensing signal mole-

cules

Bacterial enzymatic activity can also disrupt AHLs (Dong et al., 2007), which is mediated by

AHL lactonases (e.g. Bacillus spp.) (Dong et al., 2002) and AHL acylases enzymes (Fast and

Tipton, 2012). AHL lactonases open the lactone rings and most specific AHL degrading enzymes;

AHL acylases cleave AHLs, subsequently release of homoserine lactone and a fatty acid.

2.6.2 Antivirulence compounds targeting other regulatory mechanisms

Two other molecules have been reported to have other regulatory mechanisms instead of quorum

sensing disruption (Defoirdt, 2014); one (e.g. 4-[N-(1,8-naphthalimide)]-n-butyric Acid) inter-

feres with ToxR regulon in vibrios (Hung et al., 2005) and another one (e.g. N-phenyl-4-

{[(phenylamino)thioxomethyl] amino}-benzenesulfonamide) hinders catecholamine host stress


26

hormones detection by bacteria using QseC receptor (Rasko et al., 2008). Both compounds are

able to protect the expression of virulence factors; however, it is not yet tested against aquaculture

pathogens (Defoirdt, 2014).

2.6.3 Antivirulence compounds targeting one specific virulence factor

It is also possible to block specific virulence factors directly by using toxins, secretion systems or

adhesion factors (e.g. pilicides, block formation of pili (Clatworthy et al., 2007); acylated

salicylaldehyde hydrazones and thiazolidinones, inhibit type III secretion (Baron, 2010)) without

killing bacteria or inhibiting their growth (Defoirdt, 2014).

Chapter three Materials and methods

27

3 Materials and methods

3.1 Preparation of culture media

3.1.1 Luria-Bertani broth (LB35)

LB35 was prepared by dissolving 10 g tryptone (Biokar diagnostic), 5 g of yeast extract (Biokar

diagnostic) and 35 g Instant Ocean in 1 L demineralized water. The solution was homogenized by

stirring, autoclaved at 121 ºC for 20 minutes and stored at room temperature for further use.

3.1.2 Luria-Bertani Agar (LB35 agar)

LB35 agar was prepared by adding Bacteriological Type E agar powder (15 gL-1

) (Biokar diagnos-

tic) to fresh LB35, followed by autoclaving for 20 minutes at 121°C. The plates were poured with

15 ml of LB35 agar and allowed to solidify under laminar flow. After cooling the plates were ei-

ther immediately inoculated with cultured bacteria or stored at room temperature for further use.

3.1.3 Thiosulphate-citrate-bile salts-sucrose (TCBS)

TCBS agar was prepared by adding 88.0 g TCBS powder (Biokar diagnostic) in 1 L demineral-

ized water. The mixture was subsequently heated to boiling point using a magnetic stirrer. Plates

were immediately poured (15 ml per plate) under laminar flow and left to solidify for 30 minutes.

The plates were stored in plastic bags at room temperature.

3.2 Preparation of antibiotic stock solutions

The rifampicin (Sigma-Aldrich) stock solution was prepared at 25 gL-1

in dimethylsulfoxyde

(DMSO) (Sigma-Aldrich). Stock solutions of nitrofurazone (50 gL-1

) and chloramphenicol (100

gL-1

) were prepared in dimethylformamide and absolute ethanol respectively. Stock solutions of

kanamycin and ampicilin were prepared in deionized water at 100 gL-1

. All antibiotic stock solu-

tions were vortexed, filter sterilised (0.2 μm) (Whatman™ syringe filter), then wrapped in alumin-

ium foil paper to protect from light degradation and finally stored at - 20 ºC.

3.3 Bacterial strains

Wild type strains of V. crassostreae and V. tasmaniensis and quorum sensing deletion mutants

were used for the in vitro and in vivo tests on blue mussel larvae (M. edulis) (see below in table

3.1).


28

Table 3.1: Overview of Vibrio crassostreae and Vibrio tasmaniensis strains used in this study.

Strains Genotypes or relevant markers or

origin

References

J2-9 Wild strain of V. crassostreae from

where the quorum sensing mutants were

derived.

Lemire et al. (2015)

J2-9ΔLuxM Deletion of luxM (AHL synthase) The mutants of LGP32 have been constructed by dr.

Frédérique Le Roux (Station Biologique de Roscoff,

Roscoff, France)

J2-9ΔLuxS Deletion of luxS (AI-2 synthase)

J2-9ΔCqsA Deletion of cqsA (CAI-1 synthase)

LGP32 Wild type V. splendidus from which the

quorum sensing mutants were derived

Le Roux et al. (2009)

LGP32ΔLuxM Deletion of luxM (AHL synthase) The mutants of J2-9 have been constructed by dr.

Frédérique Le Roux (Station Biologique de Roscoff,

Roscoff, France)

LGP32ΔLuxR Deletion of luxR (quorum sensing mas-

ter regulator)

LGP32ΔLuxS Deletion luxS (AI-2 synthase)

3.4 Storage and culture of bacterial strains

Stock solutions of bacteria were prepared by adding autoclaved glycerol (80 %) to the bacteria

cultures at equal volume, vortexed and subsequently stored at - 80 ºC to use for long time. Before

each experiment, fresh bacteria cultures were prepared by adding 1 % bacterial stock solution

(v/v) to autoclaved LB35 medium and incubated for 24 hours at 18 ºC on a shaker (G 24 Env. Inc.

Shaker, Edison N.J.) under constant agitation.

3.5 Bacterial strain verification

To confirm that all the quorum sensing mutants (Table 3.1) derived from the respective wild

strains J2-9 and LGP32, ERIC-PCR fingerprinting was performed before the start of the experi-

ments.

3.5.1 DNA Extraction

Wild strains (J2-9 and LGP32) and the respective quorum sensing deletion mutants were grown

overnight in LB35. A pure culture of each strain was obtained after streaking on TCBS agar, fol-

lowed by incubation at 18 ºC for 48 hours. A single colony was picked up for each strain in sterile

eppendorfs containing 50 µL PCR water and mixed well by pipetting up and down. After closing

the eppendorfs, it was boiled exactly for 10 minutes and immediately put in ice to cool rapidly

and stored at - 20 ºC for future use.

3.5.2 ERIC-PCR

ERIC-PCR was done by using 10 µM primer work solutions of forward primer, ERIC2F (5’AAG-

TAA-GTG-ACT-GGG-GTG-AGC-G 3’) and reverse primer, ERIC1R (5’ ATG-TAA-GCT-CCT-

GGG-GAT-TCA-C). A PCR master mix (550 µL) was prepared by mixing reagents as described

in Table 3.2. The volume in the PCR tubes were maintained at 25 µL with 24 µL master mix and

1 µL DNA extract. Possible contaminations were checked by 2 negative controls containing 25

µL of master, one was kept open during the whole procedure and the other was closed immediate-

ly.


29

Table 3.2: PCR mastermix used for DNA amplification.

Reagents Volume (µL)

PCR water 425

10x TAQ buffer and KCl-MgCl2 55

25 mM MgCl2 33

10 mM dNTP mix 11

10 µM Primer F 11

10 µM Primer R 11

BSA 1.4

Taq polymerase (5U µL-1) 2.75

Total volume 550.15

The PCR reaction was run in a MyCycler thermal cycler (BIO-RAD). The PCR cycles was initi-

ated with a denaturation at 90 ºC for 05:00 minutes, followed by 35 cycles with the functioning

steps of denaturation at 90 ºC for 00:30 minute, primer annealing at 45 ºC for 01:00 minute, an

elongation step at 72 ºC for 00:10 minute and final elongation at 72 ºC for 20:00 minutes. The

amplified DNA was stored at - 20 ºC.

3.5.3 Gel Electrophoresis

PCR products were diluted 30x (1 µL sample + 29 µL PCR water) in sterile PCR tubes and mixed

well by pipetting. 3 µL of the diluted samples together with 1.5 µL loading buffer were mixed and

run for 105 minutes at 70 V on a 1 % agarose gel stained with gel red and in 0.5x TAE buffer. A 1

kB bp ladder (0.25, 0.5 and 0.75 µL) was loaded on the gel to identify the separated bands of the

targeted DNA samples. After electrophoresis the gel was visualized under a Bio-Rad ChemiDoc

MP gel Imaging System device.

3.5.4 Assessment of Virulence factors

A selection of virulence factors, as described in detail in the following paragraphs, was tested in

vitro. For these tests fresh bacteria cultures were prepared overnight and diluted (using Thermo

Spectronic Genesys 20 spectrophotometer) according to the respective test protocols. All assays

were repeated at least three times under strictly sterile condition and at 18 ºC. All treatments and

replicates of all virulence tests that were statistically compared derived from the same batch of

medium, were inoculated simultaneously and had the same incubation time.

3.5.5 Swimming motility

Swimming motility was tested according to Yang et al. (2014) and Rui et al. (2008) with some

modifications. In short, the plates were prepared by adding 0.2 % (w/v) of Type E Bacteriological

Agar in freshly prepared LB35, followed by stirring and autoclaving. The hot medium was cooled

down to 50 ºC, shaken vigorously and poured into plates (20 mL per plate). The lids were closed

immediately after pouring and left to solidify for 30-60 minutes. Bacterial samples (5 µL) (OD

1.0, 550 nm) were inoculated on the middle of the plates and incubated for max 24 hours. The

motility halo was measured within this time period (in less than 24 hours if the motility zone

tended to reach to the edge of the plate).


30

3.5.6 Lytic enzymes

3.5.6.1 Hemolysin

Hemolysin assay was done according to Natrah et al. (2011a), Liuxy et al. (1996), Zhang and

Austin (2000) and Austin et al. (2005) with some modifications. In short, the plates were prepared

with autoclaved LB35 agar that was supplemented with 5 % (v/v) defibrinated sheep blood (Ther-

mo Scientific, Oxoid Limited). Bacterial samples (2 µL) (OD 0.5, 550 nm) were inoculated in

triplicate on the blood agar plates. The plates were incubated for 4 days, and then the plates were

further incubated at 4 ºC for some days to allow the produced hemolysin to lyse the blood cells.

Subsequently, the colony diameter and clearing zone were measured and the ratio was calculated.

3.5.6.2 Caseinase

Caseinase activity was observed by mixing autoclaved double strength LB35 agar (20 gL-1

tryptone, 10 gL-1

yeast extract, 70 gL-1

instant ocean and 30 gL-1

type E bacteriological agar) with

equal volumes of 4 % (w/v) sterilized (121 ºC for 5 minutes) skimmed milk powder suspension

(Oxoid, Basingstoke, Hampshire, UK). Bacterial samples (2 µL) (OD 0.5, 550 nm) were inoculat-

ed in triplicate on the plates. The plates were incubated for 4 days, then the plates were further

incubated at 4°C for some days to allow the produced caseinase lyse the casein. Subsequently, the

colony diameter and clearing zone were measured and the ratio was calculated after both incuba-

tion periods.

3.5.6.3 Gelatinase

An autoclaved mixture of LB35 agar and 0.5 % gelatin (v/w) (Sigma-Aldrich) was used to assess

the gelatinase activity. Plates were poured (15 mL) and allowed to solidify. Bacterial samples (2

µL) (OD 0.5, 550 nm) were spotted in triplicate on the agar plates. The plates were incubated for

7 days or for 2 days followed by another 5 days at 4 ºC. After incubation the plates were poured

with saturated ammonium sulphate (80 % (w/v)). Colony diameter and clearance zones were

measured after 2 minutes. The ratio was calculated.

3.5.6.4 Lipase

Lipase and phospholipase activity were assayed according to Natrah et al. (2011a), Liuxy et al.

(1996) and Austin et al. (2005) with some modifications. LB35 agar was supplemented with 1 %

Tween80 (Sigma-Aldrich), followed by autoclaving and pouring into the plates. Diluted (OD 0.5,

550 nm) bacterial samples (2 µL) were spotted in triplicate on the plates. The plates were incubat-

ed for 4 days or for 1 day followed by another 3 days at 4 ºC. Colony diameter and/or clearance

zone were measured and the ratio was calculated.


Autoclaved LB35 agar was mixed with 1 % egg yolk emulsion (Sigma-Aldrich) to prepare the

culture plates for phospholipase activity assay. The plates were inoculated in triplicate with dilut-


31

ed (OD 0.5, 550 nm) bacterial culture (2 µL), followed by 4 days incubation. Hence, the colony

diameter and the clearance zone were measured and ratio was calculated.

3.5.7 Biofilm formation

Biofilm formation by the wild strains and quorum sensing mutants were measured according to

Brackman et al. (2008) and Li et al. (2014) with some modifications. The assay was performed in

96 well polystyrene microtiter plates. Overnight bacterial culture (1 % (v/v)) were diluted to

OD550 = 0.5 and inoculated into the wells of a microtiter plate (200 µL per well). Each strain was

measured in 3 independent assays, with 4 replicates each, including a negative control (empty

wells). The plates were then incubated for 24 hours. The wells were washed three times gently

with 300 µL sterile physiological saline to remove all non-adherent bacteria, followed by 30

minutes drying. The remaining adherent bacteria were fixed by adding 200 µL per well of 99 %

methanol. After 2 hours of fixation the plates were emptied and air dried overnight. The plates

were then stained with 200 µL 0.1 % crystal violet for 20 minutes, followed by rinsing under tap

water to remove excess stain and air drying. After complete drying (about 3 hours air drying) the

dye bounded to the adherent cells was resolubilised with 200 µL per well of 95 % ethanol. The

absorbance of each well was measured at 570 nm using Tecan Microplate Reader (Infinite M200).

3.5.8 Exopolysaccharide production

Exopolysaccharide production was assayed in 96-wells microtiter plates (Brackman et al., 2008;

Li et al., 2014) with some modifications. Overnight cultured bacteria (1 % (v/v)) were diluted to

OD550 = 0.5 and inoculated at 200 µL per well in 96 well plates. Each of the bacterial strains was

inoculated as 4 replicates in 3 independent assays together with a negative control. The plates

were then incubated for 24 hours. After incubation, each of the microtiter plates was washed three

times with sterile physiological saline (300 µL per well) to remove all non-adherent bacteria, fol-

lowed by air drying for 30 minutes. After complete drying, the wells were poured with 100 µl per

well of phosphate buffer saline (pH ± 7) containing 5 mM Calcofluor white staining (Sigma Al-

drich) (freshly prepared and light protected). After 60 minutes of incubation with Calcofluor

white staining, fluorescence (excitation 405 nm, emission 500 nm) was measured using Tecan

Microplate Reader (Infinite M200).

3.6 Larval challenge test

3.6.1 Blue mussel D-larvae

Mussel broodstock was collected from culture stocks of the Netherlands by a company named

Roem Van Yerseke (Yerseke, the Netherlands). The broodstock was transported dry and after

arrival in the laboratory the fouling organisms were removed. Hence, they were thoroughly

cleaned by rinsing with filtered seawater (FSW) (0.2 µm) and stored dry at 4 ºC until further use.

For spawning good quality broodstocks were selected and conditioned at 18 ºC with FSW for 20-

30 minutes, followed by thermal shocks of 5 ºC and 20-28 ºC. For better response water flow was

provided. Once gametes were released the males and females were transferred to individually

sterile plastic cups containing filtered autoclaved seawater (FASW) under the laminar flow. Ferti-


32

lisation was allowed by mixing fresh eggs and sperm (max 15 min) at a ratio of 1:10 in a sterile

beaker. Fertilisation rates were checked under a light microscope. When almost 90 % of the eggs

showed polar bodies, the eggs were rinsed on a sterile 30 µm sieve to avoid polyspermy. Another

method used was collecting the freshly released eggs on a sterile 30 µm sieve and placing the

sieve with the eggs for 5-10 seconds in a beaker with freshly collected sperm after which the eggs

were immediately washed with FASW also to avoid polyspermy. In either way, the fertilized eggs

were incubated in sterile 5 liter glass bottles containing FASW and antibiotic mixtures ((chloram-

phenicol, nitrofurazone and enrofloxacin) or (rifampicin, kanamycin and ampicillin)) at the rate of

100, 50 and 10 larvae per mL. The concentration of each of the antibiotics was 10 mgL-1

and all

of the antibiotics were prepared according to the section 3.2. Unfortunately, after many attempts

the fertilised larvae never developed to D-larvae, only misformed trochophora larvae with a high

percentage of mortality. Each attempt the procedure was optimized and all kind of variations were

applied to the original protocol (variation in larval density 10-50-100-200 L mL-1

; variation in

antibiotic (Ab) mixtures) in order to detect the origin of the development failure. Unfortunately

the reason why the larval development was not successful was never revealed. The same protocol

was used successfully for years by different MSc and PhD students, but this year all attempts

failed. Fortunately we could use 2 day old D-larvae from the research institute NIOZ (Yerzeke,

The Netherlands) which we stocked on arrival for 12-48 hours in an Ab mixture (chlorampheni-

col, nitrofurazone, enrofloxacin and rifampicin, kanamycin, ampicillin at the concentration of 10

mgL-1

of each of the antibiotic) at 125 larvae per mL. After incubation with antibiotics they were

gently dough thoroughly washed (with FASW) to eliminate all Ab traces on a sterile 60 µm sieve

and put in a sterile 2 L beaker at a density of 200 ± 30-40 larvae per ml. During stocking in the 24

wells plates for the challenge tests they were distributed homogenously by plunging.

3.6.2 Experimental design

The bacteria strains used for the challenge tests are mentioned in Table 3.1. In total 3 challenge

tests were performed. For each test the larvae were stocked in 24-well plates at an approximate

stocking density of 200 larvae per mL (1 mL per well) with 4 replicates each. All challenge tests

ran for 5 days. Since the larvae needed to be sacrificed in order to count them, a separate plate

was inoculated for each day. To determine the original stocking density as accurately as possible,

an extra well was filled on a separate ‘counting’ plate for each treatment on each plate. The origi-

nal stocking density was then calculated separately for each 24-well plate, minimising the stock-

ing error by calculating the average of these extra wells per plate. Larval survival was determined

daily by counting the live larvae with lugol staining, using a binocular microscope (Euromex,

Holland). The bacterial load of each treatment was checked daily on TCBS agar using a spiral

plater (Spiral System, L.E.D. Techno, nv). All TCBS plates were incubated at 18 ºC for at least 24

hours.


33

3.6.2.1 Challenge test 1

The goal of this test was to visualise the bacterial load of the well content in time during the test.

Therefore, three treatments were chosen: non challenged larvae, to check for contamination;

ME5, a pathogenic Photobacterium sp. and ME6, a pathogenic Splendidus clade Vibrio (Table

3.3, Fgiure 3.1).

Figure 3.1: a) Challenge of mussel larvae with pathogenic vibrios Photobacterium sp. ME5 and Vibrio

sp. ME6 containing 0.1 % LB35 and 10 mg mL-1

rifampicin, b) control for challenge test 1 (containing

0.1 % LB35 and 10 mg mL-1

rifampicin) and 2 (containing 0.1 % LB35), c) larvae after 1 day from

control of challenge test 1.

Table 3.3: In vivo challenge test 1 of blue mussel larvae with bacterial strains ME5 and ME6.

Treatment Larvae

200 ± 30-40 mL-1 Rif

10 mgL-1 LB35

0.1 % (v/v) Inoculum

105 CFU mL-1

Negative control + + + -

ME5 + + + ME5

ME6 + + + ME6

NB: Rif = Rifampicin, (+) = addition, (-) = no addition


In the second challenge test, the survival of mussel larvae was determined after challenge with V.

crassostreae (J2-9 wild type), V. tasmaniensis (LGP32 wild type) and a selection of quorum sens-

ing mutants as described in Table 3.4 and Figure 3.2. More info regarding these strains can be

found in Table 3.1.

Figure 3.2: a) Challenge of mussel larvae with experimental wild strain LGP32 and its quorum

sensing mutants containing 0.1 % LB35, b) wild strain J2-9 and its quorum sensing mutants

containing 0.1 % LB35, c) larvae after 2 days challenge with quorum sensing mutants LGP32ΔluxM.


34

Table 3.4: In vivo challenge test 2 of blue mussel larvae with V. crassostreae, V. tasmaniensis and a

selection of quorum sensing deletion mutants.

Treatment Larvae

200 ± 30-40 mL-1 LB35

0.1 % (v/v) Inoculum

105 CFU mL-1

Negative Control + + -

V. crassostreae J2-9 (wild type) + + J2-9

V. crassostreae J2-9ΔluxM + + J2-9ΔluxM

V. crassostreae J2-9ΔluxS + + J2-9ΔluxS

V. crassostreae J2-9ΔcqsA + + J2-9ΔcqsA

V. tasmaniensis LGP32 (wild type) + + LGP32

V. tasmaniensis LGP32ΔluxM + + LGP32ΔLuxM

V. tasmaniensis LGP32ΔluxR + + LGP32ΔluxR

V. tasmaniensis LGP3ΔluxS + + LGP32ΔluxS

NB: (+) = addition, (-) = no addition


The third challenge test was conducted to determine the effect of cinnamaldehyde at the concen-

trations of 0 µM, 1 µM and 10 µM on the survival of mussel larvae (200 ± 30-40 mL-1

) chal-

lenged with V. crassostreae J2-9, V. tasmaniensis LGP32, Photobacterium sp. ME5 and Vibrio

sp. ME6. For all the treatments, 0.1 % LB35 was added, and the bacteria were inoculated at 105

CFU mL-1

. For ME5 and ME6, 10 mgL-1

rifampicin was added to the water. A negative control

without bacterial inoculum was maintained.

Figure 3.3: a) Challenge of mussel larvae with pathogenic vibrios ME5 and ME6 containing

cinnamaldehyde (0, 1 and 10 µM), 0.1 % LB35 and 10 mg mL-1

rifampicin, b) challenge with

experimental wild strain LGP32 and J2-9 containing cinnamaldehyde (0, 1 and 10 µM) and 0.1 %

LB35, c) larvae after 5 days challenge with ME6 at the cinnamaldehyde concentration 1 µM.

3.7 Statistical analysis

Significance tests between and among the variables were done using t-test and one way anova

with SPSS 23.0 version.

Chapter four Results

35

4 Results

The first part of this master dissertation aimed at investigating the impact of quorum sensing on

virulence factor production of V. crassostreae and V. tasmaniensis in vitro. Before performing

these tests, ERIC-PCR fingerprinting was performed in order to confirm that the fingerprints of

the mutants were identical to those of the respective wild types and thus that the strains were not

contaminated. In the second part, the virulence of the wild type strains, their quorum sensing (QS)

mutants and two other mussel pathogenic isolates (Photobacteriun sp. ME5 and Vibrio sp. ME6)

were tested in vivo towards blue mussel larvae.

4.1 Molecular characterization of wild types and selected quorum sens-

ing mutants

The derivation of the selected quorum sensing mutants from the wild type strains J2-9 and LGP32

was confirmed by ERIC-PCR fingerprinting. The band patterns of the selected QS mutants were

identical to those of their respective wild type strains, which confirmed that they were indeed

derived from the correct wild type strains and that they were not contaminated (Figure 4.1).

Figure 4.1: ERIC-PCR band patterns of V. crassostreae J2-9 and V. tasmaniensis LGP32 wild types

and their selected QS mutants, and of Photobacterium sp. ME5 and Vibrio sp. ME6. NC1: Negative

control 1, which was kept open during and after adding PCR master mix, NC2: Negative control 2,

which was closed immediately after adding the PCR master mix, JC: J2-9ΔcqsA, JS: J2-9ΔluxS, JM:

J2-9ΔluxM, LS: LGP32ΔluxS, LR: LGP32ΔluxR, LM: LGP32ΔluxM.

4.2 Virulence factor production by wild types and quorum sensing mu-

tants

In this study, we determined the production of the (putative) virulence factors swimming motility,

hemolysin, caseinase, gelatinase, lipase, phospholipase, exopolysaccharide production and bio-

film formation in the wild types J2-9 and LGP32 and their selected QS mutants.


36

4.2.1 Swimming motility

Swimming motility was determined on soft agar (conatining 0.2 % agar). After 24 hours of incu-

bation, wild type strains and their QS mutants showed swimming motility on soft agar. The luxM

and luxS deletion mutants showed significantly higher motility compared to wild type LGP32 in

three independent experiments (Table 4.1). In trial 1, the ΔluxR mutant showed significantly low-

er motility than the wild type, but the difference was relatively small and this was not confirmed

in the 2 other experiments. In the case of V. crassostreae, the ΔcqsA showed significantly lower

motility when compared to the wild type J2-9 in the three independent trials. In the 3rd

trial, the

ΔluxS mutant showed significantly higher motility compared to wild type J2-9, but the difference

was relatively small.

Table 4.1: Swimming motility halo diameter (mm) (mean ± standard deviation of three replicates) of

wild types and QS mutants of Vibrio tasmaniensis LGP32 and Vibrio crassostreae J2-9 after 24 hours

of incubation on soft LB35 agar.

Strains Trial 1 Trial 2 Trial 3

LGP32 65 ± 0 64 ± 1 57 ± 2

LGP32ΔluxM 78 ± 3* 80 ± 1* 76 ± 1*

LGP32ΔluxR 61 ± 1* 66 ± 2 61 ± 1

LGP32ΔluxS 76 ± 1* 72 ± 1* 67 ± 1*

J2-9 78 ± 3 70 ± 1 69 ± 0

J2-9ΔluxM 76 ± 1 72 ± 2 69 ± 1

J2-9ΔluxS 79 ± 1 73 ± 3 72 ± 1 *

J2-9ΔcqsA 58 ± 1* 57 ± 2* 52 ± 0*

Mutants that are significantly different from the wild type are marked with an asterisk (independent samples

t-test; p < 0.01).

4.2.2 Production of lytic enzymes

4.2.2.1 Hemolysin

Hemolytic activity was determined on agar containing sheep blood. Wild type V. tasmaniensis

LGP32 and its QS mutants did not show any hemolytic activity on sheep blood agar after 4 days

of incubation at 18 ºC, even after prolongued storage at 4 ºC for some days (Table 4.2). In con-

trast, J2-9 wild type and QS mutants produced hemolysin but there were no significant differences

between the strains.

Table 4.2: Haemolytic activity (mean ± standard deviation of three replicates) of wild types and QS

mutants of Vibrio tasmaniensis and Vibrio crassostreae after 4 days of incubation on LB35 agar with

defibrinated sheep blood.

Trial 1 Trial 2

Strains Colony Clearing zone Ratio Colony Clearing zone Ratio

LGP32 12 ± 0 0 ± 0 0.0 ± 0 10 ± 0 0 ± 0 0.0 ± 0

LGP32ΔluxM 13 ± 0 0 ± 0 0.0 ± 0 11 ± 1 0 ± 0 0.0 ± 0

LGP32ΔluxR 13 ± 1 0 ± 0 0.0 ± 0 12 ± 0 0 ± 0 0.0 ± 0

LGP32ΔluxS 14 ± 0 0 ± 0 0.0 ± 0 11 ± 0 0 ± 0 0.0 ± 0

J2-9 13 ± 0 19 ± 0 1.5 ± 0 12 ± 0 18 ± 0 1.5 ± 0

J2-9ΔluxM 14 ± 0 19 ± 0 1.4 ± 0 12 ± 0 18 ± 0 1.5 ± 0

J2-9ΔluxS 13 ± 0 19 ± 1 1.5 ± 0 11 ± 0 17 ± 0 1.5 ± 0

J2-9ΔcqsA 13 ± 1 18 ± 1 1.4 ± 0 12 ± 0 17 ± 0 1.5 ± 0


t-test; p < 0.01).


37

4.2.2.2 Proteases

Caseinase

Caseinolytic activity was screened using agar containing skimmed milk powder. All strains were

found to show caseinase activity. There were no differences between the quorum sensing mutants

and their respective wild types, except for the luxS deletion mutant of LGP32, which was found to

produce significantly higher caseinase activity than wild type LGP32 (in trial 2 and 3) (Table

4.3). It should be noted, however, that although the differences were significant, they were very

small and probably not physiologically relevant.

Table 4.3: Caseinase activity (mean ± standard deviation of three replicates) of wild types and QS


skim milk powder.

Trial 1 Trial 2 Trial 3

Strains Colony Clearance

zone Ratio Colony

Clearance

zone Ratio Colony

Clearance

zone Ratio

LGP32 12 ± 0 15 ± 1 1.3 ± 0 13 ± 1 18 ± 1 1.4 ± 0 12±0 16±0 1.4±0

LGP32ΔluxM 13 ± 1 18 ± 1 1.4 ± 0 14 ± 1 20 ± 1 1.4 ± 0 14±1 20±1 1.4±0

LGP32ΔluxR 14 ± 0 19 ± 0 1.4 ± 0 15 ± 0 21 ± 0 1.4 ± 0 13±1 19±0 1.5±0

LGP32ΔluxS 14 ± 0 21 ± 1 1.5 ± 0 15 ± 1 23 ± 1 1.6 ± 0* 14±1 22±1 1.6±0*

J2-9 14 ± 1 19 ± 1 1.4 ± 0 15 ± 1 21 ± 1 1.4 ± 0 16 ± 1 23 ± 1 1.4 ± 0

J2-9ΔluxM 15 ± 0 22 ± 1 1.4 ± 0 16 ± 0 23 ± 1 1.4 ± 0 17 ± 0 24 ± 0 1.4 ± 0

J2-9ΔluxS 15 ± 1 20 ± 1 1.4 ± 0 16 ± 0 23 ± 1 1.4 ± 0 17 ± 1 23 ± 1 1.4 ± 0

J2-9ΔcqsA 15 ± 1 19 ± 0 1.3 ± 0 16 ± 0 23 ± 1 1.4 ± 0 16 ± 1 22 ± 1 1.3 ± 0


t-test; p < 0.01).

Gelatinase

Gelatinase activity was determined by using agar containing 0.5 % gelatin. All strains were found

to produce gelatinase activity. However, there were no differences between the quorum sensing

mutants and their corresponding wild types (Table 4.4).

Table 4.4: Gelatinase activity (mean ± standard deviation of three replicates) of wild types and QS


0.5 % gelatin.



zone Ratio Colony

Clearance

zone Ratio Colony

Clearance

zone Ratio

LGP32 16 ± 1 41 ± 1 2.6 ± 0 17 ± 1 40 ± 1 2.4 ± 0 16 ± 1 42 ± 0 2.7 ± 0

LGP32ΔluxM 17 ± 0 44 ± 1 2.6 ± 0 17 ± 0 44 ± 1 2.6 ± 0 16 ± 0 45 ± 1 2.8 ± 0

LGP32ΔluxR 16 ± 0 47 ± 1 2.9 ± 0 18 ± 1 46 ± 1 2.6 ± 0 17 ± 0 44 ± 0 2.7 ± 0

LGP32ΔluxS 18 ± 0 45 ± 0 2.5 ± 0 18 ± 0 45 ± 1 2.5 ± 0 16 ± 0 44 ± 0 2.8 ± 0

J2-9 19 ± 0 44 ± 1 2.4 ± 0 20 ± 0 47 ± 2 2.4 ± 0 17 ± 1 45 ± 0 2.7 ± 0

J2-9ΔluxM 18 ± 1 46 ± 1 2.5 ± 0 20 ± 1 48 ± 2 2.4 ± 0 17 ± 0 47 ± 0 2.8 ± 0

J2-9ΔluxS 17 ± 0 44 ± 0 2.6 ± 0 17 ± 1 46 ± 1 2.7 ± 0 15 ± 0 44 ± 1 2.9 ± 0

J2-9ΔcqsA 18 ± 1 46 ± 2 2.5 ± 0 19 ± 1 47 ± 1 2.5 ± 0 16 ± 0 47 ± 0 2.9 ± 0


t-test; p < 0.01).


38


All strains showed phospholipase activity after four days of incubation on agar with 1 % egg yolk

emulsion. However, quorum sensing did not have significant effects on this virulence factor (Ta-

ble 4.5).

Table 4.5: Phospholipase activity (mean ± standard deviation of three replicates) of wild types and

QS mutants of Vibrio tasmaniensis and Vibrio crassostreae after 4 days of incubation on LB35 agar

with 1 % egg yolk emulsion.

Trial 1 Trial 1


zone Ratio Colony Clearance zone Ratio

LGP32 10 ± 0 30 ± 0 3.1 ± 0 10 ± 0 30 ± 1 3.2 ± 0

LGP32ΔluxM 10 ± 0 32 ± 1 3.1 ± 0 11 ± 0 33 ± 0 3.1 ± 0

LGP32ΔluxR 10 ± 1 33 ± 1 3.2 ± 0 10 ± 0 33 ± 1 3.2 ± 0

LGP32ΔluxS 10 ± 0 32 ± 0 3.3 ± 0 11 ± 0 33 ± 1 3.1 ± 0

J2-9 13 ± 0 31 ± 1 2.3 ± 0 14 ± 0 31 ± 0 2.3 ± 0

J2-9ΔluxM 13 ± 0 32 ± 1 2.4 ± 0 13 ± 1 32 ± 1 2.4 ± 0

J2-9ΔluxS 13 ± 0 30 ± 1 2.3 ± 0 14 ± 1 31 ± 1 2.3 ± 0

J2-9ΔcqsA 14 ± 0 31 ± 1 2.3 ± 0 14 ± 0 33 ± 1 2.3 ± 0


t-test; p < 0.01).

4.2.2.4 Lipase

None of the strains showed any lipase activity on agar with 1 % Tween80 (Table 4.6).

Table 4.6: Lipase activity (mean ± standard deviation in three replicates) of wild types and QS

mutants of Vibrio tasmaniensis and Vibrio crassostreae after 4 days of incubation on LB35 agar with 1

% tween80.



zone Ratio Colony

Clearance

zone Ratio Colony

Clearance

zone Ratio

LGP32 12 ± 1 0 ± 0 0.0 ± 0 14 ± 0 0 ± 0 0.0 ± 0 11 ± 0 0 ± 0 0.0 ± 0

LGP32ΔluxM 15 ± 1 0 ± 0 0.0 ± 0 16 ± 1 0 ± 0 0.0 ± 0 12 ± 1 0 ± 0 0.0 ± 0

LGP32ΔluxR 13 ± 1 0 ± 0 0.0 ± 0 17 ± 0 0 ± 0 0.0 ± 0 12 ± 1 0 ± 0 0.0 ± 0

LGP32ΔluxS 13 ± 0 0 ± 0 0.0 ± 0 17 ± 0 0 ± 0 0.0 ± 0 12 ± 1 0 ± 0 0.0 ± 0

J2-9 16 ± 2 0 ± 0 0.0 ± 0 18 ± 1 0 ± 0 0.0 ± 0 19 ± 1 0 ± 0 0.0 ± 0

J2-9ΔluxM 21 ± 1 0 ± 0 0.0 ± 0 19 ± 2 0 ± 0 0.0 ± 0 15 ± 1 0 ± 0 0.0 ± 0

J2-9ΔluxS 21 ± 1 0 ± 0 0.0 ± 0 19 ± 2 0 ± 0 0.0 ± 0 15 ± 1 0 ± 0 0.0 ± 0

J2-9ΔcqsA 20 ± 2 0 ± 0 0.0 ± 0 19 ± 1 0 ± 0 0.0 ± 0 13 ± 0 0 ± 0 0.0 ± 0

4.2.3 Biofilm formation

All strains were found to produce biofilms in 96 well polystyrene microtiter plates. In case of V.

crassostreae J2-9, there were no differences between the wild type and the mutants, ewcept for

Experiment 2, in which all mutants showed significantly higher biofilm production than the wild

type (Figure 4.2). In the case of V. tasmaniensis, the luxR deletion mutant was found to produce

significantly higher biofilm than the wild type LGP32 in all three experiments (although the dif-

ference was not significant in Experiment 2). In addition, the ΔluxM mutant also showed higher

biofilm production in Experiment 3 (Figure 4.3). It should be noted, however, that although sig-

nificant, the differences were relatively small and might not be physiologically relevant.


39


mutants of Vibrio crassostreae J2-9 (three independent experiments with four replicates in each

experiment). Error bars represent the standard deviation. Mutants that are significantly different

from the wild type are marked with asterisks (independent samples t-test; *: p < 0.05, **: p < 0.01).

0

0.04

0.08

0.12

0.16

0.2

Bio

film

fo

rma

tio

n (

OD

57

0)

Experiment 1

** ** **

0

0.04

0.08

0.12

0.16

0.2

Bio

film

fo

rma

tio

n (

OD

57

0)

Experiment 2

0

0.04

0.08

0.12

0.16

0.2

Bio

film

fo

rma

tio

n (

OD

57

0)

Experiment 3

*

0

0.04

0.08

0.12

0.16

0.2

Bio

film

fo

rm

ati

on

(O

D5

70

) Experiemnt 1

0

0.04

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0.12

0.16

0.2

Bio

film

fo

rma

tio

n (

OD

57

0) Experiement 2


40


mutants of Vibrio tasmaniensis LGP32 (three independent experiments with four replications in each

experiment). Error bars represent the standard deviation. Mutants that are significantly different

from the wild type are marked with asterisks (independent samples t-test; *: p < 0.05, **: p < 0.01).

4.2.4 Exopolysaccharide production

Exopolysaccharide production was assessed using biofilms from 96-well polystyrene microtiter

plates. All strains were found to produce exopolysaccharides (Figure 4.4 and 4.5). However,

there were no significant differences between the quorum sensing mutants and their respective

wild types. The experiment was repeated three times and we found the same results.

Figure 4.4: Production of exopolysaccharides by wild type V. crassostreae J2-9 and its QS mutants.

Error bars represent the standard deviation of four replicates. Astertisks indicate a significant

**

**

0

0.04

0.08

0.12

0.16

0.2 B

iofi

lm f

orm

ati

on

(O

D5

70

)

Experiment 3

0

5000

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25000

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35000

Exo

po

lysa

cch

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Experiment 1

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41

difference in exopolysaccharide production when compared to the wild type; *: p < 0.05, **: p < 0.01

(independent samples t-test).

Figure 4.5: Production of exopolysaccharides by wild type V. tasmaniensis LGP32 and its QS

mutants. Error bars represent the standard deviation of four replicates. Astertisks indicate a

significant difference in exopolysaccharide production when compared to the wild type; *: p < 0.05,

**: p < 0.01 (independent samples t-test).

4.3 Blue mussel larval challenge tests

4.3.1 Challenge test 1: mussel larvae challenged with pathogenic isolates

Photobacterium sp. ME5 and Vibrio sp. ME6

In the first challenge test, the mussel larvae were challenged with ME5, a pathogenic

Photobacterium sp. strain and ME6, a pathogenic Vibrio sp. strain, that were isolated before from

an induced mass mortality event in blue mussel adults. The results of this five day study revealed

that both isolates caused significant mortality in both independent experiments (Figure 4.6).

Another goal of this challenge test was to monitor the bacterial density in the rearing water during

the five days of challenge by plate-counting on TCBS agar. Photobacterium sp. ME5 showed

faster growth than ME6 (Figure 4.7). Indeed, already after one day of incubation, ME5 reached

the maximum density of 106 CFU mL

-1. The density of Vibrio sp. ME6 increased up to day 3, also

reaching 106 CFU mL

-1. In the negative controls, which were not inoculated with bacteria, plate

counts were generally much lower, indicating that the larvae contained relatively low numbers of

Vibrionaceae, but were not completely free of them.

0

5000

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Experiment 1

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Experiment 2

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Experiment 3


42

** *

0%

20%

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60%

80%

100%

120%

NC ME6 ME5

Su

rviv

al

(Da

y 1

)

Challenge test 1 (a)

**

0%

20%

40%

60%

80%

100%

120%

NC ME6 ME5

Su

rviv

al

(Da

y 1

)

Challenge test 1 (b)

**

**

0%

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60%

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NC ME6 ME5

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NC ME6 ME5

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**

0%

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NC ME6 ME5

Su

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(Da

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43

Figure 4.6: Survival of blue mussel larvae during five days of challenge with the mussel pathogenic

isolates Photobacterium sp. ME5 and Vibrio sp. ME6. Two independent experiments were performed

(named “a” and “b”). Error bars indicate the standard deviation of four replicates. Astertisks

indicate a significant difference in survival when compared to the negative control (NC); *: p < 0.05,

**: p < 0.01 (independent samples t-test).

Figure 4.7: Bacterial density in the blue mussel larval rearing water during the five days of challenge

with Photobacterium sp. ME5 and Vibrio sp. ME6. Error bars represent the standard deviation of

four replicates. NC: negative control (no pathogen added).

4.3.2 Challenge test 2: mussel larvae challenged with V. crassostreae J2-9 and

V. tasmaniensis LGP32 and their QS mutants

The second challenge test was conducted to determine the impact of QS on the virulence of V.

crassostreae and V. tasmaniensis by monitoring the survival of mussel larvae challenged with

wild type J2-9, LGP32 and their selected QS mutants. Both V. crassostreae J2-9 and V.

tasmaniensis LGP32 were found to be pathogenic to mussel larvae (Figure 4.8 and 4.9). In case

of V. crassostreae, the ΔluxS mutant caused higher mortality than the wild type in both independ-

ent experiments (Figure 4.8). In the case of V. tasmaniensis, there were no consistent differences

between the wild type and the mutants (Figure 4.9).

* *

0%

20%

40%

60%

80%

100%

120%

NC ME6 ME5

Su

rviv

al

(Da

y 5

)


**

**

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NC ME6 ME5

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)


1.00

10.00

100.00

1000.00

10000.00

100000.00

1000000.00

10000000.00

0 1 2 3 4 5

Lo

g10 (

CF

U m

L-1

)

Days

NC ME6 ME5


44

*

0%

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(Da

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)


# *

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(Da

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)



45

Figure 4.8: Survival of blue mussel larvae during five days of challenge with V. crassostreae J2-9 wild

type and three of selected QS mutants (ΔluxM, ΔluxS and ΔcqsA). Two independent experiments

were performed (named “a” and “b”). The error bars represent the standard deviation of four

replicates. # signs indicate a significant difference in survival when wild type was compared to the

negative control (NC), and asterisks indicate a significant difference in survival when the mutants

were compared to wild type (independent samples t-tests; p < 0.01).

#

*

0%

20%

40%

60%

80%

100%

120%

Su

rviv

al

(Da

y 5

)


# *

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(Da

y 5

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(Da

y 1

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(Da

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* * *

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l (D

ay

2)


#

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(Da

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)



46

Figure 4.9: Survival of blue mussel larvae during five days of challenge with V. tasmaniensis LGP32

wild type and three selected QS mutants (ΔluxM, ΔluxR and ΔluxS). The error bars indicate the

standard deviation of four replicates. Two independent experiments were performed (named “a” and

“b”). # signs indicate a significant difference in survival when wild type was compared to the negative

control (NC), and asterisks indicate a significant difference in survival when the mutants were

compared to the wild type (independent samples t-tests; p < 0.01).

# *

0%

20%

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100%

120% S

urv

iva

l (D

ay

3)


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(Da

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)


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47

4.3.3 Challenge test 3: impact of the quorum sensing inhibitor

cinnamaldehyde on the survival of mussel larvae challenged with V.

crassostreae J2-9, V. tasmaniensis LGP32, Photobacterium sp. ME5 and

Vibrio sp. ME6

The third challenge test was conducted to determine the effect of the quorum sensing inhibitor

cinnamaldehyde at different concentrations (1 µM or 10 µM) on the survival of mussel larvae

challenged with different pathogens. The results revealed that cinnamaldehyde did not have any

significant effect on the survival at the concentrations tested (Figure 4.10).

0%

20%

40%

60%

80%

100%

120%

140%

NC1 J2-9 LGP32

Su

rv

iva

l (D

ay

1)

0 µM 1 µM 10 µM

0%

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NC2 ME5 ME6

Su

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(Da

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)

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0%

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(Da

y 2

)

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)

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NC2 ME5 ME6

Su

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(Da

y 3

) 0 µM 1 µM 10 µM


48

Figure 4.10: Survival of blue mussel larvae challenged with V. crassostreae J2-9, V. tasmaniensis

LGP32, Photobacterium sp. ME5 and Vibrio sp. ME6, with and without cinnamaldehyde (1 µM or 10

µM). The error bars represent the standard deviation of four replicates.

0%

20%

40%

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120%

140%

NC1 J2-9 LGP32

Su

rviv

al

(Da

y 4

)

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10 µM

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(Da

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) 0 µM 1 µM 10 µM

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NC1 J2-9 LGP32

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(Da

y 5

)

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0%

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0 µM

1 µM

10 µM

Chapter five Discussion

49

5 Discussion

V. crassostreae (J2-9) and V. tasmaniensis (LGP32) are two important Splendidus clade vibrios

(Sawabe et al., 2014). Although non-virulent strains of those species can be found, both species

also have been reported to contain strains that are pathogenic to molluscs (Travers et al., 2015).

Quorum sensing can regulate pathogenicity by controlling the production of virulence factors

(Natrah et al., 2011a, b; Qian, 2015). V. crassostreae contains a multi-channel quorum sensing

pathway, including the autoinducer synthases CqsA, LuxM and LuxS (Lemire et al., 2015) and V.

tasmaniensis LGP32 was also found to encode homologues of the three-channel quorum sensing

system as described in V. campbellii/harveyi (Tait et al., 2010). Dr. Frédérique Le Roux has con-

structed the quorum sensing mutants ΔluxM, ΔluxR and ΔluxS from the wild type LGP32, and

mutants ΔcqsA, ΔluxM and ΔluxS from wild type J2-9 (unpublished). This dissertation work was

done with those mutants targeting the research question, whether the quorum sensing has any

impact on the virulence of V. crassostreae and V. tasmaniensis in vitro and in vivo in a challenge

with blue mussel larvae. Before starting the experiment, the purity of the mutants was checked by

ERIC-PCR. Then, various virulence factors of the wild types (J2-9 and LGP32) and quorum sens-

ing mutants were assayed. Further, blue mussel larvae were challenged with the wild types J2-9

and LGP32 and their selected quorum sensing mutants, and finally, the effect of a non-toxic quor-

um sensing inhibitor, cinnamaldehyde, was also assessed.

In this study, we investigated the impact of inactivation of single synthase mutants on the swim-

ming motility of V. crassostreae and V. tasmaniensis. The results revealed that the different mu-

tants regulate swimming motility differently. For V. tasmaniensis LGP32, the ΔluxM and ΔluxS

mutants were found to show significantly higher motility when compared to the wild type (Table

4.1). The results indicate that AHL and AI-2 single synthase mutants increase the motility of V.

tasmaniensis, which suggests that quorum sensing downregulates flagellar motility. Similar re-

sults have been obtained for other Vibrio species, in which quorum sensing downregulates swim-

ming motility (i.e. V. cholerae, V. fischeri, V. alginolyticus (Yang and Defoirdt, 2015), V.

parahaemolyticus (Gode-Potratz and McCarter, 2011; McCarter, 2004)). For example, in V.

fischeri, a mutant derived by deleting the litR gene (a homologue of the QS master regulator)

shows increased motility; therefore, quorum sensing represses flagellar motility (Bjelland et al.,

2012). In contrast, in V. crassostreae J2-9, the CAI-1 synthase deletion mutant (ΔcqsA) showed

significantly lower motility when compared to the wild type, which suggests that quorum sensing

positively regulates flagellar motility in this species (Table 4.1). Similar positive regulation of

motility by quorum sensing was shown for the ΔluxR mutant of V. tasmaniensis, although it was

only in one trial and was not confirmed by other two trials (Table 4.1, trial 1). Yang and Defoirdt

(2015) reported that quorum sensing positively regulates flagellar motility of V. harveyi, where

they found that all three and single synthase mutants and the luxR deletion mutant showed lower

motility when compared to the wild type, and this was also confirmed by reverse transcriptase

qPCR on flagellar genes.

A number of lytic enzymes have been found to be produced by Vibrio species, including

hemolysin, caseinase, gelatinase, cysteine protease, chitinase, phospholipase, lipase, serine prote-


50

ase etc. (Qian, 2015). Hemolysin is one of the most studied lytic enzymes produced by pathogenic

bacteria as their virulence factor. In this study, we found that hemolytic activity against sheep

blood agar did not show any significant differences between the quorum sensing mutants and the

wild strains J2-9 after 4 days incubation (Table 4.2). The result indicates that haemolytic activity

of V. crassostreae is independent of quorum sensing. That means that quorum sensing inhibition

will not have any impact on hemolysin production in this species. In contrast, wild type LGP32

and its mutants did not show hemolytic activity. A similar result was observed by Natrah et al.

(2011a), where they found that hemolysin was independent of quorum sensing in V. harveyi. On

the contrary, Ruwandeepika et al. (2012) reported that in Harveyi clade vibrios quorum sensing

positively regulates hemolysin and thermostable direct hemolysin. Frans et al. (2011) stated that

hpdA gene is responsible for hemolysin by V. anguillarum and V. vulnificus, which is positively

regulated by quorum sensing. In addition, it has been noted that many quorum sensing signal

molecules of gram negative bacteria (e.g. V. anguillarum, V. harveyi, V. cholerae, V. mimicus, V.

vulnificus, A. hydrophila, A. salmonicida, Edwardsiella tarda) directly or indirectly regulate the

hemolytic activity (Kim et al., 2003; Qian, 2015; Tsou and Zhu, 2010).

Proteases (e.g. caseinase, gelatinase) are important lytic enzymes, which have been found to be

produced by several pathogenic vibrios (e.g. V. anguillarum, V. harveyi, V. alginolyticus, V.

mimicus, V. vulnificus) and that is also regulated by quorum sensing in various bacteria (Li et al.,

2015; Mok et al., 2003; Rui et al., 2008; Natrah et al., 2011a). In this thesis, we found that both V.

crassostreae and V. tasmaniensis showed caseinolytic activity. However, no relevant differences

were found between wild types and quorum sensing mutants (Table 4.3). Several authors reported

that caseinase activity is regulated by quorum sensing. For instance, in V. harveyi quorum sensing

positively regulates caseinase production in marine agar with skim milk suspension (Natrah et al.,

2011a). In this study, we found that the caseinase activity of V. crassostreae was independent of

quorum sensing.

The results of gelatinase activity revealed that both V. crassostreae and V. tasmaniensis showed

gelatinase activity (Table 4.4). However, no differences were observed between wild types and

quorum sensing mutants, which indicate that gelatinase activity of V. crassostreae and V.

tasmaniensis is independent of quorum sensing. In contrast, in V. harveyi, quorum sensing posi-

tively regulates the gelatinase activity in marine agar with 0.5 % gelatin (Natrah et al., 2011a).

Quorum sensing has been found to regulate lipase and phospholipase activity of several Gram-

negative bacteria. Natrah et al. (2011a) worked with five phospholipase genes of V. harveyi of

which three of them are negatively regulated by quorum sensing. In Pseudomonas aeruginosa

quorum sensing positively regulates phospholipase activity, where AHL quorum sensing system

controls the expression of plcB gene. The plcB gene regulates the twisting mobility of P.

aeruginosa towards certain phospholipase gradient (Cao et al., 2001; Barker et al., 2004). In this

study, we found that all the strains showed phospholipase activity in type E bacterial agar with 1

% egg emulsion. However, the mutants did not show significant differences when compared to

their respective wild types (Table 4.5). That means that phospholipase activity of V. crassostreae

and V. tasmaniensis is independent of quorum sensing. Unlike phospholipase, neither V.


51

crassostreae nor V. tasmaniensis showed lipase activity in type E bacterial agar with 1 %

Tween80 (Table 4.6).

The impact of quorum sensing on biofilm formation by V. crassostreae and V. tasmaniensis was

tested in 96 well polystyrene microtiter plates. All the strains were found to produce biofilms on

these plates (Figure 4.2, 4.3). Our results indicated that quorum sensing does not have an effect

on biofilm formation in V. crassostreae and V. tasmaniensis, except luxR deletion mutant of V.

tasmaniensis. However, this difference was relatively small and was not confirmed by a higher

exopolysaccharide production. In contrast, AI-2 and CAI-1 signal molecules positively regulated

biofilm development in V. harveyi (Atkinson and Williams, 2009). They also noted that in V.

cholerae, quorum sensing negatively regulates biofilm formation. In addition, Zhu et al. (2002)

investigated that ΔhapR mutant has an enhanced and ΔluxO mutant has an impaired biofilm for-

mation. In P. aeruginosa, the LasI/LasR-RhlI/RhlR quorum sensing circuits produce AHL

autoinducers, which control the release of eDNA that is responsible for the switching motility-

mediated biofilm expansion (Gloag et al., 2013; Allesen-Holm et al., 2006). The quorum sensing

master regulator VanT of V. anguillarum has also been reported to be involved in the regulation

of biofilm formation by controlling the production of extracellular polysaccharides (Croxatto et

al., 2002; Frans et al., 2011). Yildiz and Visick (2009) reported that in V. parahaemolyticus,

OpaR (homologue of LuxR in V. harveyi), positively regulates biofilm formation. They also re-

ported that the LuxR homologues SmcR and LitR also positively regulate biofilm formation in V.

vulnificus and V. fischeri respectively.

Our study of exopolysaccharide production in 96-wells microtiter plates revealed that all strains

produced exopolysaccharides, however there were no significant differences between the quorum

sensing mutants and the respective wild types (Figure 4.4, 4.5). So, it can be noted that

exopolysaccharide production of V. crassostreae and V. tasmaniensis is independent of quorum

sensing. In several bacteria, quorum sensing has been found to regulate exopolysaccharide pro-

duction. For instance, biofilm exopolysaccharide (vps) in V. cholerae is expressed in the absence

of the quorum sensing master regulator HapR (homologous to V. harveyi LuxR), which indicates

that quorum sensing negatively regulates exopolysaccharide production in V. cholerae (Atkinson

and Williams, 2009; Hammer and Bassler, 2003). In addition, a luxO deletion mutant (that is

‘locked’ in the regulatory state mimicking high cell density) of V. cholerae cannot produce

exopolysaccharides, which means that quorum sensing negatively regulates exopolysaccharide

production (Hammer and Bassler, 2003; Yildiz et al., 2001). In vibrios, AphA is a DNA-binding

regulator which is also involved in quorum sensing and it has been shown that aphA mutant of V.

parahaemolyticus produces decreased biofilm exopolysaccharide matrix relative to the wild type

(Wang et al., 2013). Flemming et al. (2007) reported that in P. aeruginosa, quorum sensing con-

trols the release of eDNA, which functioning as an intercellular connector in the biofilm to show

an establishing role for biofilm extracellular matrix. In addition, Las quorum sensing regulates the

biofilm matrix gene (pel gene) of P. aeruginosa, which is responsible for the synthesis of Pel

exopolysaccharide that is the main component of biofilm matrix (Sakuragi and Kolter, 2007).

VanT controls the expression of both vps73 and sat genes in V. anguillarum that are involved in

the repression of exopolysaccharide production (Croxatto et al., 2002; Frans et al., 2011).


52

After characterizing the virulence factors of V. crassostreae, V. tasmaniensis and their quorum

sensing mutants in vitro, we investigated the impact of quorum sensing on the virulence of V.

crassostreae and V. tasmaniensis through a series of in vivo challenge tests with blue mussel lar-

vae. In the first in vivo test, a five day challenge of mussel larvae was done with two pathogenic

vibrios, Photobacterium sp. ME5 and Vibrio sp. ME6 to determine their effect on the survival of

blue mussel larvae and to verify that the challenge protocol was still working well. This challenge

test was repeated twice and both Photobacterium sp. ME5 and Vibrio sp. ME6 were found patho-

genic which caused significant mortalities in five day trials (Figure 4.6). In previous studies in

our laboratory, it was also observed that Photobacterium sp. ME5 and Vibrio sp. ME6 are patho-

genic towards blue mussel larvae (Ali, 2015; Yumo, 2014).

In the second challenge test the impact of quorum sensing on the virulence of V. crassostreae and

V. tasmaniensis was tested in vivo towards blue mussel larvae. In the case of V. crassostreae, the

luxS deletion mutant showed significantly lower survival compared to the wild type (Figure 4.8),

which means that AI-2 autoinducer mediated quorum sensing might downregulates the virulence

of V. crassostreae. In the in vitro tests, we had also found that the ΔluxS mutant showed signifi-

cantly higher motility (Table 4.1). That means that AI-2 signal molecules might repress the viru-

lence of V. crassostreae by reducing flagellar motility. Similar result has been repoted by

Coulthurst et al. (2007) that AHL QS of Citrobacter rodentium has an important role in viru-

lence in the mouse, where they found that croI deletion mutant is hypervirulent than the wild type.

In the case of V. tasmaniensis, quorum sensing was found not to have any effect on the virulence

towards mussel larvae (Figure 4.9).

Cinnamaldehyde interferes with AI-2 based QS in various Vibrio spp. by decreasing the DNA-

binding ability of luxR (Brackman et al., 2008). In the third challenge test, we determined the

effect of the quorum sensing inhibitor cinnamaldehyde on the survival of mussel larvae against

wild type, J2-9 & LGP32 and pathogenic bacteria, ME5 & ME6. Our study revealed that at the

experimental concentrations (1 µM and 10 µM), cinnamaldehyde neither able to protect the mus-

sel larvae against the bacteria nor toxic to the larvae (Figure 4.10). This result indicates that, the

experimental concentrations of cinnamaldehyde are not enough to save the mussel larvae. In con-

trast to this result, cinnamaldehyde (at 100 μM and 150 μM) and some of its substituted deriva-

tives can protect the gnotobiotic Artemia against virulent Vibrio harveyi BB120 (Brackman et al.,

2008). In addition, cinnamaldehyde was found to protect burbot larvae at the concentration of

0.01 mM challenged with A. hydrophyla and A. salmonicida (Natrah et al., 2012).

From the challenge test and in vitro virulence factors assays, the final conclusion can be drawn

that inhibition of the three channel quorum sensing systems cannot be used as a strategy to control

the virulence of V. crassostreae and V. tasmaniensis to blue mussel larvae. In contrast, several

authors described that quorum sensing inhibitions did have impact on the virulence of pathogenic

vibrios. For instance, AHL-degrading bacteria can save the Artemia nauplii from the pathogenic

V. harvayi (Van Cam et al., 2009). In addition, different quorum sensing signal molecules of V.

campbellii have different impacts towards the survival of different hosts (Pande et al., 2013).

Defoirdt et al. (2005) noted that mutation in AI-2 genes can abolish the virulence of V. harveyi

towards gnotobiotic A. franciscana. However, after working with the expression of virulence


53

genes of Harveyi clade vibrios Ruwandeepika et al. (2011) noted that expression level of luxR,

toxR genes negatively regulates the survival of challenged Artemia larvae. It has also been re-

ported that inactivation of AHL synthase and receptor significantly results in a higher survival of

burbot larvae challenged with A. hydrophyla and A. salmonicida (Natrah et al., 2012). Finally, as

the quorum sensing inhibition cannot control the virulence of of V. crassostreae and V.

tasmaniensis, so, the strategy to control the vibriosis caused by these bacteria should be taken by

different approaches. By knowing the exact responsible genes for the virulence we should go for

gene expression modifications and/or multipurpose or multitarget antibiotic alternatives.

Chapter six Conclusions and recommendations

54

6 Conclusions and recommendations

From the obtained results the following conclusions can be drawn:

1. Quorum sensing does not control the virulence of V. crassostreae and V. tasmaniensis

towards blue mussel larvae and therefore, quorum sensing inhibition will not be effective

to control vibriosis caused by these pathogens to molluscs.

2. Quorum sensing was observed to have direct regulatory mechanism towards virulence of

a. V. crassostreae: upregulating swimming motility, other virulence factors are inde-

pendent of quorum sensing

b. V. tasmaniensis: downregulating swimming motility, repressing caseinase activity

(probably not physiologically relevant) and reduced biofilm formation (probably not

physiologically relevant).

3. V. crassostreae and V. tasmaniensis are pathogenic to blue mussel larvae; and a luxS dele-

tion mutant of V. crassostreae is hypervirulent towards blue mussel larvae, which might

be linked to virulence factors tested in vitro (i.e. increased swimming motility).

4. The quorum sensing inhibitor cinnamaldehyde does not have any protective effect at the

experimental concentrations (1 and 10 µM).

Recommentations:

1. Determinging the genome sequence and the major genes responsible for virulence of the-

se bacteria, and further research should be conducted to unravel the mechanism behind

the pathogenicity of V. crassostreae and V. tasmaniensis.

2. Further research is needed to confirm the hypervirulence of luxS deletion in V.

crassostreae.

3. The impact of higher concentrations of cinnamaldehyde (e.g. 100 µM) on the survival of

challenged blue mussel larvae might be tested to further confirm that this compound does

not protect them from V. crassostreae and V. tasmaniensis.

4. If major virulence factors regulating genes are identified, then further research is needed

to modify (either it might be deletion or deactivation/activation) those genes to control the

pathogenicity of V. crassostreae and V. tasmaniensis.

References

55

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