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INVESTIGATIONS ON THE DISTRIBUTION OF PATHOGENIC VIBRIOS IN APALACHICOLA BAY FLORIDA AND THE APPLICATION OF CHITOSAN AS A
POSSIBLE MITIGATION STRATEGY
By
LEI FANG
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
2015
© 2015 Lei Fang
To my mom, dad, and family in China
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ACKNOWLEDGMENTS
First and foremost, I would like to express my immense gratitude to my advisor,
Dr. Anita C. Wright, who has shown me the road and helped me get started on the
journey to my Ph.D. degree. She has inspired me to become an independent
researcher and helped me realize the power of critical thinking. Without her guidance
and persistent help throughout the years, this dissertation would not have been
possible. My sincere thanks must also be given to Dr. Kwang C. Jeong, whose insightful
advice and constructive comments are indispensable to the successful completion of
this work. I am also cordially grateful to my committee members Dr. Keith R. Schneider
and Dr. Soohyoun Ahn for providing valuable career advice and continuous support
during my graduate studies. It’s my greatest honor to have these four prestigious
professors on board. Their guidance helped me in a great deal to march forward
continuously.
I would also like to thank Food Science and Human Nutrition Department at the
University of Florida for the financial support provided through awards and scholarships.
Additionally, assistance of all the helpful individuals from Apalachicola DACS lab and
Dr. Cheryl Whistler’s lab at University of New Hampshire was invaluable. More
importantly, extensive thanks are due to all of the lab mates, technicians, fellow
graduate students and friends for their constant and unconditional help throughout this
project, including Dr. Melissa Jones, Kaipeng Xu, Jessica Lepper, Amber Ginn, Zhiyao
Luo, Evan Johnson, Michael Star, Bernhard Wolmarans, Mike Hubbard, Rick Swain,
Samantha King Cekic, Shuang Wu, Xinyu Zhao, Scott Gereffi, and Ning Gao. They
made my journey towards Ph.D. degree more enjoyable.
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Finally, this dissertation is dedicated to my parents and relatives. None of my
achievements would have been possible without their love, endless support and
encouragement. They are always in my heart.
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TABLE OF CONTENTS page
ACKNOWLEDGMENTS .................................................................................................. 4
LIST OF TABLES ............................................................................................................ 9
LIST OF FIGURES ........................................................................................................ 10
LIST OF ABBREVIATIONS ........................................................................................... 11
ABSTRACT ................................................................................................................... 12
CHAPTER
1 INTRODUCTION .................................................................................................... 14
Literature Review .................................................................................................... 15 Vibrio cholerae ................................................................................................. 15
Vibrio vulnificus ................................................................................................ 22 Vibrio parahaemolyticus ................................................................................... 25
Environmental Distribution of Vibrio spp. in Gulf of Mexico .............................. 28 Monitoring Methods for V. cholerae .................................................................. 30 Molecular Typing Methods for V. cholerae ....................................................... 32
Post-Harvest Processing .................................................................................. 34
Chitosan and Chitosan Microparticles .............................................................. 37 Research Hypotheses: Rationale and Objectives ................................................... 39
Specific aim 1: Examine the Distribution of V. cholerae in Seawater and Oysters from Apalachicola Bay, Florida Relative to Different Environmental Parameters and Levels of Other Pathogenic Vibrios. ............ 40
Specific aim 2: Evaluate the Population Structure and Virulence Potential of V. cholerae from Environmental Sources in the Apalachicola Bay. ............... 41
Specific aim 3: Determine the Anti-Vibrio Potential of Chitosan in Seawater and Oysters. .................................................................................................. 41
2 DISTRIBUTION OF V. CHOLERAE IN SEAWATER AND OYSTERS FROM APALACHICOLA BAY, FLORIDA ........................................................................... 42
Introduction ............................................................................................................. 42
Materials and Methods............................................................................................ 43 Samples Collection and Processing ................................................................. 43 Isolation and Enumeration of Vibrios ................................................................ 44 DNA Extraction and Species Identification ....................................................... 45 Analysis of Abundance with Environmental Conditions .................................... 46
Results .................................................................................................................... 46 Discussion .............................................................................................................. 49
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3 GENETIC CHARACTERIZATION AND VIRULENCE POTENTIAL OF V. CHOLERAE FROM APALACHICOLA BAY ............................................................ 63
Introduction ............................................................................................................. 63
Materials and Methods............................................................................................ 64 Strains and Culture Conditions ......................................................................... 64 PCR Screening for Virulence Genes ................................................................ 64 Phylogenetic Characterization .......................................................................... 64 Antibiotic Susceptibility Test ............................................................................. 65
Results .................................................................................................................... 66 Genetic Characterization of V. cholerae from Apalachicola Bay ...................... 66 Phylogenetic Analysis of V. cholerae Population in Apalachicola Bay ............. 67 Antibiotic Resistance of V. cholerae in Apalachicola Bay ................................. 68
Discussion .............................................................................................................. 68
4 THE ANTIVIBRIOCIDAL POTENTIAL OF CHITOSAN MICROPARTICLE IN SEAWATER AND OYSTERS ................................................................................. 79
Introduction ............................................................................................................. 79
Materials and Methods............................................................................................ 81 Bacterial Strains and Inoculum Preparation ..................................................... 81 Chitosan Microparticles (CM) Preparation ........................................................ 82 In vitro Evaluation of Effects of CM on Growth of Vibrio spp. ........................... 82 Effects of CM Treatment on Survival of V. vulnificus and V.
parahaemolyticus in Artificially Inoculated Oysters ....................................... 83
Effects of CM Treatment on Survival of Indigenous Vibrio spp. in Oysters....... 84
Statistical Analysis ............................................................................................ 85 Results .................................................................................................................... 85
Chitosan Inhibits Growth of Vibrio spp. in Broth Culture ................................... 85
Effects of CM on Survival of Vibrio spp. in ASW .............................................. 86 Effect of CM Treatment on Survival of Vibrio spp. in Artificially Inoculated
Live Oysters .................................................................................................. 87 Effect of CM Treatment on Survival of Indigenous Vibrios in Live Oysters ....... 88
Discussion .............................................................................................................. 89
5 ROLE OF CAPSULAR POLYSACCHARIDE IN THE ACTIVITY OF CHITOSAN FOR VIBRIO VULNIFICUS ................................................................................... 100
Introduction ........................................................................................................... 100 Materials and Methods.......................................................................................... 102
Bacterial Strains and Culture Conditions ........................................................ 102 Effects of Chitosan Microparticles (CM) Treatment on Survival of Individual
V. vulnificus Strains ..................................................................................... 102 Effects of CM Treatment on Competitive Survival of V. vulnificus Strains ...... 102
Phase Variation .............................................................................................. 103 Statistical Analyses ........................................................................................ 103
Results .................................................................................................................. 103
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Strain Variation in the Sensitivity of V. vulnificus to CM Activity ..................... 103
Effects of CM on Survival of V. vulnificus as a Function of CPS Expression. . 104 Effects of CM Treatment on Competitive Survival of V. vulnificus Strains. ..... 104
Effects of CM Treatment on CPS Phase Variation of V. vulnificus Strains. .... 105
Discussion ............................................................................................................ 106
6 SUMMARY AND CONCLUSIONS ........................................................................ 113
LIST OF REFERENCES ............................................................................................. 118
BIOGRAPHICAL SKETCH .......................................................................................... 133
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LIST OF TABLES
Table page 2-1 Sequences of oligonucleotides used for molecular analysis ............................... 53
2-2 Spatial distribution of V. cholerae in water and oysters from Apalachicola Bay. .................................................................................................................... 55
2-3 Spatial distribution of V. vulnificus in water and oysters from Apalachicola Bay. .................................................................................................................... 56
2-4 Spatial distribution of V. parahaemolyticus in water and oysters from Apalachicola Bay. ............................................................................................... 57
2-5 Distribution of Vibrios by general location. .......................................................... 58
2-6 Overall relationship of environmental conditions to prevalence of V. cholerae. .. 60
2-7 Overall relationship of environmental conditions with abundance of V. cholerae. ........................................................................................................ 62
3-1 Summary of V. cholerae strains used in this study ............................................. 73
3-2 Virulence potential of V. cholerae strains isolated from environmental samples in Florida. ............................................................................................. 77
3-3 Antibiotic susceptibility test result ....................................................................... 78
4-1 Effects of CM on indigenous V. vulnificus in oysters .......................................... 97
4-2 Effects of CM on indigenous V. parahaemolyticus in oysters ............................. 98
4-3 Effects of CM on heterotrophic aerobic bacteria in naturally infected oysters. ... 99
5-1 Summary of V. vulnificus strains used in this study. ......................................... 109
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LIST OF FIGURES
Figure page 2-1 Sampling sites in Apalachicola Bay. ................................................................... 52
2-2 Overall distribution of Vibrio spp. in water and oysters.. ..................................... 54
2-3 Relationship of V. cholerae prevalence to various water parameters.. ............... 59
2-4 Relationship of V. cholerae levels to various water parameters.. ....................... 61
3-1 Maximum-likelihood tree for MLST. .................................................................... 71
4-1 Effects of CM on growth of Vibrio spp. in broth culture. ...................................... 94
4-2 Effects of CM on survival of Vibrio spp. in ASW. ................................................ 95
4-3 Effects of CM on survival of Vibrio spp. in artificially inoculated oysters. ............ 96
5-1 Effects of CMs on survival of individual V. vulnificus strains in ASW with CM.. 110
5-2 Competitive survival of V. vulnificus MO6-24/O wild-type strain vs. CPS mutant. ............................................................................................................. 111
5-3 Comparison of phase variation in colony morphology of V. vulnificus strains.. . 112
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LIST OF ABBREVIATIONS
APW Alkaline Peptone Water
CM Chitosan Microparticles
CPS Capsular Polysaccharide
EPS Exopolysaccharide
FDA Food and Drug Administration
GRAS Generally Regarded as Safe
IQF Individual Quick Frozen
mCPC Modified Cellobiose-polymyxin B-colistin
MLST Multilocus Sequence Typing
MPN Most Probable Number
VBNC Viable but Nonculturable
PCR Polymerase Chain Reaction
PFGE Pulsed Field Gel Electrophoresis
PHP Post-Harvest Processing
TCBS Thiosulfate-citrate-bile-sucrose
TCP Toxin-coregulated Pilus
TTSS Type Three Secretion Systems
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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy
INVESTIGATIONS ON THE DISTRIBUTION OF PATHOGENIC VIBRIOS IN
APALACHICOLA BAY FLORIDA AND THE APPLICATION OF CHITOSAN AS A POSSIBLE MITIGATION STRATEGY
By
Lei Fang
August 2015
Chair: Anita C. Wright Major: Food Science
A cholera outbreak in Florida in 2011 was attributed to the consumption of raw
oysters contaminated by toxigenic Vibrio cholerae O75. Little is known about the
ecology of this pathogen in the North Florida Gulf Coast. In this research, levels of
V. cholerae were examined over three years in Apalachicola Bay, Florida. V. cholerae
was found in 48% of seawater samples but was not as widely distributed as V. vulnificus
(89%) and V. parahaemolyticus (83%), which were isolated throughout Apalachicola
Bay. In contrast positive V. cholerae samples were more likely associated with near
shore (71%) compared to off shore (29%) sites. Regression analysis showed inverse
correlation of salinity and conductivity to the abundance/prevalence of V. cholerae in
seawater and oysters, while dissolved oxygen was positively correlated with V. cholerae
in oysters. Multi-locus sequence typing revealed a genetically diverse population of
V. cholerae from seawater, while oyster isolates in Apalachicola Bay were more clonal.
Most strains (88% of strains showed all genes) exhibited close identity (88-100%) to
known virulence genes (toxR, rtxA, hlyA, opmU), but all strains lacked genes necessary
for expression (ctxA, ctxB) and/or acquisition of cholera toxin (tcpA). Antibiotic
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resistance profiles included resistance to kanamycin, streptomycin, amoxicillin,
amikacin, tetracycline, and cephalothin.
Taking a broader perspective, efforts to reduce vibriosis in the United States (US)
have not been entirely successful, as evidenced by recent increases in the incidence of
cases, mostly attributed to V. parahaemolyticus. Effective post-harvest processing
(PHP) to reduce Vibrios in oysters does not address the risk of V. cholerae, and these
procedures are not suitable for the raw “half shell” market, as they also kill the mollusks.
Therefore, chitosan, a non-toxic derivative of chitin, was investigated as an alternative
PHP in live oysters. Chitosan microparticles showed strong anti-Vibrio activity during
growth in broth culture, and significantly reduced survival of these bacteria in seawater
and live oysters. Additionally, a V. vulnificus mutant strain lacking capsular
polysaccharide was more resistant to this activity than the wild-type strain, indicating a
role for this structure in the interactions of chitosan and Vibrios. Overall, this research
provides a foundation for a model to enhance policy and management decisions and
may assist the seafood industry through a novel PHP to prevent or eliminate the risks of
pathogenic Vibrio spp. in Florida oysters.
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CHAPTER 1 INTRODUCTION
Vibrios are gram-negative, rod-shaped bacteria that occur commonly in estuarine
or costal riverine environments. V. cholerae, V. vulnificus, and V. parahaemolyticus are
three principle pathogens in this genus. Globally, V. cholerae is responsible for an
estimated 3 to 5 million diarrheal cases and 100,000 to 120,000 deaths annually, but
the disease is relatively rare in the US (WHO, 2012). Other Vibrio pathogens are
responsible for 75% of seafood-borne bacterial infections (mostly V. parahaemolyticus)
and 95% of associated fatalities (V. vulnificus) in the US (Scallan et al., 2011). Vibrio
infections from these species result in an estimated 80,000 illness, 500 hospitalizations
and 100 deaths annually in the US, which has a great impact on the US seafood
industry (Newton et al., 2012). Consumption of undercooked seafood, especially raw
oysters, can result in a severe, systemic vibriosis. Disease symptoms vary with the
causative species but include mild to severe diarrhea, cramps, fever, nausea, wound
infections, and rapidly fatal septicemia (Dechet et al., 2008; Horseman and Surani,
2011; Jones and Oliver, 2009; Mahmud et al., 2010).
Although cholera in the US is rare, recent outbreaks attributed to Asiatic,
pandemic V. cholerae O1 El tor in South and Central America (Wachsmuth et al., 1993),
Haiti (Weil et al., 2012), and Cuba (Mascarello et al., 2013) highlight the threat to the
western hemisphere. Furthermore, multiple sporadic cases (Tobin-D'Angelo et al.,
2008) and one outbreak (Onifade, 2011) in Florida implicated oysters contaminated with
V. cholerae O75 as the likely source of disease. For this reason, the distribution of
V. cholerae in Apalachicola Bay, the primary site of oyster harvest in Florida, was
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examined in the context of various physical environmental parameters and the
prevalence of the other major pathogenic Vibrio spp.
Efforts to reduce Vibrio infections have also been mostly directed towards
V. vulnificus and V. parahaemolyticus because they pose the most immediate threat to
the US. These efforts have not been entirely successful, as evidenced by recent
increases in the incidence of vibriosis cases (Newton et al., 2012). Furthermore,
potential mitigations aimed at reducing the risks associated with cholera have not been
widely studied. The currently available post-harvest processing (PHP) includes thermal,
irradiation, and high-pressure interventions; however, these also methods kill the
mollusks, rendering them unfit for the live “half shell” market. Therefore, this research
also examined the susceptibility of the three major Vibrio pathogens to chitosan, a novel
non-toxic alternative to traditional PHP, and evaluated its potential application in
seafood industry.
Literature Review
Vibrio cholerae
Vibrio cholerae is the etiological agent of cholera, which has been recorded
among ancient civilizations for 2500 years in the Ganges River Delta (Lacey, 1995). The
modern era of cholera originated with what is often referred to as the first pandemic and
arose from the region of Bengal with an outbreak in 1817, and subsequently ravaged
Middle Eastern, Western European, and other Asian countries (Siddique et al., 1994).
Sporadic cholera continued to devastate nearly the entire world and killed millions
throughout additional three acknowledged pandemics from 1829 to 1879. V. cholerae
was named by Robert Koch, and recognized as the causative agent of contagious
disease in May 1884 during the fifth pandemic (Islam et al., 1993). The fifth (1881–
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1896) and sixth (1899–1923) pandemics were believed to be dominated by serotype O1
biotype classical strain, whereas the current ongoing seventh pandemic is attributed to
serotype O1 biotype El Tor strain (Reidl and Klose, 2002).
Classical and El Tor biotypes were characterized by several phenotypic
properties, such as expression of hemolysis, sensitivity to polymyxin B and specific
phages, Voges-Proskauer phenomenon, and agglutination of chicken erythrocytes. In
1992, the emergence of new serotype O139 was renowned to cause a large outbreak of
cholera in Bangladesh and India, and later disseminated to Southeast Asia and South
and Central America, indicating the ability of non-O1 serotypes to cause cholera
epidemics (Albert et al., 1993; Ramamurthy et al., 1993). Although more than 200
serotypes of V. cholerae have been discovered to date, only serotypes O1 and O139
are associated with major cholera epidemics. V. cholerae O1 and O139 strains produce
cholera toxin, and interestingly the genes of cholera toxin are rarely present in non-
O1/O139 strains (Asakura and Yoshioka, 1994).
Patients infected by cholera usually suffer with severe watery diarrhea and rapid
loss of body fluids, which leads to dehydration and hypotensive shock. Without
treatment, death can occur within hours. The symptoms of cholera are primarily caused
by cholera toxin, which is responsible for the devastating diarrhea. Cholera toxin (CTX)
is a potent A-B type enterotoxin that consists of two subunits: a single enzymatic A
subunit and five identical pore-forming penameric B subunits, encoded by ctxA and ctxB
genes, respectively (Asakura and Yoshioka, 1994). CTX is released from bacteria and
binds specifically to GM1 receptors on enterocytes via B subunits, triggering
endocytosis. The A subunit enzymatically activates a G protein and locks it into its GTP-
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bound form through an ADP-ribosylation reaction. Constitutive G protein activity leads to
the activation of adenylate cyclase and increases intracellular cAMP levels. High cAMP
levels in turn results in the activation of the membrane-bound CFTR protein, giving rise
to the osmotic imbalance. The dramatic efflux of chloride, sodium, and water from the
intestinal epithelium causes massive diarrhea and subsequent dehydration that is the
characteristic of cholera (Asakura and Yoshioka, 1994). V. cholerae also cause wound
infection and septicemia, though these are less frequent and less severe compared to
those caused by V. vulnificus.
Most V. cholerae strains isolated from environment are non-toxigenic and lack
the genes encoding CTX, but horizontal gene transfer and genetic reassortment allow
the emergence of new toxigenic strains through the acquisition of several mobile
genetic elements. The CTX genes reside on a distinct filamentous bacteriophage
(CTXФ), encoding six important virulence genes, including ctxA, ctxB, zot, ace, cep, and
orfU (Waldor and Mekalanos, 1996; Waldor et al., 1997). This phage-mediated transfer
results in the new pathogenic strains as well as clone diversity. The evolution of the
pathogenic potential of V. cholerae can be also achieved by transducing the V. cholerae
pathogenic island (VPI), which contains tcp genes encoding the toxin-coregulated pilus
(TCP), a critical receptor for CTXФ (Faruque et al., 1998). It is noteworthy that CTXФ
has been detected sporadically not only in environmental V. cholerae isolates (non
O1/O139), but also appeared in V. mimicus strains, indicating a fascinating mixture of
genetic elements among different species (Boyd et al., 2000).
A number of other putative virulence factors commonly present in both O1/O139
and non-O1/O139 strains may contribute to pathogenesis. Along with the expression of
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toxins, colonization of the intestine is used by V. cholerae to survive inside a host and
cause disease. The presence of toxin-coregulated pilus (TCP) is essential for the
colonization of V. cholerae in the human intestinal tract. TCP is a type-IV bundle-
forming pilus serving as the receptor for CTXФ, (Waldor and Mekalanos, 1996). RTX
toxin (repeats in toxin) is an exotoxin generally produced by a variety of Gram-negative
bacteria. RtxA exhibits covalent cross-linking activity on cellular actin, causing
depolymerization of actin stress fiber and cytotoxicity in HEp-2 cells. Several genes
encoding RTX (rtxR, rtxA, rtxB) are involved in CTXФ DNA replication and integration
(Butler and Camilli, 2005). V. cholerae also elaborates zonula occludens toxin (Zot) and
accessory cholera enterotoxin (Ace), which decreases tissue resistance and increases
intestinal fluid secretion, respectively (Watnick and Kolter, 1999; Watnick et al., 2001).
The genes coding Zot and Ace are part of the chromosomally integrated genome of
CTXФ that are located immediately upstream of the ctx genes, and have been believed
to play roles in phage packaging and secretion (McLeod et al., 2005).
The LPS serves as a barrier to protect bacteria from avoiding host defense, and
increases survival in the presence of bile and other external stresses (Yildiz and
Schoolnik, 1999). The type of LPS (O1 and O139 vs. non-O1/O139) may also contribute
to virulence (Nesper et al., 2001). The O1 pandemic strains are unencapsulated,
whereas O139 strains have capsular polysaccharide (CPS) associated with the
presence of an O139 LPS, which is considered as a principle factor for the initial cholera
outbreaks invaded by O139 in India and Bangladesh. Both O1 LPS and O139 capsule
are involved in direct mucosal adherence to human hosts, and help in the formation of
biofilm in respect of the survival of organism in the hostile natural environments;
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however, O139 strains appear to be more resistant to complement killing by normal
human serum than non-encapsulated mutants (Morris et al., 1996).
In addition to the virulence factors described above, V. cholerae has developed
several survival strategies to increase persistence in its surrounding environmental
habitat. V. cholerae, along with all other Vibrio species, is chitinolytic, producing
chitinase, and is capable of using chitin as the sole carbon source. Many studies have
demonstrated that the ability of chitin utilization system to increase survival under
starvation conditions (Yildiz and Schoolnik, 1999). Vibrios are often reported at higher
levels in association with chitinous organisms (e.g., copepods, amphipods, algae)
relative to the water column community (Simidu et al., 1971).
Furthermore, V. cholerae forms a three-dimensional biofilm in response to many
adverse environmental conditions. Biofilm-associated cells undergo a phenotypic shift in
behavior by generation of an exopolysaccharide (EPS), reduction of growth rate, and
gene transcription. Specifically, copious EPS expression results in the production of
large three-dimensional biofilm and rugose colony morphology, providing enhanced
chlorine resistance and phage resistance (Morris et al., 1996; Yildiz and Schoolnik,
1999). Surprisingly, strains that are able to build mature biofilm have shown reduced
virulence expression in the infant mouse cholera model, indicating that organisms have
a selective advantage in their ability to enhance persistence of the species at specific
environmental conditions (Watnick and Kolter, 1999; Watnick et al., 2001). In light of
biofilm formation, flagellum, mannose-sensitive hemagglutinin (MSHA), and quorum-
sensing system all apparently play important roles in the attachment process. Flagellum
overcomes the repulsive forces associated with the substratum in the initial bacterial
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attachment specifically of the upper small intestine (Butler and Camilli, 2005). The ability
of O1/O139 serotypes to agglutinate chicken erythrocytes is attributed to the production
of mannose-sensitive hemagglutinin (MSHA). MSHA belongs to the type IV family of pili,
and promotes both adherence to zooplankton and biofilm formation (Chiavelli et al.,
2001).
Quorum sensing is another distinct survival strategy for many bacteria to regulate
genes in response to cell density. Quorum sensing allows V. cholerae to generate
biofilm at low cell density, which facilities intestinal colonization, in contrast that the
repression of biofilm production at high cell density promotes V. cholerae adaptation to
an environmental reservoir. Along with the biofilm formation, successful environmental
persistence can also be achieved by entering a “dormancy” state, referred as viable but
nonculturable (VBNC) state. In this state, V. cholerae cells reduce in size, and cannot
be recovered by routine microbiological media; however, they are still metabolically
active and retain virulence. Strains can enter VBNC state spontaneously to protect
themselves against adverse environmental conditions that are inappropriate for
bacterial growth.
The essential role of water in transmitting V. cholerae has been extensively
studied for more than a century. V. cholerae is clearly one of the very important
inhabitants of the riverine, estuarine, and marine aquatic environment. Waterborne
V. cholerae infection is primarily caused by feces-contaminated drinking water and in
part by transmission between patients. As opposed to V. parahaemolyticus infection,
which causes foodborne illness in both industrialized and developing countries,
V. cholerae infection is a sign of poverty and the result of poor water treatment and
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sanitation. The global burden of cholera is still huge. This causative agent continues to
thrive wherever the water treatment and sanitation facilities are suboptimal. The World
Health Organization (WHO) estimated that there are 3 to 5 million cases of cholera
occurring annually and causing 100,000 to 120,000 deaths worldwide (WHO, 2012).
Following a devastating earthquake, a cholera epidemic started from October 2010 in
Haiti, a country that had been cholera-free for a century and recorded more than
250,000 cases and 4,000 deaths in the first six months (Weil et al., 2012). In 2013,
Mexico reported several cholera cases due to heavy seasonal rainfall. An outbreak in
Cuba was also reported in 2014 (Mascarello et al., 2013).
In the US, sporadic cases of cholera occur most frequently in states bordering
the Gulf Coast, and the ingestion of contaminated seafood is frequently implicated.
However, the largest outbreak (16 cases) was attributed to contaminated rice in 1981
(Morris, 1990). Only 40 domestic cases have been reported to the CDC in US since
1995 from only eight southeastern states between 2003 and 2007 (Tobin-D'Angelo et
al., 2008). These cases are distinguished from pandemic V. cholerae in that they are
not serotype O1 but rather serotype O75, which was recently responsible for a small
outbreak in Florida in 2011 (Onifade et al., 2011). The Florida outbreak was associated
with oyster consumption and aroused concerns around seafood safety (Onifade et al.,
2011). Increasing dependence on imported seafood from endemic areas also pose a
potential public health threat to the US. Changing environmental conditions in Florida
waters and inappropriate food safety practices have been proposed as critical factors
that may have contributed to this outbreak. Anthropogenic impacts along coastal water
may also increase the growth of V. cholerae in Florida waters and facilitate the
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emergence of new pathogenic lineages from other non-O1 and O139 serotypes by
horizontal transfer and recombination. V. cholerae is one of oldest and most recognized
pathogens of humans, providing a valuable paradigm for the connection between
infectious disease and the influence of the surrounding environment.
Vibrio vulnificus
Vibrio vulnificus is found ubiquitously in the water column, sediment, plankton,
shellfish, and some types of fish (Blackwell and Oliver, 2008). It is also a component of
the natural microflora of the Eastern oyster, Crassostrea virginica, in the Gulf of Mexico
and occurs at especially high density in oysters during warm summer months (DePaola
et al., 2009). V. vulnificus is considered one of the most invasive bacterial species, and
causes systemic infections that are responsible for more than 50% mortality rate and
95% seafood-related deaths in US (Feldhusen, 2000). Symptoms range from mild
gastroenteritis to severe septicemia and include fever, vomiting, diarrhea, abdominal
pain, and the formation of secondary lesions on the extremities of patients. In addition,
V. vulnificus is noteworthy in being able to cause wound infections, carrying a 20% to
25% fatality rate when patients expose open wounds or broken skin to contaminated
seawater (Oliver, 2005). People with certain underlying and chronic disease, such as
alcoholism, cirrhosis, hemochromatosis, diabetes, or immune system disorders, have
the greatest possibility to contract serious infections (Blake et al., 1979).
Strains of V. vulnificus can be divided into three distinct biotypes, which differ in
several phenotypic characteristics, including LPS structure, sugar fermentation,
optimum growth temperature, and host specificity (Biosca et al., 1993; Biosca et al.,
1996). Biotype 1 is almost exclusively associated with human infections, while Biotype 2
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strains are more commonly associated with fish disease. Biotype 3 was identified with
human wound infection in Israel and is a genetic hybrid of Biotypes 1 and 2 (Zaidenstein
et al., 2008). Although Biotype 1 is the predominant human pathogen, causing a wide
spectrum of illnesses attributed to different virulence factors, all 3 biotypes are potential
human pathogens.
The capsule polysaccharide (CPS) is the most characterized virulence factor of
V. vulnificus and functions to evade host defense. CPS is present in nearly all strains
and provides resistance to opsonization by complement and subsequent avoidance of
phagocytosis by macrophage (Kashimoto et al., 2003; Tamplin et al., 1985). In addition,
the expression of CPS is able to mask immunogenic structure and retard the clearance
of bacteria in the blood stream, which would facilitates invasion in subcutaneous tissues
(Yoshida et al., 1985). The degree of capsule expression is also related to the relative
virulence of strains in this species, and undergoes phenotypic phase variation between
opaque (encapsulated) and translucent (reduced or acapsular) colony morphologies.
Encapsulated strains with opaque colonies are dramatically more virulent than
acapsular strains that present translucent colony morphology. For example, the opaque
colonies yield <102 in 50% lethal doses (LD50) following mice injection, whereas the
translucent morphotype LD50 is typically >106 bacteria (Wright et al., 1999). Moreover,
encapsulated cells are able to use transferrin-bound iron better than CPS mutants.
The expression of CPS on cell surface involves proteins encoded by three genes
(wza, wzb, wzc) that transport CPS to the cell surface, and these three genes are
genetically similar to E. coli group 1 capsule (Wright et al., 2001a). In recent years, a
rugose phenotype was found responsible for biofilm formation in this species, but its
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role in virulence is still unclear (Grau et al., 2005). Virulence of the bacterium is also
closely linked to the presence of lipopolysaccharide (LPS) that plays a critical role in
pathogenesis by developing endotoxic shock. The ability of V. vulnificus to cause
septicemia, tissue edema, hemorrhage, and significant hypotension is associated with
the production of LPS which is considered as a pyrogen to elicit a cytokine response
and release Tumor Necrosis Factor (Rhee et al., 2005). In animal studies, injection of
purified V. vulnificus LPS to mice results in a sudden decrease of arterial blood pressure
and heart rate, leading to death within one hour (Elmore et al., 1992).
A number of other virulence factors have also been described for V. vulnificus,
including iron acquisition system, elaboration of hemolysin and repeats-in-toxin (RTX),
flagella, and production of various proteins to facilitate attachment. The ferrophilic
characteristics of V. vulnificus demand higher levels of iron than that of other pathogens
for its initial growth. V. vulnificus possesses several iron-scavenging siderophores and
heme receptors mediated by iron-uptake systems that help strains to acquire iron from
transferrin, lactoferrin, siderophores, heme proteins, or directly from human hosts
(Litwin and Byrne, 1998; Simpson and Oliver, 1983, 1987). An intriguing finding
indicated that iron transport is linked to other virulence. The elevated iron levels in
human serum further stimulate the expression of cytolysin-hemolysin (VvhA), which
contributes to strain virulence through hemolytic activity and other cytotoxic effects (Kim
et al., 2009). V. vulnificus cytolysin-hemolysin is capable of forming small pores in
plasma membrane that cause rapid necrosis of soft tissue, cell death, and hypertension
(Kim et al., 1993). However, mutants lacking the gene for VvhA were not impaired for
virulence in a mouse model (Wright and Morris, 1991). At the same time, Vv-RTX
25
hemolysin also appears to be an accessory toxin implicated in cytotoxicity, leading to
membrane integrity losing and cell apoptosis of intestinal epithelium (Lee et al., 2007).
Furthermore, flagella and several proteins, such as outer membrane protein (Goo et al.,
2006), surface lipoprotein (McPherson et al., 1991), pili structural protein (Paranjpye
and Strom, 2005), and prepilin peptidase, are responsible for attachment, motility, and
pathogenesis enhancement (Paranjpye et al., 1998).
Not all strains of V. vulnificus are equally virulent. Strains exhibit significant
genomic heterogeneity, and are separated into two distinct genotypes based on
16S rRNA analysis, named as type A and B (Nilsson et al., 2003), as well as by a
virulence-correlated gene, identified as vcgC (type C) and vcgE (type E) (Nilsson et al.,
2003; Rosche et al., 2005). Genotype also indicates a clear association to clinical or
environmental isolation, with 90% of the strains isolated from clinical samples being
classified as “C-type” and 87% of environmental isolates possessing the vcgE variant.
However, the relevance of genotypic characterization to virulence is not yet fully
understood. A recent study found that both C-type and E-type strains can cause high
levels of skin infection in subcutaneously inoculated mice model, and that some E-types
were capable of causing severe systemic diseases (Thiaville et al., 2011). Therefore,
further research is needed to elucidate the determinants that are necessary for causing
infections in human hosts.
Vibrio parahaemolyticus
V. parahaemolyticus was first identified and described by Fujino et al. (1965) in
Japan following a seafood-borne outbreak with 272 illnesses and 20 deaths in the
1950s. Currently, it is recognized as the most common cause of seafood-borne bacterial
26
illness internationally, including both industrialized and less-developed countries
(Newton et al., 2012). The bacterium is native to coastal, estuarine and marine waters,
occurring in especially high numbers in fish and filter-feeding shellfish. Consumption of
V. parahaemolyticus-contaminated seafood can cause gastroenteritis often
accompanied with abdominal cramping, nausea, vomiting, and fever. The diarrhea is
watery, mucoid, and sometimes bloody with reddish stools. Fatalities caused by
V. parahaemolyticus are extremely rare.
The exact mechanism of V. parahaemolyticus gastroenteritis has been reviewed
extensively but not yet fully understood (DePaola et al., 2003b). Many studies reported
that the diarrheal disease was primarily mediated by toxins, namely thermostable direct
hemolysin (TDH) and TDH-related hemolysin (TRH) (Shirai et al., 1990). TRH shows
approximately 67% homology with TDH based on amino acid sequences, and involves
similar biological activities in the pathogenesis of V. parahaemolyticus (Broberg et al.,
2011). Since most V. parahaemolyticus environmental isolates lack the characteristic
tdh/trh hemolysin genes, the presence of these virulence makers was used to
differentiate the pathogenic from nonpathogenic strains (West et al., 2013). TDH
positive strains are hemolytic on Wagatsuma blood agar, which is known as the
Kanagawa phenomenon (KP), whereas TRH positive strains may or may not produce β
hemolysis on Wagastsuma agar (Joseph et al., 1982; Miyamoto et al., 1969). TDH is a
pore-forming toxin, and causes an influx of ions into certain cells, disrupting signal
transduction and osmotic balance, which results in cell death (Shirai et al., 1990).
Consequently, it is thought to be the cause of the large fluid secretion involved in
production of watery diarrhea in human hosts. However, some animal models have
27
disputed this relationship, whereby deletion mutants lacking TDH or TRH still produced
symptoms (Matsumoto et al., 2000). Furthermore, some human diseases have been
attributed to strains that lack these hemolysins.
Beside TRH and TDH, V. parahaemolyticus possess genes for the type three
secretion system (TTSS), which is also proposed as virulence factors. TTSS is a
needle-like complex that is generally used in several gram negative pathogens to attach
and invade host cells by virulence factor proteins secretion and translocation (Hueck,
1998). Many foodborne pathogens, such as E. coli, Yersinia, Salmonella, and Shigella,
have TTSS to infect eukaryotic cells. There are two sets of TTSS genes (TTSS1 and
TTSS2) encoded in the large and small chromosomes in the clinical strain
V. parahaemolyticus RIMD2210633 (Makino et al., 2003). TTSS1 is detected in all
environmental and clinical isolates, and has a role in the cytotoxic effects of the
organism in HeLa cells (Paranjpye et al., 2012). Conversely, TTSS2 is present
exclusively in clinical V. parahaemolyticus strains, and coincides with TRH and TDH
among a number of strains causing different degrees of inflammatory diarrhea in
humans (Makino et al., 2003).
Different V. parahaemolyticus strains are primarily distinguished according to a
serotyping scheme, which depends on distinct combinations of the somatic (O) and
capsular (K) antigens. Before 1996, V. parahaemolyticus associated with foodborne
cases presented with a variety of serotypes in Japan, India, Thailand, and many other
Asian areas (Elhadi et al., 2004; Deepanjali et al., 2005). At the beginning of February
1996, a unique serotype O3:K6 of V. parahaemolyticus was involved with increased
hospitalizations in Calcutta, India. This new clone showed tdh-positive and trh-negative
28
genotypic characteristics, and rapidly spread in Japan, Taiwan, Bangladesh,
supplanting other serotypes in these given areas (Okuda et al., 1997). Now serotype
O3:K6 is considered as the pandemic strain of V. parahaemolyticus, and has spread
throughout the world (Matsumoto et al., 2000). There are three large outbreaks
documented in the US on the Atlantic, Gulf and Pacific Coasts (Cook et al., 2002a;
Daniels et al., 2000). Unlike most Japanese outbreaks, which implicated fish, US
outbreaks of V. parahaemolyticus gastroenteritis involved primarily oysters, and the
associated strains were divergent from the pandemic O3:K6 strains, indicating an
alteration of the O and K antigens (Chowdhury et al., 2000). The vast majority of
V. parahaemolyticus infections are now attributed to Pacific and Northern Atlantic
coasts, causing an estimated 4500 cases per year in the US (Newton et al., 2012).
Thus, V. parahaemolyticus is an emerging pathogen that has acquired the potential of
causing a global pandemic.
Environmental Distribution of Vibrio spp. in Gulf of Mexico
The northern Gulf of Mexico is a leading resource for fish and shellfish production
in the world, with an estimated contribution of 1.3 billion pounds of seafood valued at
$639 million (EPA, 2010). The Gulf also has eight of the top 20 fishing ports, where an
estimated 2.8 million anglers participate in more than 7 million fishing trips annually
(Tao et al., 2012). Vibrios are temperature-sensitive and tend to be more common in
warm climates that are typical of the northern Gulf of Mexico (Motes et al., 1998). In
warm summer months, almost all oysters harvested in US Gulf Coast carry V. vulnificus
or V. parahaemolyticus, and the total number can exceed 104 organisms/g of oysters
(Cook et al., 2002b). A recent study found that 87% oyster samples were positive for
V. vulnificus and V. parahaemolyticus in the Northern Gulf of Mexico during a 19-month
29
survey period (Johnson et al., 2010). Total Vibrio density in the intestines of some fish
species was reported to be up to 108 CFU/g, greatly exceeding levels in the surrounding
seawater, oysters and sediments (Depaola et al., 1994).
Vibrios undergo a striking seasonal fluctuation in costal estuaries, with
temperature being a primary factor affecting their abundance and numbers. At low
temperatures (10°C) they become non-detectable (Nilsson et al., 1991), perhaps as a
consequence of the loss of culturability upon entering to the VBNC state (Oliver et al.,
1995). The warming of coastal waters is thought to contribute to growth and persistence
of Vibrio spp. and has been postulated as a possible factor that contributes to
increasing outbreaks. Annually, reported vibriosis incidence per 100,000 population
increased from 0.09 to 0.28 in COVIS and from 0.15 to 0.42 in FoodNet in last 15 years
(Newton et al., 2012). Thus, global warming coincides with increases in the pathogenic
potential of environmental reservoirs, as well as disease transmission to more outbreak
incidences (Paz et al., 2007a). Epidemiological studies have revealed an association
between water temperature and V. parahaemolyticus densities in oysters harvested
from the Northern Gulf of Mexico (Zimmerman et al., 2007). A correlation has been also
postulated between V. parahaemolyticus occurrence and the levels of turbidity and
chlorophyll (Lobitz et al., 2000; Watkins and Cabelli, 1985). Salinity is another important
factor notable for demonstrating a strong association with the presence of V. cholerae in
Chesapeake Bay (Louis et al., 2003).
Clearly, the abundance and distribution of these three human pathogens
(V. parahaemolyticus, V. vulnificus and V. cholerae) have been linked to environmental
factors, most notably temperature, depending on the pathogen and its habitat, and the
30
geographic location. River inflow and salinity also serve as useful indicators to predict
the occurrence of V. cholerae in Chesapeake Bay and Great Bay Estuary (Louis et al.,
2003; Schuster et al., 2011). However, the effects of conditions specifically influencing
the prevalence of pathogenic V. cholerae in Gulf Coasts are relatively underreported
relevant to the other Vibrio spp. A better understanding of environmental features based
on physical and biological water quality parameters is critical for the identification of
useful strategies to monitor and forecast the incidence of V. cholerae illnesses in the
US.
Monitoring Methods for V. cholerae
Considering the important public health concerns, a monitoring program to detect
V. cholerae is of utmost importance. Alkaline peptone water (APW) is recommended by
the FDA Bacteriological Analytical Manual (BAM) as an enrichment medium in MPN
analysis of Vibrio concentrations in environmental samples (FDA, 2011). The presence
of high pH and salt concentration are responsible for inhibiting many other
environmental bacteria and enriching the growth of V. cholerae and other pathogenic
Vibrios (DePaola et al., 2003a). The addition of electrolyte supplements such as NaCl,
KCl and MgCl2 may also enhance the growth of Vibrios. Generally, samples are
enriched in APW for 16 to 24 h at 37°C. Selective agars are then employed for the
isolation and purification of presumptive V. cholerae colonies. Thiosulfate-citrate-bile-
sucrose (TCBS) is the most commonly employed medium to isolate V. cholerae and
other enteropathogenic Vibrios from a variety of sources, including clinical,
environmental, or contaminated food. The bile salts provide an alkaline pH and high
salinity to inhibit Gram-positive organisms and suppress coliforms. Unlike other
pathogenic Vibrio spp., such as V. vulnificus and V. parahaemolyticus, V. cholerae is
31
able to ferment sucrose and produces a characteristic yellow colony when plated on this
medium. Although TCBS agar is recommended for isolation of V. cholerae from stool
specimens, it is usually unsatisfactory to achieve isolation and enumeration of this
pathogen from environmental samples. Retrospective studies found that even in areas
of endemicity, it is often difficult to isolate V. cholerae O1 or O139 from the environment,
particularly during inter-epidemic periods (Huq et al., 1990; Khan et al., 1984). Since
TCBS allows for the growth of many false positive typical colonies from the microflora of
enriched food samples, MPN enumeration with this agar is generally not recommended
for V. cholerae by the FDA BAM.
CHROMagar™ Vibrio (Hara-Kudo et al., 2001) is an another widely used
differential medium that clearly distinguishes colonies of V. parahaemolyticus (mauve
color) from other Vibrios which grow as milky white, pale blue or colorless colonies;
however, it is still difficult to distinguish V. cholerae from V. vulnificus, since both
pathogens grow as green blue to turquoise blue colonies on this medium. A recently
available alternative chromogenic agar from Hardy diagnostics is more suitable for
V. cholerae isolation, as the species produces the more distinctive mauve color and can
be easily distinguished from other pathogenic Vibrios on this agar (unpublished data).
Putative V. cholerae isolates presenting typical colony morphology on selective
agars are usually subjected to PCR assays for species identification. Confirmation of
V. cholerae isolates is achieved using species-specific primers. Several studies
demonstrated that ompW and toxR genes can be targeted for the species-specific
identification of pathogenic V. cholerae strains, but require additional PCR targets to
detect the presence of virulence markers (Nandi et al., 2000). In addition, it is
32
worthwhile to mention that ompW sequence is located in smaller chromosome, while
toxR sequence is present in larger chromosome (Chapman et al., 2015; Nandi et al.,
2000). The 16S rRNA sequences are highly conserved among different V. cholerae
biotypes and serotypes, and also used frequently in confirmation of V. cholerae strains
from both clinical and environmental sources (Chun et al., 1999).
Molecular Typing Methods for V. cholerae
Methods to characterize V. cholerae in clinical and environmental samples
increasingly rely upon molecular typing assays. These assays have been applied in
conjunction with other typing methods, such as serotyping, to investigate the virulence
potential of V. cholerae strains. Generally, there are two purposes for molecular typing
of V. cholerae strains: 1) investigate the phylogenetic relationships of environmental
strains to epidemic strains and 2) monitor temporal and geographic distribution of these
strains. In the genus of Vibrio, sequence similarities for the 16S rRNA are more than
97.6% among species and are even greater within a species, making intra-species
typing level very difficult (Chun et al., 1999). As a result, other typing techniques have
been deemed more suitable to discriminate phenotypic and genotypic traits for
V. cholerae. Molecular based typing (DNA fingerprinting) approaches have included
pulsed field gel electrophoresis (PFGE), repetitive element-based PCR (Rep-PCR),
ribotyping, random amplification of polymorphic DNA (RAPD), and multilocus sequence
typing (MLST).
Many studies have applied PFGE for V. cholerae outbreak investigation since
separated fragments are highly discriminatory for epidemiological and phylogenetic
analyses (Bakhshi et al., 2012; Bhuiyan et al., 2012; Reimer et al., 2011). However,
some strains are untypeable by PFGE, and the overall diversity obtained by this method
33
is prohibitively high for epidemiological use (Chowdhury et al., 2000). In MLST studies,
several housekeeping genes and /or virulence genes are usually analyzed. Each
sequence for a given locus is screened for similarity comparisons with known
sequences for gene locus. If the sequence is different, it is considered to be a new allele
and is assigned a unique allele number. Each sequence type is defined by cluster of
similar alleles, and genetic distance is determined by number of accumulated
differences among alleles. MLST provides more coherent phylogenetic comparison than
other typing methods but is laborious and time consuming. MLST using only three
housekeeping genes (recA, pgm, gyrB) and three virulence genes (ctxA, ctxB, tcpA)
was shown to offer superior discriminatory ability for typing compared to PFGE
(Kotetishvili et al., 2003). This study analyzed 22 V. cholerae isolates, including the
epidemic O1 and O139 strains and other serogroups, and authors found greatest
diversity among tcpA and ctxAB genes, providing a better measurement of phylogenetic
relatedness than PFGE.
Whole genome sequencing is rapidly having great impact on pathogen
discrimination and characterization. Whole genome sequencing can be used to derive
more comprehensive MLST but also provides enormous information in single nucleotide
polymorphisms (SNPs) to detect minute differences between different strains. In
retrospect, the completion of first sequencing of human genome spent ten years,
involving thousands of researchers and millions of dollars. Today, with the reduction of
time and cost for sequencing technology, life science research is shifting from gene-
based genotyping to whole genome sequencing. FDA has been utilizing whole genome
sequencing to determine the exact source of food outbreak since 2008. It is now
34
becoming reality for pipelines to fully sequence a whole genome and perform key
interpretation for pathogen typing and identification in one day. For example, genomic
sequencing greatly enhanced epidemiological investigation of cholera outbreak in Haiti
in 2010. Its ability to differentiate close related organisms provides the opportunity to
prevent future outbreaks and offers strong data for government policy decision. With the
continuous declining costs and the development of bioinformatics, the technology of
whole genome sequencing will be used in more laboratories for performing basic
foodborne pathogen identification during foodborne illness outbreaks.
Post-Harvest Processing
Pathogenic Vibrios are clearly autochthonous inhabitants in the coastal and
estuarine environments and are commonly found in oysters. Oysters destined for raw
half-shell consumption are increasingly being exposed to post-harvest processing
methods (PHP) to reduce Vibrios and other bacteria to non-detectable level. Current
PHP technologies used for summer-harvested Gulf oysters include cool pasteurization,
high hydrostatic processing (HHP), irradiation, and individual quick frozen (IQF).
“Cool pasteurization” is a method of killing microorganisms in oysters by
subjecting them to warm water, followed by a rapid cooling and frozen storage. The
process was developed and patented in 1995 by AmeriPure in Franklin, Louisiana. Prior
to heat treatment, oysters are washed first, and then individually banded to stay close.
Banded oysters are further loaded by worker onto trays, and submerged in a 55°C
7,500 gallon water tank for 24 min to obtain an internal temperature of 48-52°C. The
oysters are then transferred into a similar tank containing cool water (2-4°C) for another
15 min to stop the cooking process (Andrews et al., 2000; Cook and Ruple, 1992). Mild
thermal processing is effective in reducing pathogens to non-detectable level but
35
causes unwanted side effects in flavor, appearance, color and taste attributes of oysters
(Cruz-Romero et al., 2007). Currently AmeriPure is the only Gulf oyster processor using
cool pasteurization for oysters (Muth et al., 2013).
High hydrostatic pressure (HHP) is a high pressure treatment of oysters in the
shell that was patented by Motivatit Seafoods, L.L.C. in Houma, Louisiana, in 1999. This
process involves washing, grading, sorting and banding in oysters preparation.
Following treatment, oysters are placed in a stainless steel cylinder, dipped into the
water-filled pressure chamber and 35,000 to 40,000 psi pressure are applied for 3 to 5
min (Manas and Pagan, 2005). This technology attracts a great deal of interest in the
seafood industry because it successfully eliminates pathogenic or spoilage
microorganisms in the oysters but minimally affects their chemical and physical changes
in comparison to thermal processing. Currently, two Louisiana and one Texas Gulf
oyster processors apply HHP to treat summer harvested oysters.
Irradiation involves washing, packaging and labeling oysters, which are then
transported to an irradiation facility. In irradiation processing, oysters are exposed to
either gamma rays, machine generated electrons, or X-rays. Irradiation to remediate of
pathogenic Vibrio spp. in oysters has proven effective with low doses of gamma
irradiation (<1.0 kGy) from Cobalt-60 (Basak, 1996). More recently, X-ray treatment with
1.0 kGy was reported to achieve more than 4.7 log CFU/g reduction of
V. parahaemolyticus in pure culture, half shell and whole shell oysters (Mahmoud,
2009). This technology was also approved by FDA as a post-harvest process, although
it has not yet been applied commercially to Gulf oysters. However, the consumer
36
acceptance and preference towards irradiated molluscan shellfish will warrant further
investigation.
Individual quick frozen (IQF) processing of oysters was initially developed in the
US in 1989 and is now popularized in other countries, including Canada, Australia and
New Zealand (DePaola et al., 2009). This type of processing starts with rinsing and
shucking oysters, followed by a rapid temperature reduction by liquid carbon dioxide or
nitrogen immersion through a freezer tunnel. The next step is to spray the oysters with a
fine mist of water in order to freeze products into a glaze of ice through a glazing
machine. IQF processing is capable of reducing freeze-sensitive pathogens, particularly
V. parahaemolyticus and V. vulnificus. In addition, IQF with extended frozen storage
greatly satisfies market demand to consume raw oyster throughout the year. IQF
processing of oysters is presently used by many oyster operations in the Gulf (two in
Texas, one in Louisiana, one in Mississippi, one in Alabama, and four in Florida).
However, IQF-treated oysters appear to have limited visual aesthetics, which can affect
consumer acceptance. Specifically, freezing oysters alters the texture of natural oyster
meat, making them grainy or poor quality that is not comparable to fresh oysters
(Songsaeng et al., 2010).
Established PHP methods effectively reduce pathogen loads to non-detectable
levels (<30 MPN/g) but fail to maintain the viability of the oysters. Since many
consumers prefer to purchase and consume a living product, these demands encourage
development of new PHP alternatives that can reduce illness in seafood industry and
maintain freshness of seafood at the same time. High salinity treatment is one of the
management strategies used to reduce V. vulnificus in live oysters. This process
37
involves relaying oysters from their point of harvest with moderate salinity and more
abundant V. vulnificus levels to offshore waters with much higher salinity and non-
detectable V. vulnificus levels. High salinity relay (≥30 ppt) resulted in a 2 to 3 log
CFU/g reduction of V. vulnificus levels as compared to two moderate salinity sites (22 to
25 ppt) in the Chesapeake Bay (Audemard et al., 2011). Unfortunately, this technology
results in high oyster morality, and consumer acceptance of a higher-quality product still
needs further evaluation. Depuration is another PHP technology that allows shellfish to
purge environmental contaminants by immersing themselves in tanks of clean
seawater. However, this process showed limited efficiency to remove pathogens to non-
detectable level. The persistence of V. parahaemolyticus and V. vulnificus in tissues of
Gulf Coast oysters was reported in several studies (Chae et al., 2009; Tamplin and
Capers, 1992). Therefore, development of novel PHP alternatives is vital to the seafood
industry for alleviating issues of pathogenic Vibrio spp. in raw oysters.
Another issue with current PHP is that they have focused on V. vulnificus and
V. parahaemolyticus, as they are the primary risk for Vibrio disease from Gulf Coast
oysters and the most prevalent pathogens geographically. No research has evaluated
the effects of oyster PHP on V. cholerae. The potential risk of cholera to the US, as
indicated by recent outbreaks, supports the need to validate PHP that will be effective
for all three pathogenic Vibrios.
Chitosan and Chitosan Microparticles
The primary response to reduce Vibrios in seafood industry relies on the
implementation of postharvest processing (PHP), but current PHP efforts have failed to
prevent increasing rates of vibriosis. Evaluation and development of new PHP methods
for the mitigation of Vibrio spp. is needed. Chitin is the second most abundant polymer
38
in nature and is the most abundant in saline aquatic environments. This polysaccharide
is a major component of the shells from shrimp, crabs, and other crustaceans that are
abundant in ecosystems in rivers, oceans and other estuarine environments. Many
Vibrio spp. are able to adhere to chitin and use chitin as nutrients (Pruzzo et al., 2008).
Chitosan is an aminopolysaccharide biopolymer produced primary from chitin, which is
composed of β-1,4-linked glucosamine (deacetylated units) and N-acetyl-D-glusoamine
(acetylated units). Chitosan was approved as a feed additive in 1983, and has been
accepted as functional food ingredients in health department of Japan in 1992
(Chistoserdova, 2010; Taylor, 2011). To date, chitosan is commercially available as
food additives or dietary supplement on a worldwide scale in Korea, Japan, Finland,
England, and Italy. In the US, chitosan derived from shrimp recently achieved a
generally regard as safe (GRAS) status as a food additive by FDA (FDA, 2012).
The unique chemical structure of chitosan has several biological properties that
have been the focus of scientific research. For example, chitosan-mediated delivery
systems significantly improve the bioavailability of drugs and are categorized either as
nanoparticle, microparticle, or macro delivery systems. Specifically, these microparticles
and nanoparticles were found to have beneficial biological effects including anti-tumor
(Hallaj-Nezhadi et al., 2011), antimicrobial (Schlievert, 2007), cholesterol-reducing
(Hossain et al., 2007), immune system booster (Kim et al., 2006) and free radical
scavenging activity (Cho et al., 2008). In addition to its lack of toxicity and allergenicity,
the biodegradability and biocompatibility of chitosan make it potentially useful for
biomaterial, medical, and pharmaceutical applications.
39
Finally, chitosan has been investigated as an antimicrobial material against a
wide range of target organisms like algae, bacteria, yeasts and fungi in both vivo and
vitro (Chirkov, 2002; Rabea et al., 2003). Early research describing the antimicrobial
potential of chitosan and their derivatives dated from the 1980-1990s. Chitosan
microparticles (CMs) are derived from chitosan with minor cross-linking modification. In
a recent study, CM was first found to successfully reduce E. coli O157:H7 shedding in
cattle as feed additives, which suggested that CM may be applied as a possible
treatment of bacterial infections (Jeong et al., 2011). Chitosan was previously shown to
be effective against V. vulnificus in vitro and in mice (Lee et al., 2009). Unfortunately the
effects of CM against gram-negative species in seafood are still greatly understudied.
The exact mechanism for chitosan activity is not fully understood, and multiple factors
are likely to contribute to this antibacterial action. Chitosan antimicrobial activity is
influenced by various intrinsic (type, molecular weight, viscosity, concentration) and
extrinsic factors (pH, temperature, ionic strength, metal ions, organic matter).
Understanding how these factors interplay of these factors with each other is required in
order to optimize the potency of chitosan preparation for any type of application.
Research Hypotheses: Rationale and Objectives
Pathogenic Vibrio species are the primary risk for seafood safety, particularly for
raw oysters that harbor V. vulnificus and V. parahaemolyticus. The Florida cholera
outbreak was associated with oyster consumption and further aroused concerns around
seafood safety (Onifade et al., 2011). Increasing dependence on imported seafood from
cholera endemic areas also pose a potential public health threat to the U.S. Changing
environmental conditions in Florida waters and inappropriate food safety practices have
been proposed as critical factors that may have contributed to this outbreak. V. cholerae
40
is one of oldest and most recognized pathogens of humans, providing a valuable
paradigm for the connection between infectious disease and the influence of the
surrounding environment. Anthropogenic impacts along coastal water may also
increase the growth of Vibrio spp. in Florida waters and facilitate the emergence of new
pathogenic lineages by horizontal transfer and recombination. Taking a broader
perspective, efforts to reduce vibriosis in the US have not been entirely successful, as
evidenced by recent increases in the incidence of cases, mostly attributed to
V. parahaemolyticus. Effective post-harvest processing (PHP) to reduce Vibrios in
oysters does not address the risk of V. cholerae, and these procedures are not suitable
for the raw “half shell” market, as they also kill the mollusks.
Therefore, the focus of this study was threefold: 1) to investigate the hypothesis
that the association of cholera with Apalachicola Bay Oysters was related to the
distribution and virulence potential of V. cholerae in Apalachicola Bay, 2) to provide data
for the hypothesis that relative abundance of Vibrios in Apalachicola Bay was influenced
by environmental conditions, and 3) to evaluate the hypothesis that the antimicrobial
properties of chitosan could have application to the mitigation of Vibrios in seawater and
oysters as a potential PHP for the seafood industry.
The objective of this study includes the following specific aims:
Specific aim 1: Examine the Distribution of V. cholerae in Seawater and Oysters from Apalachicola Bay, Florida Relative to Different Environmental Parameters and Levels of Other Pathogenic Vibrios.
The distribution of V. cholerae was surveyed under different ecological conditions
at different sites of Apalachicola Bay during various seasons over three years. In
particular, the occurrence and abundance of V. cholerae was investigated with respect
to other pathogenic Vibrios, as well as to salinity, conductivity, dissolved oxygen,
41
temperature, and pH in order to understand how environmental dynamics affect the
ecology of V. cholerae. These results are described in Chapter 2.
Specific aim 2: Evaluate the Population Structure and Virulence Potential of V. cholerae from Environmental Sources in the Apalachicola Bay.
The potential risks of V. cholerae populations in Apalachicola Bay, Florida were
determined by multiple molecular methods, which were used for comparison to
toxigenic 7th pandemic V. cholerae O1/O139 strains and to the O75 strain associated
with the 2011 Florida outbreak. These results are described in Chapter 3.
Specific aim 3: Determine the Anti-Vibrio Potential of Chitosan in Seawater and Oysters.
The effect of chitosan against pathogenic Vibrio spp. was examined under
commercially relevant conditions and in live oysters in order to determine the feasibility
of chitosan as a PHP treatment for seafood industry. The contribution of capsular
polysaccharide to the activity of chitosan against Vibrios was evaluated. These results
are described in Chapter 4 and 5.
42
CHAPTER 2 DISTRIBUTION OF V. CHOLERAE IN SEAWATER AND OYSTERS FROM
APALACHICOLA BAY, FLORIDA
Introduction
Vibrio cholerae is the causative agent of cholera, which remains a global health
problem, as the World Health Organization (WHO) reported that the annual number of
cholera cases increased in the past few years to more than half a million cases with
7816 related deaths from all reporting regions (WHO, 2013). Unlike many human
pathogens, Vibrio spp. including V. cholerae are clearly autochthonous in estuarine
ecosystems. The warming of coastal waters is likely to enhance the growth and
persistence of this bacterium in estuarine niches, and has been proposed as a
contributing factor to increased occurrence of outbreaks (Vezzulli et al., 2012). For
example, recent emergence of cholera in Haiti (2010) and Cuba (2012) is intricately
linked to the 7th pandemic strain of Asiatic V. cholerae O1 El tor, demonstrating a return
of cholera from east to west (Weil et al., 2012). Conversely, a recent outbreak (Onifade
et al., 2011) on the Florida Gulf Coast involved a divergent serotype O75, which is
genetically distinct from the pandemic strain and may represent a resident population
that is sustained in this geographic location. Deciphering the role of environmental
conditions as a factor in driving the persistence and abundance of V. cholerae is the key
to mitigating potential health risks of this pathogen to humans.
Ecological-based models that define environmental parameters that are relevant
to the potential risk of cholera have been proposed in several studies. For example, the
function of temperature and salinity served as a predictive model for the presence of
V. cholerae in Chesapeake Bay (Louis et al., 2003), and rainfall was confirmed as an
43
effective predictor of V. cholerae prevalence in the Great Bay Estuary of New
Hampshire (Schuster et al., 2011). However, the occurrence, survival, and public health
threat of V. cholerae in Gulf Coast states warrant further investigation. The study
described herein characterized the distribution of V. cholerae in seawater and oysters
collected at various sites in Apalachicola Bay, Florida, which is the prime harvest site for
oysters, contributing about 90% of the state’s oyster harvest. A smaller subset of
samples was collected from fish, sediment and plant. The role of environmental
parameters, as well as the prevalence and distribution of the other major pathogens in
this genus, namely V. vulnificus and V. parahaemolyticus, were also evaluated with
respect to V. cholerae occurrence and abundance.
Materials and Methods
Samples Collection and Processing
Water and oyster samples were collected seasonally from 2012 to 2014 among
17 sites of Apalachicola Bay in the northern Gulf of Mexico with the assistance of the
Florida Department of Agriculture and Consumer Services (DACS), the Florida
Department of Environmental Protection, and the Apalachicola National Estuarine
Research Reserve (Figure 2-1). Environmental parameters were recorded, which
included temperature, salinity, pH, dissolved oxygen, turbidity and conductivity. Live
oysters were collected using oyster tongs, and 1 to 2 L of surface seawater was
collected into sterile plastic containers or autoclaved glass bottles. In addition, various
plants, the top 5 cm sediments and trawled fish samples were collected occasionally
and screened for Vibrio spp. All samples were collected in triplicates for each date, site
44
and environmental source. All the samples were transported with ice packs and
assessed within 4 h.
Isolation and Enumeration of Vibrios
Any dirt or debris found on oysters shells was cleaned, and live oysters were
shucked and weighted in sterile bottles to avoid cross-contamination. Samples (n=3) of
oyster meats were diluted 1:1 in phosphate-buffered saline (PBS; pH 7.4) and
homogenized for 60 s in a Waring blender, and serially diluted in PBS. Homogenate (2
ml) was transferred to 8 ml alkaline peptone water (APW; pH 8.5; 1% NaCl), and diluted
samples (1 ml) were transferred to 9 ml APW (1:10) for inoculation of triplicate
enrichment cultures. Volumes of 10, 1.0 and 0.1 ml seawater samples were also
inoculated into a three-tube multiple analysis series with APW selective enrichment.
Plant or sediment samples (n=3) were placed in APW enrichment tubes (10 or 25 ml)
and vortexed 30 s. Large fish were killed with ms222 and intestine (approximately 10
cm) was dissected and placed in 25 ml APW. Small fish were swabbed at anal cavity
and swabs placed in 25 ml APW enrichment tubes. All enrichment tubes were incubated
for 16-24 h at 37°C and subsequently streaked onto modified cellobiose-polymyxin B-
colistin (mCPC) (Massad and Oliver, 1987), thiocitrate bile salts sucrose agar (TCBS,
Difco), CHROMagar™ Vibrio (CHROMagar Microbiology), and Vibrio chromogenic agar
(Hardy Diagnostics) and incubated for another 16-24 h at 37°C or room temperature
(25°C). Typical V. cholerae colonies displaying yellow on TCBS or light blue on
CHROMagar™ or mauve on Hardy chromogenic agar were cross-streaked to another
selective agar. Following incubation 16-18 h at 37°C, a representative colony that was
positive on multiple plates would be recognized putatively as V. cholerae. Similarly,
typical colonies displaying yellow on mCPC and mauve on CHROMagar™ Vibrio were
45
recognized putatively as V. vulnificus and V. parahaemolyticus respectively. All
presumptive positive were swabbed to non-selective Luria-Bertani broth with NaCl
(LBN, 1.0% tryptone, 0.5% yeast extract, 1.0% NaCl in DI water, pH 7.4) agar (LA) and
LBN in 50% glycerol for further test. Frozen stocks were used for PCR confirmation as
described below, and MPN calculations were performed based on confirmed positive
cultures of V. cholerae, V. parahaemolyticus, and V. vulnificus using MPN calculator
according to the BAM (FDA, 2011).
DNA Extraction and Species Identification
All putative V. cholerae, V. parahaemolyticus and V. vulnificus isolates collected
from selective agar were subjected to a PCR assay for species identification using
primers derived from species-specific DNA (Table 2-1). DNA was extracted from all
isolates using a boiling method. Briefly, all isolates were inoculated into each 5 ml LBN
and incubate at 37°C overnight. Bacteria was collected by centrifugation at 13,000 rpm
for 3 min, and the cell pellet was resuspended by 400 l phosphate-buffered saline and
boiled at 100°C for 7 min followed by the same centrifuge step. The supernatant was
transferred to a new microcentrifuge tube and stored at -20°C until used as a template
for a PCR reaction. V. cholerae positive was confirmed by16S-23S rRNA intergenic
spacer region based on PCR assay (Chun et al., 1999). One l DNA extract was mixed
with 2.5 l 10x buffer (5 PRIME), 400 nM of each deoxinucleotidetriphospate (dNTP,
Invitrogen), 400 nM of each primer (Sigma-Aldrich) and 0.25 l of Taq polymerase (5
PRIME) in a final volume of 25 l in PCR water. The amplification cycle was initial
denaturation at 94°C for 2 min, followed by 35 cycles of 94°C for 45 s, 60°C for 1 min,
72°C for 45 s, and one final extension at 72°C for 5 min at the end of 35 cycles. Each
46
amplified fragment (10 l) was mixed with one l of 6 x loading dye (Qiagen), separated
by electrophoresis on a 1% agarose gel and visualized under UV light after staining with
0.5 g/ml ethidium bromide (EtBr, Fisher Scientific Inc.). Presumptive V. vulnificus and
V. parahaemolyticus isolates were confirmed by PCR based on the hemolysin gene
vvhA (Warner and Oliver, 2008) and thermolabile hemolysin gene tlh (Bej et al., 1999),
respectively, following the same PCR protocol as described above, except the
annealing temperature for V. vulnificus and V. parahaemolyticus was 58°C.
Analysis of Abundance with Environmental Conditions
To achieve normally distributed data sets, all bacteria concentrations in water
and oysters were transformed to log MPN/100 ml and log MPN/10 g. Normal logistic
regression analysis was used to determine environmental condition correlations with the
distribution of V. cholerae occurrence or abundance in water and oysters. All statistical
analyses were performed using JMP pro 11 (SAS, Cary, NC). An alpha level of 0.05
was considered the minimum level for statistical significance.
Results
Distribution of V. cholerae in Apalachicola Bay, Florida Relative to Other Pathogenic Vibrios.
Following the outbreak of toxigenic V. cholerae serogroup O75 in 2011, a survey
was conducted to better understand the ecology and potential risk of V. cholerae related
to Gulf Coast oysters. From 2012 to 2014 water (n=138), oyster (n=60) and small set of
fish samples (n=33) were collected from 17 sites to survey for V. cholerae in
Apalachicola Bay. Comparison of the percent of positive samples was used to infer
prevalence of one species over the other two across sample types. Overall,
V. parahaemolyticus and V. vulnificus were significantly more frequent in all seawater
47
and oyster samples than V. cholerae (p<0.001). All oyster samples were positive for
V. vulnificus and V. parahaemolyticus, and the occurrence of positive seawater samples
was slightly lower for both V. vulnificus (93%) and V. parahaemolyticus (76%) (Figure 2-
1). Conversely, V. cholerae prevalence was 48% and 50% for oysters and seawater,
respectively. Furthermore, no fish samples were found to be positive for V. cholerae.
On the contrary, the prevalence of V. parahaemolyticus (83%) and V. vulnificus (67%) in
these fish samples was relatively high and showed a large diversity of fish species
(king, spot, croaker, file fish, sand sea trout, silver trout, catfish, flounder, and
anchovies).
Sampling sites were widely distributed throughout the Bay, and occurrence of
V. cholerae was compared to that of V. vulnificus and V. parahaemolyticus on a site-by-
site basis (Tables 2- 2, 2-3, and 2-4). Levels of V. cholerae ranged from 0.3 to 3.6 log
MPN/10 g with a mean of 1.63 log MPN/10 g for oyster samples when V. cholerae was
detected. In seawater, V. cholerae levels ranged from 0.6 to 3.0 log MPN/100 ml with a
mean of 1.64 log MPN/100 ml, which was relatively similar regarding mean levels of V.
parahaemolyticus (1.63 log MPN/100 ml) and V. vulnificus (1.69 log MPN/100 ml) in
seawater. Although sampling time points were limited, the distribution of V. cholerae
appeared to be more site-specific compared to that of the other species. As shown in
Table 2-5, bay shore (BS) sites located nearer the mouth of the Apalachicola River
(sites 4, 5, 6, 7, 8, 12, and 15) were more likely to show positive for V. cholerae, as
compared to mid-bay (MB) site (sites 1, 2, 3, 13, 14, 17, and 18) and sites with highest
salinity on the ocean side of the barrier island (BI) (sites 9, 10, 11, and 16). Conversely,
48
V. parahaemolyticus was less prevalent for BS sites (77%) compared to MB (88%) and
BI (90%) sites (Table 2-5).
Relationship of V. cholerae Occurrence and Abundance to Environmental Parameters.
Various environmental factors appeared to influence V. cholerae prevalence in
Apalachicola Bay. Salinity ranged from 0 to 36 ppt, temperature from 18 to 30.4°C,
dissolved oxygen levels from 5.5 to 11.8 mg/L, pH from 6.5 to 8.6, and conductivity from
0.63 to 53.1 s/m (Figure 2-3). As expected, high temperature (up to 30.4°C) was
observed in the summer months compared to the winter and went down to 18°C, while
pH was relatively constant. However, much greater fluctuation was seen over time for
pH with unusually high levels, presumably as a consequence of reported river flow due
to diversion of water from the Apalachicola River (Petes et al., 2012).
Positive V. cholerae samples were more frequently detected at sites where the
salinity level was relatively low, and logistic regression analysis (Table 2-4) revealed a
strong negative relationship between the salinity and the presence of V. cholerae in
oysters (R2 = 0.46, p = 0.0096) and water samples (R2=0.50, p < 0.001). For example,
77% of collected water samples were positive when the salinity was lower than 12 ppt
(Figure 2-3), although no positive samples were detected at the site 12 up the river with
0 ppt. Furthermore, V. cholerae for oyster and water samples was more frequently
(72%) detected when water temperature exceeded 22°C, but overall there was no
significant association between temperature levels and the occurrence of V. cholerae.
The presence of V. cholerae in oyster samples also showed a positive correlation to
dissolved oxygen (R2 = 0.41, p < 0.0447), but not with the occurrence in the seawater.
V. cholerae was detected at similar frequencies in water and oysters over the pH ranges
49
observed, except no oyster samples were collected below pH 7.2, and it is likely that the
pH conditions at all of the study sites were within the optimal range for this species
(Table 2-6). Results with respect to the effects of conductivity were similar to those for
salinity and showed a strong inverse relationship to the detection of V. cholerae for both
water (R2=0.60, P < 0.001) and oyster samples (R2=0.99, P < 0.001).
The levels of culturable V. cholerae isolates varied from 0.3 to 3.6 log MPN/10 g
and 0.6 to 3.0 log MPN/100 ml in oysters and seawater, respectively. Highest levels
were observed with conditions of low salinity and conductivity, medium temperature,
and high oxygen (Figure 2-4). Logistic correlations in respect to environmental
parameters and abundance of V. cholerae were similar to those observed for
prevalence (Table 2-7).
Discussion
The role of climate in determining the abundance of Vibrios in marine and
estuarine environments is not a new concept. In this study, salinity and temperature
were monitored along with other environmental parameters that may affect the
occurrence and abundance of V. cholerae and other Vibrio spp. in the water column and
in oysters from Apalachicola Bay from 2010 to 2014. The study showed that V. cholerae
was detected more often in water and oysters, as compared to fish samples, and was
significantly less abundant in Apalachicola Bay than the other pathogenic Vibrios
examined in this study. A significant negative correlation between the prevalence of
V. cholerae and salinity has been described previously (Louis et al., 2003) and was
reaffirmed by this survey. V. cholerae isolates in Apalachicola Bay were mostly
collected from sites with salinity ranging from 0 ppt to 12 ppt. These observations are in
agreement with previous studies done in Southern California, where V. cholerae
50
detected at salinities ranging from 0 to 10 ppt (Jiang, 2001). In addition to affect the
abundance of V. cholerae in environment, low salinity also contributes to the expression
of virulence genes in V. cholerae. Previous study shown that salinity between 2 ppt to
2.5 ppt is optimal for the expression of CTX (Tamplin and Colwell, 1986)
This study also showed conductivity had a significant association with
V. cholerae in both water and oysters. Conductivity measures water quality based on
total inorganic dissolved solids, which mostly reflects salinity in seawater. Most bodies
of water maintain a constant conductivity that affect water quality and aquatic life. Based
on the data, the prevalence of V. cholerae in both seawater and oysters more strongly
correlated with conductivity than salinity, and this was particularly noticeable regarding
the abundance of V. cholerae in oysters. Since most previous studies focus on how
salinity affect the distribution of V. cholerae in environment, this study offers new insight
into conductivity that also accounts for the complexity of V. cholerae prevalence.
A positive relationship with dissolved oxygen in the water column was observed
with V. cholerae density in oysters but not V. cholerae density in water column. This
may reflect the patchiness of the data or potentially, this relationship could reflect a
close association of V. cholerae with zooplankton environments, as dissolved oxygen
content is vital for algae and aquatic animal growth. Perhaps, oysters harbor algal
populations during conditions of oxygen stress, which might affect the survival and
persistence of V. cholerae. Some studies demonstrated that blooms of algae and other
phytoplankton have a moderate effect on the population of V. cholerae and other Vibrio
spp. (Romalde et al., 1990; Spira et al., 1981).
51
The temperature ranged from 17C to 30C throughout the course of sampling
period. Even in the winter months, water temperatures still exceeded 20C in some
sampling sites, which probably accounts for the lack of seasonality in this survey.
Although this was not a comprehensive survey, seasonal observations showed no
significant trend for V. cholerae occurrence versus temperature in any sample types,
which is surprising in light of retrospective studies (Blackwell and Oliver, 2008). In
comparing these observations to the findings of previous studies, it should also be
noted that a significant linear relationship cannot be identified when temperature does
not vary over a sufficiently wide range. Studies that identified the importance of
temperature to V. cholerae included more exhaustive sampling and all were collected
over a wider range of temperature than that of the current study. Nevertheless, warm
temperature in combination of sufficient dissolved oxygen and elevated pH still plays an
important role in V. cholerae growth and multiplication in aquatic environment,
particularly in association with copepods (Huq et al., 1990).
In an era of warming coastal waters and increasing vibriosis incidence, this study
provides new data for understand the complex and dynamic factors affecting the
distribution of toxigenic V. cholerae in Florida oysters and shellfish harvesting waters.
At present regulations and mandates pertaining to Vibrios do not include V. cholerae.
The data indicate that although the public health risk of cholera in the US from Florida
seafood in extremely low, the prevalence and close relationship of environmental
isolates to outbreak strains warrants future monitoring and study. These data will
support future policy decisions and influence management practices to reduce or
eliminate the risk of this pathogen for the seafood industry.
52
Figure 2-1. Sampling sites in Apalachicola Bay.
53
Table 2-1. Sequences of oligonucleotides used for molecular analysis
Targeta Geneb
Primer set
Fragment length (bp)
V. cholerae 16S gene rrnA F 5’ TTAAGCSTTTTCRCTGAGAATG 3’
301
rrnA R 5’ AGTCACTTAACCATACAACCCG 3’
V. cholerae cholera toxin gene
ctxA F 5’ GGCTGTGGGTAGAAGTGAAACGG 3’
1140
ctxA R 5’ CTAAGGATGTGGAATAAAAACATC 3’
V. cholerae toxin co-regulated pilus gene
tcpA R 5’ AAAACCGGTCAAGAGGG 3’
600
tcpA F
tcpA R
5’ CAAAAGCTACTGTGAATGG 3’ 5’ CAAATGCAACGCCGAATGG 3’
V. vulnificus hemolysis gene
vvhA F 5’ AGCGGTGATTTCAACG 3’
411
vvhA R 5’ GGCCGTCTTTGTTCACT 3’
V. parahaemolyticus thermal labile hemolysin gene
tlhA F 5’ GCTACTTTCTAGCATTTTCTCTGC 3’ 450
tlhA R 5’ AAAGCGGATTATGCAGAAGCACTG 3’
a. Gene target and species are given. b. Gene name for forward (F) and Reverse (R) primers are shown.
54
Figure 2-2. Overall distribution of Vibrio spp. in water and oysters. The occurrence rate
of V. cholerae (Vc), V. vulnificus (Vv) and V. parahaemolyticus (Vp) in water oysters, and fish are presented as determined by percent of positive samples (%). ND= No bacteria were detected from MPN plated to selective agars.
55
Table 2-2. Spatial distribution of V. cholerae in water and oysters from Apalachicola Bay.
Matrix Site (Location)
Abundance of V. cholerae in Apalachicola Bay (log MPN/10 g of oyster or log MPN/100 ml of
seawater)a
Jul.-Aug. 2012
Nov. 2012
Feb. 2013
June 2013
Dec. 2013
May 2014
Oyster 1 (MB) - - - 2.6 ND - 2 (MB) ND - - - - - 3 (MB) ND - ND* 1.3 - - 7 (BS) - - - ND 2.4 1.2 14 (MB) - - - ND - - 15 (BS) ND - - 2.3 0.6 ND 18 (MB) - - - - - ND Water 2 (MB) ND - - - ND - 3 (MB) ND - ND 2.4 ND - 4 (BS) 1.9 3.0 - 2.4 1.6 1.3 5 (BS) 1.6 1.0 2.2 2.4 0.8* ND 6 (BS) 0.9 - - - 1.9 ND 7 (BS) - - - ND 1.2 ND 8 (BS) - - - - ND - 9 (BI) ND 1.0 - - ND ND 10 (BI) 1.4 - - ND ND - 11 (BI) - - - - ND ND 12 (BS) - - - ND - - 13 (MB) 1.0 ND - ND - - 14 (MB) - ND - 1.6 - - 15 (BS) - - - 3.0 1.3* 1.0 16 (BI) ND - - - - - 17 (MB) - - - - ND -
a. Abundance (log MPN/10 g of oyster or log MPN/100 ml of seawater) was determined by MPN as described in the methods.
b. “*” denotes sites that were sampled twice on specific month. “–” denotes sites that were not sampled during sampling month. ND = No bacteria were detected from MPN plated to selective agars. MB= Mid-Bay; BS= Bay Shore; BI= Barrier Island.
56
Table 2-3. Spatial distribution of V. vulnificus in water and oysters from Apalachicola Bay.
Matrix Site (Location)
Abundance of V. vulnificus in Apalachicola Bay (log MPN/10 g of oyster or log MPN/100 ml of
seawater)a Jul.-Aug. 2012
Nov. 2012
Feb. 2013
June 2013
Dec. 2013
May 2014
Oyster 1 (MB) - - - 2.4 2.0 - 2 (MB) 1.5 - - - - - 3 (MB) 1.4 - 0.9* 4.0 - - 7 (BS) - - - 2.7 2.0 3.0 14 (MB) - - - 3.4 - - 15 (BS) 1.4 - - 3.7 1.2 1.0 18 (MB) - - - - - 2.7 Water 2 (MB) 2.3 - - - ND - 3 (MB) 2.4 - 0.9 1.6 ND - 4 (BS) 1.5 1.2 - ND 3.0 ND 5 (BS) 2.5 3.0 3.0 0.6 1.1* ND 6 (BS) 1.4 - - - 1.1 0.9 7 (BS) - - - 2.7 3.0 3.0 8 (BS) - - - - 3.0 - 9 (BI) 2.0 1.1 - - ND 1.2 10 (BI) 2.1 - - 0.5 0.8 - 11 (BI) - - - - 1.5 3.0 12 (BS) - - - ND - - 13 (MB) 1.2 0.5 - 0.6 - - 14 (MB) - 2.4 - 1.4 - - 15 (BS) - - - 1.0 2.5* 1.4 16 (BI) 0.8 - - - - - 17 (MB) - - - - 0.5 -
a. Abundance (log MPN/10 g of oyster or log MPN/100 ml of seawater) was determined by MPN as described in the methods.
b. “*” denotes sites that were sampled twice on specific month. “–” denotes Sites that were not sampled during sampling month. ND = No bacteria were detected from MPN plated to selective agars. MB= Mid-Bay; BS= Bay Shore; BI= Barrier Island.
57
Table 2-4. Spatial distribution of V. parahaemolyticus in water and oysters from Apalachicola Bay.
Matrix Site (Location)
Abundance of V. parahaemolyticus in Apalachicola Bay (log MPN/10 g of oyster or log MPN/100 ml of
seawater)a Jul.-Aug. 2012
Nov. 2012
Feb. 2013
June 2013
Dec. 2013
May 2014
Oyster 1 (MB) - - - 2.0 0.3 - 2 (MB) 1.2 - - - - - 3 (MB) 1.1 - 3.0* 4.0 - - 7 (BS) - - - 3.7 0.5 3.0 14 (MB) - - - 4.0 - - 15 (BS) 1.3 - - 3.7 2.0 3.0 18 (MB) - - - - - 3.0 Water 2 (MB) 2.4 - - - 1.4 - 3 (MB) ND - 0.8 1.5 1.2 - 4 (BS) 0.5 1.7 - 1.4 ND ND 5 (BS) ND 2.3 1.2 0.6 1.4* ND 6 (BS) 0.8 - - - ND 3.0 7 (BS) - - - 1.9 1.0 3.0 8 (BS) - - - - 1.4 - 9 (BI) 1.2 2.1 - - 1.2 0.9 10 (BI) 1.2 - - 1.6 2.7 - 11 (BI) - - - - ND 2.2 12 (BS) - - - ND - - 13 (MB) 3.0 ND - 1.2 - - 14 (MB) - 0.6 - 1.4 - - 15 (BS) - - - 1.4 2.2* 3.0 16 (BI) 2.0 - - - - - 17 (MB) - - - - ND -
a. Abundance (log MPN/10 g of oyster or log MPN/100 ml of seawater) was determined by MPN as described in the methods.
b. “*” denotes sites that were sampled twice on specific month. “–” denotes Sites that were not sampled during sampling month. ND = No bacteria were detected from MPN plated to selective agars. MB= Mid Bay; BS= Bay Shore; BI= Barrier Island.
58
Table 2-5. Distribution of Vibrios by general location.
Species Percent of Positive Samples by Locationa
Bay Shore Mid-bay Barrier Island
V. cholerae 71% 32% 20%
V. vulnificus 87% 92% 90%
V. parahaemolyticus 77% 88% 90%
a. Locations for include Bay Shore (site 4, 5, 6, 7, 8, 12 and 15), Mid bay (site 1, 2, 3, 13, 14, 17 and 18), and Barrier Island (site 9, 10, 11, and 16).
59
Figure 2-3. Relationship of V. cholerae prevalence to various water parameters. The %
of V. cholerae positive samples are shown as a function of high, medium and low A) salinity (ppt), B) conductivity (S/m), C) dissolved oxygen (mg/L), D) pH, and E) temperature (°C).
60
Table 2-6. Overall relationship of environmental conditions to prevalence of V. cholerae.
Environmental
parameters
Seawater (N=138) Oyster (N=60)
Relative
importance a
P Value Relative
importance
P Value
Salinity 0.498 <0.001 0.457 0.0096
pH 0.023 0.451 0.033 0.591
Dissolved O2 0.018 0.559 0.410 0.0447
Temperature 0.001 0.832 0.021 0.625
Conductivity 0.600 <0.001 0.990 <0.001
a Relative importance is based on pseudo-R2 statistics derived from logistic regression analysis.
61
Figure 2-4. Relationship of V. cholerae levels to various water parameters. V. cholerae levels (log MPN/100 ml or 10 g) are shown with respect to A) salinity (ppt), B) conductivity (S/m), C) dissolved oxygen (mg/L), D) pH, and E) temperature (°C). ND = No oyster samples were collected in low pH and high conductivity level.
62
Table 2-7. Overall relationship of environmental conditions with abundance of V. cholerae.
Environmental
parameters
Seawater (N=138) Oyster (N=60)
Relative
importance a
P Value Relative
importance
P Value
Salinity 0.21 0.0013 0.55 0.0010
pH 0.08 0.2355 0.16 0.5956
Dissolved O2 0.01 0.7560 0.40 0.0368
Temperature 0.03 0.4394 0.00 0.9320
Conductivity 0.31 0.0047 0.82 0.0003
a Relative importance is based on pseudo-R2 statistics derived from logistic regression analysis.
63
CHAPTER 3 GENETIC CHARACTERIZATION AND VIRULENCE POTENTIAL OF V. CHOLERAE
FROM APALACHICOLA BAY
Introduction
Although V. cholerae is comprised of more than 200 serotypes, epidemic cholera
is generally limited to toxigenic strains of V. cholerae serogroups O1 and O139 that
carry the genes for cholera toxin (ctxA, ctxB) (Lee et al., 2006). The signs and
symptoms of cholera are primarily caused by cholera toxin, and most V. cholerae from
environmental sources do not have the genes required for expression of this toxin.
Although non-O1/ O139 strains are occasionally associated with diseases such as
diarrhea and septicemia (Singh et al., 2001), the pathogenic potential of these non-
epidemic V. cholerae populations is relatively limited. Emerging serotypes could be
reservoirs of new pathogenic lineages that have acquired increased virulence potential
by horizontal gene transfer and recombination. For example, only eight sporadic cases
of non-pandemic V. cholerae O75 in the US were reported to CDC between 2003 and
2007 (Tobin-D'Angelo et al., 2008); however, a recent cholera outbreak attributed to
serogroup O75 caused at least 10 confirmed cases in March 2011 (Onifade et al.,
2011). This outbreak aroused concerns among governmental officials and scientists
because it is the first documented US outbreak attributed to a toxigenic V. cholerae non
O1/O139 strain and oyster consumption.
In this chapter, V. cholerae strains collected from Apalachicola Bay were
characterized in order to understand the virulence potential and genetic structure of
environmental V. cholerae populations relative to toxigenic 7th pandemic V. cholerae
and to the strains and the 2011 Florida outbreak.
64
Materials and Methods
Strains and Culture Conditions
All Apalachicola Bay isolates confirmed as V. cholerae by 16S rRNA were stored
in Luria broth (LBN; 1.0% tryptone, 0.5% yeast extract, 1.0% NaCl in deionized water,
pH 8.4) with 50% glycerol at -80°C. For each experiment, strains were retrieved from
the frozen stock and streaked for isolation on LB with 1.5% agar (LA). Two Tampa Bay
strains were provided by Dr. Harwood for genetic characteristic comparison.
PCR Screening for Virulence Genes
All isolates confirmed as V. cholerae by 16S rRNA gene were further analyzed
for virulence potential by PCR analysis of genes encoding cholera toxin (ctxA) and toxin
co-regulated pilus (tcpA), using the same protocol as described in chapter 2 (Table 2-1).
Phylogenetic Characterization
A total of 35 strains were selected from different sites and sampling times as a
representative subset for genetic characterization. Whole genome sequencing was
performed for those isolates using the Genome Analyzer IIx system (Illumina, Inc., San
Diego, CA) according to the manufacturer‘s methods. Raw reads of these genomes
were assembled with UFRC Galaxy program. Genome-to-genome comparisons,
identification and characterization of molecular genetic elements were through Center
for Genomic Epidemiology (CGE) pipeline (http://www.genomicepidemiology.org/).
Subsequently, a fifteen-locus MLST (recA, gyrB, topA, pyrH, gapA, adk, mdh,
mete, pnta, purm, pyrc, toxR, ompU, hlyA, and rtxA) was performed to assess the
phylogenetic and epidemiological relatedness of identified environmental V. cholerae
strains. DNA sequences were concatenated manually and aligned by Clustal W and
downloaded to MEGA program version 5.0 to generate a dendrogram. Phylogenetic
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trees were constructed by the neighbor joining algorithm with 1000 bootstrap.
Sequences from toxigenic V. cholerae strains including O1 classical biotype (O395, I-
1300, IEC-224), O1 El Tor biotypes (2010EL-1786, 16961, MJ-1236, 2012EL-2176,
M66-2), and O75 were derived from NCBI GenBank and included in phylogeny
analyses. Sequences for additional reference V. cholerae strains MO10 (O139;
Bangladesh), V51 (O141; United States), 1587 (O12; Peru), AM-19226 (O39;
Bangladesh), MZO-2 (O14; Bangladesh), and MZO-3 (O37; Bangladesh) were retrieved
from the Broad Institute (http://www.broadinstitute.org/) and included in phylogeny
analyses. All Environmental strains (35) and clinical genomes (17) used in this study are
summarized in Table 3-1
Antibiotic Susceptibility Test
Antimicrobial susceptibility test was performed for those 35 isolates against a
variety of antimicrobial agents using disc diffusion assay and following the
manufacturer’s protocol (BBL™ Sensi-Disc™). Briefly, V. cholerae strains were
streaked on LA agar for isolation and incubated overnight at 37C. One or two overnight
colonies from LA plates were emulsified to one ml of sterile saline (0.85% NaCl), and
the turbidity of the bacterial suspension was compared to a 0.5 McFarland Standard.
Subsequently, culture was dipped and spread evenly as lawn by a sterile cotton swab
onto Sensitest agar plates (Oxoid, England), and the plates were allowed to dry for 10
min. Antibiotic discs with the following drug concentrations, including kanamycin
(30 g), sulfamethoxazole (23.75 g)-trimethoprim (1.25 g), ciprofloxacin (5 g),
ceftriaxone (30 g), nalidixic acid (30 g), streptomycin (10 g), cephalothin (30 g),
amikacin (30 g), amoxicillin-clavulanic acid (30 g), and tetracycline (10 g) were
66
placed on the plates. No more than five equidistant antibiotic discs were applied per
plate to prevent the overlapping of zones of inhibition. Zones of inhibition of each
antibiotic were measured after 18-24 h incubation at 37C, and susceptibility or
resistance pattern of the V. cholerae isolate to the antibiotic was compared with the
recorded diameters of the control organism E. coli ATCC 25922. Bacteria were
classified as resistant, intermediate or sensitive based on clinical laboratory test
standard (Clinical and Laboratory Standards Institute, 2009)
Results
Genetic Characterization of V. cholerae from Apalachicola Bay
Out of more than 400 putative V. cholerae colonies recovered from selective
media, 119 isolates were confirmed positive by 16S rRNA - specific PCR. All isolates
confirmed as V. cholerae by 16S rRNA gene were screened for virulence potential by
PCR analysis; however, cholera toxin gene (ctxA) and toxin co-regulated pilus gene
(tcpA), the genes associated with cholera epidemics, were absent from all Apalachicola
Bay isolates. A representative subset of 35 isolates was further selected from different
sites, sources, and sampling time points and further analyzed for genetic
characterization and antibiotic resistance (Table 3-2). Whole genome sequence was
used to map specific genes as closed reference genome from V. cholerae O1 biovar El
Tor str. N16961, a (GenBank accession no. NC_002505.1 and NC_002506.1),
representing the ongoing seventh pandemic clone.
All Apalachicola isolates presented the virulence gene for the outer membrane
protein U (ompU), and most isolates encoded other virulence-associated genes that
encode virulence regulation protein ToxR (toxR; 94%), hemolysin (hlyA; 94%), and
repeats-in-toxin (rtxA; 88%), indicating a broad distribution of these virulence factors in
67
environmental strains (Table 3-3). Protein genes were also compared to clinical strains
through CGE database. An average of 138 protein genes were matched to pathogenic
families, and a mean of three protein genes not matched to pathogenic families (Table
3-2). It is not clear if these virulence genes and protein genes influence the ecology or
virulence potential of these isolates; however, they do demonstrate an appreciable level
of genomic diversity among environmental V. cholerae strains from Apalachicola bay.
Phylogenetic Analysis of V. cholerae Population in Apalachicola Bay
In order to examine the diversity and phylogenetic relationships of those
environmental strains to epidemic strains, a MLST analysis was performed based on
eleven housekeeping genes (recA, gyrB, topA, pyrH, gapA, adk, mdh, mete, pnta, purm,
and pyrc) and four virulence genes (toxR, ompU, hlyA, and rtxA). Results demonstrated
that V. cholerae O75 formed a monophyletic lineage with V. cholerae V51, a clinical
V. cholerae O141 serogroup strain associated with sporadic cholera-like diarrhea in the
US, suggesting O75 and O141 serogroup had a common ancestor after it had diverged
from other V. cholerae lineages responsible for cholera epidemics (Figure 3-1). In
addition, V. cholerae O75 strain was phylogenetically close to other non-O1 and non-
O139 serotype clinical strains, including O12, O14, O37 and O39 strains, but was
divergent from the V. cholerae 7th pandemic strains O1 El tor, classical O1, and O139
strains. The closest Florida isolate from the present study was an oyster isolate from
Tampa Bay. Some Apalachicola strains clustered more closely with clinical O1 strains
and the O139, while others were grouped with O75 and other non-O1/O139, illustrating
the genetic diversity of V. cholerae strains isolated from environmental sources.
Interestingly, the oyster isolates from Apalachicola Bay were more clonal than either
seawater isolates from the same source or oyster isolates from Tampa Bay.
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Antibiotic Resistance of V. cholerae in Apalachicola Bay
In order to determine the susceptibility of these 35 strains to antibiotics, each
V. cholerae isolate was subjected to antibiotic susceptibility assay to antimicrobial
agents. Most strains (77%) were sensitive or showed only intermediate resistance to all
antibiotics assessed. All isolates were sensitive to sulfamethoxazole-trimethoprim,
ciprofloxacin, ceftriaxone, and nalidixic acid (Table 3-3). V. cholerae strains were
resistant or showed intermediate resistance to amoxicillin-clavulanic acid (40%),
followed by streptomycin acid (43%), amikacin (22%), and tetracycline (11%). Five
isolates (14%) exhibited multidrug resistance (resistant to two or more antibiotics),
including three isolates that were resistant to four antibiotics and two isolates that were
resistant to three antibiotics.
Discussion
V. cholerae continues to cause devastating diarrheal disease in many areas of
the world and remains as a significant global concern. To date, the vast majority of
research focuses on O1 and O139 strains, and a recent outbreak of pandemic O1
V. cholerae in 2010 in Haiti highlights the potential for spread to the western
hemisphere. Furthermore, a V. cholerae O75 Florida outbreak in 2011 demonstrated
the potential role of non-O1/O139 strains in causing epidemics (Chapman et al., 2015;
Onifade et al., 2011). Changing environmental conditions in Florida waters and
inappropriate food safety practice have been proposed as critical factors that may have
contributed to this outbreak. Anthropogenic impacts that are increasing along coastal
water may function to enhance the growth and persistence of V. cholerae in Florida
waters, and this may be further aggravated by the emergence of new pathogenic
69
lineages derived from environmental strains through horizontal gene transfer and
recombination (Haley et al., 2014).
MLST analysis revealed a diverse population of V. cholerae from Apalachicola
Bay; however, these results elucidate the relationship of a phyletic lineage of
V. cholerae O75 outbreak strain to other Florida isolates, as most strains from
Apalachicola Bay (52%) were more closely related to that clade, as compared to clades
with O1 strains (24%) or more distant non-O1/O139 strains (24%). Strains from Tampa
Bay (n=2) were also in the O75 clade, and one strain showed the closest genetic
relationship to the O75 strain among all other Florida isolates. Although all
environmental strains lacked ctxA and tcpA genes, the presence of virulence genes (rtx,
ompU, hlyA, toxR) in Florida isolates suggests a wide distribution of those virulence
factors in environmental and clinical isolates. From a public health perspective, these
results elucidate the persistence of V. cholerae O75- related strains in Gulf Coasts as a
cause of infectious disease. The present study provides a snapshot of the genetic
complexities that are present in V. cholerae strains in US northern Gulf region. Oyster
reservoirs in Apalachicola Bay may play an important role in the positive selection and
dissemination of genetic elements since all the V. cholerae strains (n=5) isolated from
oysters in Apalachicola Bay exhibited a close genetic similarity in contrast to the oyster
isolates from Tampa Bay, which were more diverse.
The antibiotic resistance analysis revealed a range of patterns among
V. cholerae strains in Apalachicola Bay. Diverse resistance to different classes of
antibiotics, including β-lactams (14% for amoxicillin-cavulanic acid), cephalosporins (9%
for cephalothin), aminoglycosides (11% for amikacin and 3% for kanamycin), as well as
70
streptomycin and tetracycline (both 11%), were observed in these environmental
isolates. All Florida isolates were sensitive to the antibiotics in the classes for inhibiting
DNA or folic acid synthesis. These results were differed from a report of Haitian
environmental isolates which shown resistant to nalidixic acid (DNA synthesis inhibitor)
and cotrimazole (folic acid synthesis inhibitor). The emergence of multi-drug antibiotic
resistant phenotype was seen from five strains, suggesting a contribution to persistence
in environmental reservoirs. The presence of multiple antibiotic resistance genes was
also confirmed from two isolates and provided evidence of potential dissemination of
antibiotic resistance genes from clinical pathogens to environmental bacteria.
In summary, V. cholerae from Apalachicola Bay seawater are highly diverse,
which strains from oysters were more clonal. The absence of ctxA/B and tcp genes
associated with pandemic strains suggests limited virulence potential and human health
risks associated with these strains. However, some Apalachicola Bay isolates were
genetically more similar to pandemic isolates from infections than to other strains from
environmental reservoirs. In an era of warming coastal waters and increasing vibriosis
incidence, this study provides initial data for understanding the complex and dynamic
factors affecting the distribution of toxigenic V. cholerae in Florida oysters and shellfish
harvesting waters. Further studies are needed to support future policy decisions and
management practices to reduce or eliminate the risk of this pathogen in seafood
industry.
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Figure 3-1. Maximum-likelihood tree for MLST. Phylogenetic relationship of 25 Apalachicola Bay isolates (5 from oysters and 20 from seawater) was constructed from concatenated sequences of 15 loci, including recA, gyrB, topA, pyrH, gapA, adk, mdh, mete, pnta, purm, pyrc, toxR, ompU, hlyA, and rtxA sequences. Eleven serotype O1 (four El Tor biotype and one classical biotype), one serotype O139 and four non-O1/O139 serotype reference strains are shown, including the O75 outbreak strain (boxed). Two Tampa Bay oyster isolates are included for comparison. Isolates from Apalachicola Bay are designated as “OY”, “SW” and “SD” and were isolated from oysters, seawater and sediment respectively. “AB” and “S” were Apalachicola Bay and the site collected from respectively.
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O1/0139
Non-O1/O139
Tampa Bay Oysters
Apalachicola Bay Oysters
Apalachicola Bay Oysters
Non-O1/O139
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Table 3-1. Summary of V. cholerae strains used in this study
Strains Name Geographical
Origina Source Year
Serogroup (Biotype)b
1587 Lima, Peru Clinical 1994 O12
AM-19226 Bangladesh Clinical 2001 O39
MO10 Madras, India Clinical 1992 O139
MZO-2 Bangladesh Clinical 2001 O14
MZO-3 Bangladesh Clinical 2001 O37
V51 USA Clinical 1987 O141
IEC224 Brazil Clinical 1994 O1 (EI Tor)
LMA3984-4 Brazil River water 2007 O1 (EI Tor)
M66-2 Indonesia Clinical 1937 O1
MJ-1236 Bangladesh Clinical 1994 O1 (EI Tor)
MS6 Thailand Clinical 2007 O1 (EI Tor)
N16961 Bangladesh Clinical 1975 O1 (EI Tor)
2010EL-1786 Haiti Clinical 2010 O1 (EI Tor)
O395 India Clinical 1965 O1
2012EL-2176 Haiti Clinical 2012 O1 (EI Tor)
I-1300 Russia Clinical 1999 O1
457 USA Clinical 2011 O75
30OY-Tampa Bay-2012 Upper Bay, Tampa Oyster 2012 ND
24OY-Tampa Bay-2012 Upper Bay, Tampa Oyster 2012 ND
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Table 3-1. Continued
Strain Name Geographical Origina Source Year Serogroup (Biotype)
12SW-AB-S5-2012 Shore Town Boat Ramp,
Apalachicola Bay Seawater 2012 NDb
21SW-AB-S5-2012 Shore Town Boat Ramp,
Apalachicola Bay Seawater 2012 ND
15SW-AB-S5-2012 Shore Town Boat Ramp,
Apalachicola Bay Seawater 2012 ND
14SG-AB-S5-2012 Shore Town Boat Ramp,
Apalachicola Bay Sea grass 2012 ND
45SW-AB-S4-2013 Shore DACS, Apalachicola
Bay Seawater 2013 ND
46OY-AB-S1-2013 Bay-Ward’s Lease, Apalachicola Bay
Oyster 2013 ND
52SW-AB-S15-2013 Bay- 98 Bridge N, Apalachicola Bay
Seawater 2013 ND
53SW-AB-S3-2013 Bay-Cat Point, Apalachicola
Bay Seawater 2013 ND
47OY-AB-S3-2013 Bay-Cat Point, Apalachicola
Bay Oyster 2013 ND
48OY-AB-S13-2013 Bay- Dry bar, Apalachicola
Bay Oyster 2013 ND
49OY-AB-S1-2013 Bay-Ward’s Lease, Apalachicola Bay
Oyster 2013 ND
50SW-AB-S4-2013 Shore DACS, Apalachicola
Bay Seawater 2013 ND
51SW-AB-S5-2013 Shore Town Boat Ramp,
Apalachicola Bay Seawater 2013 ND
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Table 3-1. Continued
Strains Name Geographical Origin Source Year Serogroup (Biotype)
13SW-AB-S5-2012 Shore Town Boat Ramp,
Apalachicola Bay Seawater 2012 ND
31OY-AB-S15-2013 Bay- 98 Bridge N, Apalachicola Bay
Oyster 2013 ND
32SW-AB-S3-2013 Bay-Cat Point, Apalachicola
Bay Seawater 2013 ND
33SW-AB-S6-2014 Shore Waterman Park,
Apalachicola Bay Seawater 2014 ND
34SW-AB-S6-2014 Shore Waterman Park,
Apalachicola Bay Seawater 2014 ND
35SW-AB-S5-2013 Shore Town Boat Ramp,
Apalachicola Bay Seawater 2013 ND
36SW-AB-S18-2013 East Point, Apalachicola Bay Seawater 2013 ND
22SW-AB-S4-2012 Shore DACS, Apalachicola
Bay Seawater 2012 ND
37SW-AB-S4-2012 Shore DACS, Apalachicola
Bay Seawater 2012 ND
38SG-AB-S13-2012 Bay- Dry bar, Apalachicola
Bay Sea grass 2013 ND
1SG-AB-S5-2012 Shore Town Boat Ramp,
Apalachicola Bay Sea grass 2012 ND
39SW-AB-S4-2012 Shore DACS, Apalachicola
Bay Seawater 2012 ND
40SW-AB-S5-2012 Shore Town Boat Ramp,
Apalachicola Bay Seawater 2012 ND
41SW-AB-S4-2012 Shore DACS, Apalachicola
Bay Seawater 2012 ND
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Table 3-1. Continued
Strains Name Geographical Origin Source Year Serogroup (Biotype)
43SW-AB-S5-2013 Shore Town Boat Ramp,
Apalachicola Bay Seawater 2013 ND
44SW-AB-S4-2012 Shore DACS, Apalachicola
Bay Seawater 2012 ND
42SW-AB-S4-2012 Shore DACS, Apalachicola
Bay Seawater 2012 ND
a) Strains from Apalachicola Bay were derived from this study. Strains from Tampa Bay were kindly provided by Dr. V. Harwood. Information for remaining were derived from descriptions in GenBank.
b) ND = Not Determined
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Table 3-2. Virulence potential of V. cholerae strains isolated from environmental samples in Florida.
Virulence Genesa
toxR rtxA hlyA opmU ctxA/ctxB tcpA
94% 88% 94% 100% 0% 0% a) The presence or absence of virulence genes (toxR, rtxA, ompU, ctxA/ctxB, and tcpA) was
determined from whole genomic sequencing as described in the text.
78
Table 3-3. Antibiotic susceptibility test result
a) Antibiotics are grouped based up the mechanism for antimicrobial activity. b) Resistant, intermediate and sensitive percentages refer to the percentage of strains showing a
particular zone of inhibition as defined in text.
Antibiotic Mechanisma Resistantb Intermediate Sensitive
Cell Wall Synthesis Inhibitors
Ceftriaxone 0% 0% 100% Amoxicillin-Clavulanic Acid 14% 26% 60% Cephalothin 9% 0% 91%
Protein Synthesis Inhibitors
Tetracycline 11% 0% 89% Kanamycin 3% 3% 94% Streptomycin 11% 32% 57% Amikacin 11% 11% 78%
DNA Synthesis Inhibitors
Nalidixic Acid 0% 0% 100% Ciprofloxacin 0% 0% 100%
Folic Acid Synthesis Inhibitors
Sulfamethoxazole-Trimethoprim
0% 0% 100%
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CHAPTER 4 THE ANTIVIBRIOCIDAL POTENTIAL OF CHITOSAN MICROPARTICLE IN
SEAWATER AND OYSTERS
Introduction
Vibrio species cause a significant proportion of human infections associated with
consumption of raw or undercooked shellfish, particularly raw oysters (Newton et al.,
2012). The primary pathogens in the US are V. vulnificus and V. parahaemolyticus;
however, in 2011 the first US outbreak of cholera in recent history was attributed to the
consumption of oysters contaminated by V. cholerae O75 (Onifade et al., 2011). Unlike
other foodborne pathogens associated with seafood, Vibrio spp. occur naturally in
estuarine environments, and their abundance is seasonal (Motes et al., 1998; Tamplin
et al., 1982). During warmer months (water temperature > 20C), nearly all oysters
harvested from US Gulf Coast waters harbor V. vulnificus and/or V. parahaemolyticus,
with the highest densities periodically exceeding 104 MPN/g (Cook et al., 2002b).
Despite extensive efforts employing HACCP approaches and improved sanitation by the
seafood industry, incidence of seafood-associated cases continues to escalate,
particularly during summer months, perhaps as a consequence of increasing global
water temperatures (Paz et al., 2007b). Annual reports of Vibrio-related disease per
100,000 population increased from 0.09 to 0.28 in COVIS and from 0.15 to 0.42 in
FoodNet in the last 15 years (Newton et al., 2012).
In response to the Vibrio risk assessment, FDA implemented guidance regarding
post-harvest processing (PHP) of Gulf Coast oysters harvested during summer months
Published in Fang, L., Wolmarans, B., Kang, M.Y., Jeong, K.C., Wright, A.C., 2015. Application of Chitosan Microparticles for Reduction of Vibrio Species in Seawater and Live Oysters. Appl Environ Microbiol 81, 640-647.
80
(Feldhusen, 2000). Established PHP methods to reduce Vibrio numbers in oysters
include thermal, gamma irradiation, freezing, and high hydrostatic pressure treatments
(Andrews et al., 2000; Kural et al., 2008; Mahmoud, 2009). Although effective in
reducing pathogen loads to non-detectable levels (<30 MPN/g), these approved PHPs
generally kill the shellfish and may lead to undesirable changes in shelf life, color, flavor
and texture (Cruz-Romero et al., 2007). Furthermore, substantial demand for live
oysters is apparent (Muth et al., 2013). Ice immersion (Quevedo et al., 2005),
depuration (immersion in recirculating, sanitized seawater) (Tamplin and Capers, 1992),
and relaying (transport to offshore high salinity/low Vibrio sites) (Motes and DePaola,
1996) methods maintain product integrity but are less effective. Therefore, development
of novel PHP alternatives is vital to the seafood industry for alleviating issues of
pathogenic Vibrio spp. in raw oysters.
Chitin is the second most abundant natural biopolymer after cellulose and is a
component of various marine organisms, such as the shells of crab, lobster and shrimp
(Kurita, 2006; Tharanathan and Kittur, 2003). Because of the low biodegradation of
chitin, a large amount of crustacean exoskeleton waste accumulates after seafood
processing, accounting for 50-90% total solid waste landing in the US (Knorr, 1984; Tan
et al., 1996). In this respect, commercial application of chitin derivatives from
inexpensive seafood refuse is both an environmentally acceptable use of an oceanic
resource and an economically feasible solution for waste disposal. In recent decades,
chitosan has attracted a great deal of attention with a wide range of applications
(Prashanth and Tharanathan, 2007). Chitosan is a deacetylated derivative of chitin, and
chitosan derived from shrimp was recently approved for GRAS status as a food additive
81
by the US Food and Drug Administration (Alam et al., 2006). In addition, Japan and
Korea have approved chitosan as a food additive since 1983 and 1995, respectively
(Chistoserdova, 2010; Taylor, 2011). Chitosan-mediated systems can significantly
improve bioavailability of drug delivery and are categorized as nanoparticle,
microparticle, or macro delivery systems (Hossain et al., 2007; Kurita, 2006; Pal et al.,
2013). Furthermore, the antimicrobial activity of chitosan has been well demonstrated
for both Gram-positive and Gram-negative pathogens as well as for food spoilage
bacteria (Chen et al., 2002; Liu et al., 2004).
Chitosan microparticles (CM) are derived from chitosan with minor cross-linking
modification, and a recent study showed application of CM as a feed additive resulted in
reduced shedding of E. coli O157:H7 in cattle (Jeong et al., 2011). Chitosan was
previously shown to be effective against V. vulnificus in vitro and in mice (Lee et al.,
2009), but the effects of CM against pathogenic Vibrio spp. and possible applications to
live oysters have not been studied. Therefore, the objective of the study of this chapter
was to investigate the effects of CM treatment on pathogenic Vibrio spp., and evaluate
the potential feasibility of CM as a PHP treatment for live oysters.
Materials and Methods
Bacterial Strains and Inoculum Preparation
Three clinical strains of Vibrio spp. used in this study included V. vulnificus
CMCP6 (encapsulated biotype 1 with “C” genotype commonly found in clinical strains)
(Kim et al., 2003), V. parahaemolyticus TX2103 (Serotype O3:K6) (DePaola et al.,
2003b), and V. cholerae 139 classical O1 (Johnson et al., 1994), and were provided by
Drs. P. Gulig, A. DePola and J. Johnson, respectively. Strains were stored as -80°C
frozen stock cultures in Luria-Bertani broth with NaCl (LBN: 1.0% tryptone, 0.5% yeast
82
extract, 1.0% NaCl in deionized water, pH 8.4) in 50% glycerol. For each experiment,
bacteria were retrieved from frozen stock onto LBN agar (LA) and individual colonies
were used inoculate LBN for preparation of liquid inocula. All media were from Difco
(Sparks, MD), and unless otherwise stated; all other reagents were from Sigma Aldrich
(St. Louis, MO).
Chitosan Microparticles (CM) Preparation
Preparation of CM followed a previously described protocol (Jeong et al., 2011;
van der Lubben et al., 2001). Briefly, chitosan was purchased from Sigma-Aldrich
(448869-250G), and a 1% (w/v) chitosan solution was prepared in 2% (v/v) acetic acid
with 1% (w/v) Tween®80. After addition of 2 ml of 10% (w/v) aqueous sodium sulfate,
the chitosan solution was stirred and sonicated for 20 min to generate microparticles.
The chitosan microparticles were collected by centrifugation at 6000 rpm for 10 min,
washed with deionized water three times, and freeze-dried for further use.
In vitro Evaluation of Effects of CM on Growth of Vibrio spp.
To evaluate growth inhibition, bacteria were streaked for isolation to LA from
frozen stock cultures for each experiment, and plates were incubated at 37°C overnight.
Inocula of each species were picked from LA plates, and were cultured separately
overnight (18-23 h) in LBN broth at 37C with shaking (100 rpm). The overnight cultures
were serially diluted in PBS, enumerated by absorbance at 600 nm compared to a
standard curve, and diluted in LBN (40 ml) at pH 7.4 to ca. 104 log CFU/ml. Each strain
was incubated in LBN with different CM concentrations (0.0, 0.1, 0.3, and 0.5%, wt/vol)
in 250 ml Erlenmeyer flasks with shaking at 37°C, and CFU/ml determined by plating on
LA plates at 0, 3, 6, 9 and 12 h post-inoculation.
83
For survival studies, sterile artificial seawater (ASW, Instant Ocean Sea Salt,
Aquarium Systems, Blacksburg, VA) was prepared in DI water at 20 ppt, pH=7.4.
Inocula were prepared as described above except at levels of ca. 107 log CFU/ml in
flasks of ASW (40 ml) with different CM concentrations (0, 0.1, 0.3, and 0.5%), and
incubated at 37, 25 or 4°C without shaking. Survival of each species was determined by
plate counts on LA after 24 and 48 h incubation. All in vitro results were reported as
mean log CFU/ml ± standard deviation from three independent experiments with three
flask replicates for each experiment.
Effects of CM Treatment on Survival of V. vulnificus and V. parahaemolyticus in Artificially Inoculated Oysters
Live oysters (C. virginica) were obtained from a local seafood market,
transported in coolers on ice packs, and brought to the laboratory within 2 h. Oysters
were acclimated in air at room temperature (25 ± 1C) for 30 min in order to avoid
temperature shock and then cleaned under tap water to remove any dirt or debris.
Subsequently, oysters (up to 30 oysters/tank) were placed in 30-gal tanks (Nalgene
heavy duty rectangular HDPE tank with cover 24 x 18 x 18”) in 20 L of ASW (20 ppt,
pH= 7.4) for 24 h acclimation at room temperature (25 ± 1C) using two pumps with
charcoal filtration (Tetra Whisper Internal Power Filter). Following acclimation in ASW,
tetracycline was used as previously described (Srivastava et al., 2009) to reduce the
indigenous Vibrio levels prior to experimental inoculation. Oysters (n=6) were
transferred to smaller tanks (Nalgene HDPE pans 21 x 17 x 5”), containing 6 L of ASW
with tetracycline (10 µg/ml final concentration) and incubated at room temperature
without filtration for 24 h. Exposure to antibiotics was discontinued by transferring
84
oysters to fresh ASW in new 6 L tanks, followed by incubation for 24 h with charcoal
filtration to remove residual tetracycline.
Oysters were artificially inoculated by addition of V. vulnificus or
V. parahaemolyticus to the ASW (ca. 106 CFU/ml) in fresh tanks, covered with foil, and
incubated without filtration for 24 h. Oysters were then transferred to a new tank
containing 6 L of ASW and various concentrations (0.0, 0.1, 0.3, and 0.5% w/v) of CM
and individually evaluated for survival of V. vulnificus or V. parahaemolyticus after 0, 24
and 48 h exposure to CM. Oysters were removed from tanks, transferred to a biological
safety cabinet, shucked under sterile conditions using shucking knives that had been
rinsed with ethanol (70%) and flamed. Oyster meats were collected aseptically in 50 ml
sterile conical tubes, weighed, and homogenized for 30 s with an equal volume of PBS
using a sterile mini blender (Seward, Stomacher® 80 Biomaster, Lab System) to prepare
1:2 dilution sample suspensions. Serial 10-fold PBS dilutions were used to enumerate
Vibrio spp. by spread plate on selective agars, namely modified cellobiose-polymyxinB-
colistin (mCPC) agar for V. vulnificus (Warner and Oliver, 2007) or on Vibrio
CHROMagar™ (CHROMagar Microbiology, Paris, France) for V. parahaemolyticus.
Presumptive V. vulnificus (yellow colonies on mCPC) or V. parahaemolyticus (mauve
colonies on Vibrio CHROMagar™) were counted and reported as log CFU/g. All
experiments were independently conducted three times using three oyster replicates for
each experimental condition and time point for a total of nine oysters per treatment.
Effects of CM Treatment on Survival of Indigenous Vibrio spp. in Oysters
Market oysters were obtained in the summer to ensure high levels of Vibrio and
acclimated to laboratory conditions in holding tanks as described above. Oysters were
then transferred to experimental tanks and treated with various concentrations (0, 0.1,
85
0.3, and 0.5%) of CM as above. Oysters were individually evaluated for survival of
V. vulnificus, V. parahaemolyticus, V. cholerae, and heterotrophic aerobic bacteria after
0, 24 and 48 h exposure to CM by plate counts on mCPC agar, Vibrio CHROMagar™,
thiosulfate-citrate-bile salts-sucrose (TCBS) agar, and LA, respectively, as described
previously. Typical colonies were assessed by PCR in trial 3, using species-specific
primers for V. vulnificus (Warner and Oliver, 2008), V. parahaemolyticus (Bej et al.,
1999), and V. cholerae (Chun et al., 1999). Results represent three independent
experiments using three oysters per experimental condition and time point in the first
and second trials and six oysters per sample in trial 3 for a total of 12 oysters per
treatment.
Statistical Analysis
Results of microbiological tests were log transformed for statistical analysis.
Analyses of variance (ANOVA) were performed to test the null hypotheses that there
were no effects of chitosan treatment on CFU/g levels of bacterial populations in
samples. If a null hypothesis was rejected at the 0.05 level, a Tukey’s multiple mean
comparison test was used to identify differences in treatments. Another ANOVA was
also performed in all the in vivo tests based on the differences between day 1 and day 2
and pretreatment. Student t tests were then used to determine if mean differences were
significantly different from zero. Analysis was run using JMP pro 11 (SAS, Cary, NC).
Results
Chitosan Inhibits Growth of Vibrio spp. in Broth Culture
A range (0.1, 0.3, and 0.5%) of CM concentrations was evaluated for inhibition of
growth of the three pathogenic Vibrio spp. under optimal culture conditions. Exposure to
0.5% CM resulted in growth cessation, and levels of all three Vibrio spp. were
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significantly (p<0.0001) reduced compared to untreated control samples (0.0% CM) and
became non-detectable at 3 h post-treatment (Figure 4-1). Similar results were obtained
with 0.1 and 0.3% CM for V. vulnificus and with 0.3% for V. parahaemolyticus
(p<0.0001). However, V. cholerae showed more gradual inhibition at 0.3% CM
compared to control samples, and no inhibition was observed for V. cholerae and
V. parahaemolyticus in 0.1% CM. Reductions of V. cholerae were significantly less than
were observed for V. vulnificus or V. parahaemolyticus (p<0.02) for 0.3% CM, and
reductions of V. parahaemolyticus and V. cholerae (p<0.03) were significantly less than
those for V. vulnificus at 0.1% CM. Thus, the efficacy of CM to eliminate these
pathogenic Vibrios spp. varied among species.
Effects of CM on Survival of Vibrio spp. in ASW
The effects of CM on survival of pathogenic Vibrio spp. under simplified estuarine
conditions, namely ASW at 20 ppt pH=7.4, were investigated using high levels of
bacteria (ca. 107 CFU/ml). As shown in Figure 4-2, dramatic reductions (>7 mean log
CFU/ml) were observed for all three species in comparison to untreated control cultures
following exposure to 0.5% CM at 37C (p<0.001). V. vulnificus was the most sensitive
of the species to the deleterious effects of CM and was no longer detected in either
0.3% or 0.5% CM by 24 h at all incubation temperatures examined. V. vulnificus also
became non-detectable even in 0.1% CM by 48 h at 25C and 37C but not at 4C.
Thus, sensitivity to CM also varied with temperature and appeared to increase with
increasing temperature, as reduction to non-detectable levels was not achieved at 4 and
25C for V. parahaemolyticus and V. cholerae. However, significant effects (p<0.05) of
all CM concentrations, as compared to untreated controls, were evident for all
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temperatures examined for V. vulnificus and V. parahaemolyticus by 48 h exposure.
However, 0.1% CM did not result in any significant inhibition compared to non-treated
controls for V. cholerae at any temperature over the entire experiment. Results showed
that sensitivity to CM in ASW was consistent with growth inhibition in broth culture in
that the same general trend for species sensitivity was observed with V. vulnificus >
V. parahaemolyticus > V. cholerae.
Effect of CM Treatment on Survival of Vibrio spp. in Artificially Inoculated Live Oysters
Live oyster experiments were conducted for V. vulnificus or V. parahaemolyticus,
as these species are the targets of oyster PHP in the US seafood industry. Artificial
inoculations were achieved by pretreating oysters with tetracycline to remove native
Vibrio populations and subsequently inoculating the ASW with Vibrios, which allowed
the oysters to internalize these bacteria via filter-feeding, as previously described
(Srivastava et al., 2009). Survival of Vibrio spp. in individual oysters (n=3) was
evaluated in three independent experiments after 0, 24, and 48 h exposure to CM by
plate counts on selective agars (mCPC and Vibrio CHROMagar™, respectively). PCR
confirmation was not performed as pre-screening revealed no background Vibrio levels
after tetracycline treatment (data not shown).
The pretreatment inocula in oyster meats averaged 4.6 log CFU/g (Figure 3). All
three trials showed significant (p<0.001) reductions of V. vulnificus in oyster tissues
after 24 h exposure for all concentrations of CM, as compared to untreated controls,
and levels continued to decline at 48 h. Reductions for V. vulnificus averaged >4.0 log
CFU/g by 48 h from three independent experiments following treatment with 0.5% CM.
Even at 0.1% CM, decreases in V. vulnificus levels were ca. 2.0 log CFU/g and were
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significantly (p<0.0001) different from levels observed in untreated control oysters.
Significant reductions (p<0.02) were also obtained for V. parahaemolyticus compared to
untreated controls treated for both 0.3% and 0.5% CM, resulting in a 2.2 and 3.3 log
CFU/g reductions, respectively, after two days. No significant differences in
V. parahaemolyticus levels were observed for 0.1% CM treatment compared to
untreated oysters. Thus, results were consistent with the in vitro experiments in that
both species were sensitive to CM, but V. parahaemolyticus response was somewhat
attenuated compared to V. vulnificus.
Effect of CM Treatment on Survival of Indigenous Vibrios in Live Oysters
To further examine the antimicrobial effect of CM, fresh summer oysters with
indigenous populations of Vibrio spp. were subjected to CM treatment in three
independent experiments. V. vulnificus and V. parahaemolyticus mean log CFU/g were
determined by plate counts on selective agars. PCR confirmation was performed only in
experiment 3 and showed >80% agreement with presumptive identifications (data not
shown), which is consistent with the reported accuracy of Vibrio identification on these
agars (Di Pinto et al., 2011; Warner and Oliver, 2007). V. cholerae was not detected in
these oysters. Heterotrophic aerobic bacteria were also enumerated in oyster
homogenates by standard plate counts on nonselective LA.
As expected, initial concentrations of V. vulnificus (Table 4-1) and
V. parahaemolyticus (Table 4-2) before treatment showed greater variation relative to
artificially inoculated oysters, presumably due to distinct conditions at harvest or during
storage. Although results were consistent among the three independent experiments,
data were not averaged due to variation in initial levels. Significant reductions (p<0.05)
in V. vulnificus levels compared to untreated controls were observed for all CM
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concentrations by 24 h, and levels continued to decline after 48 h post-treatment.
Exposure to 0.5% CM was more effective than the lower concentrations, and reductions
on day 2 post-treatment compared to untreated controls ranged from 1.9 to 3.9 log
CFU/g for V. vulnificus and from 1.9 to 2.6 log CFU/g for V. parahaemolyticus over the
three experiments. Furthermore, greater vibriocidal activity was observed for treated
samples compared to initial levels and reached 4.0 and 4.7 log CFU/g reductions for
V. vulnificus and V. parahaemolyticus, respectively, in some experiments. Heterotrophic
aerobic bacteria also declined following CM treatment compared to untreated controls,
and reductions ranged from 1.4 to 3.4 log CFU/g (Table 4-3). Results may have been
confounded by differences in initial levels among the three trials, but results clearly
demonstrate the significant effects of CM on Vibrio spp. in live oysters. It should also be
noted that with 0.5% CM, post-treatment levels of V. vulnificus post-treatment were < 30
CFU/g for all experiments (ISSC criteria for validation of a PHP), and levels of
V. parahaemolyticus were all <100 CFU/g (criteria for the harvest) (Terzi and
Gucukoglu, 2010).
Discussion
Currently approved PHP methods effectively lower Vibrio levels but are generally
detrimental to maintaining live oyster shell stock and can be expensive (Muth et al.,
2013). Therefore, novel and more economical PHP strategies are required for
successful treatment of oysters harvested from Gulf Coast waters. This study
demonstrated that chitosan in the form of microparticles has strong anti-Vibrio activity
on both the growth of these bacteria in culture and on their survival in seawater and
oysters. In fact, in vitro growth was completely halted, and bacteria were non-detectable
by 3 h exposure to 0.5% CM. Similar treatment in seawater also reduced levels of all
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three species by >7.0 log CFU/ml within 48 h or less at 37C. Anti-Vibrio activity was
dependent upon species, CM concentration, temperature, and exposure time. Among
the three species, V. vulnificus exhibited the greatest sensitivity, and the response of all
species was attenuated at 4C, suggesting that increased temperature serves to
augment the negative effects of CM on survival. In contrast, all species gradually
declined somewhat at 4C without CM treatment compared to untreated samples at
higher temperatures, suggesting a shift to a viable but VBNC state previously described
for these species as a response to lower temperatures (Oliver et al., 1991). Induction of
VBNC as a consequence of chitosan treatment was not investigated, but prior studies
demonstrated rapid loss of membrane integrity and viability in E. coli under similar
conditions of CM exposure (Jeon et al., 2014). CM treatment did not appear to induce
VBNC in Vibrios at low temperature, as bacteria were actually less sensitive to
treatment at lower temperature.
CM treatment was highly effective in reducing Vibrio levels in live oysters for
either inoculated or autochthonous populations of V. vulnificus and
V. parahaemolyticus. Results suggested that the mitigation of Vibrio spp. in oysters
harboring natural populations was somewhat less efficacious than artificially infected
ones. However, these differences may reflect the variability of pre-treatment bacterial
levels in naturally infected oysters, as samples with higher initial concentrations
generally exhibited greater reductions following CM treatment. Discrepancies in results
from natural vs. artificial populations may also reflect greater heterogeneity of natural
bacterial populations, as it is plausible that various strains are more resistant to CM
exposure. Alternatively, the physiological state of the natural compared to “artificial”
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Vibrio populations may provide pre-adaptation for resisting CM (Kaspar et al., 1990;
Richards, 1988; Tamplin and Capers, 1992).
Previous examination of the anti-Vibrio vulnificus activity for water soluble
fractions of chitosan (10,000 and 1,000 kDa) found greater activity with the higher
molecular weight preparation, which required 1-10 mg/ml (0.1-1.0%) for in vitro growth
inhibition. Furthermore, co-administration of 0.1-0.5 mg of chitosan with V. vulnificus
infections in mice significantly increased survival and decreased dissemination in mice
(Lee et al., 2009). Chitosan contains positively charged molecules that bind to
negatively charged structures on cell surfaces, and subsequently induce the leakage of
intracellular material from bacterial cells (Jeon et al., 2014; Liu et al., 2004; Raafat et al.,
2008). Exposure to water-soluble fractions of chitin has been shown to induce
competence in V. cholerae and V. vulnificus for uptake of DNA and is used in molecular
biology for transformation experiments (Gulig et al., 2009; Meibom et al., 2005). Metal-
binding capacity of chitosan was also considered to block pathogens by disrupting
protein synthesis of virulence factors such as cytolysin, elastase, metalloproteinase, etc.
(Lee et al., 2009; Rabea et al., 2003; Schlievert, 2007). In addition, soluble chitosan was
found to inhibit Vibrio cell-to-cell communication through the suppression of intracellular
reactive oxygen species generation, which is known to induce cell death (Lee et al.,
2009).
Chitosan microparticles were used in the present study, as Jeon et al. (2014)
demonstrated significant antimicrobial activity at pH 7-8, which coincides with optimum
pH levels for both Vibrios and oysters. They suggested that hydrophobic interactions
contribute to the mechanism of CM antimicrobial activity above neutral pH, and binding
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of CM to outer membrane protein OmpA and to LPS in E. coli O157:H7 was shown
disrupt membranes leading to cell death. The mechanism of CM activity against Vibrio
spp. has not been investigated and is likely to be complex, due to the diversity of these
species. Significant species differences in sensitivity to CM were reported for Vibrio spp.
based on strains tested, and investigations into the basis for these differences may
provide better understanding of mechanisms of activity. It is plausible that differences
among these species in the composition of capsular polysaccharide, LPS, or outer
membrane proteins, contribute to altered surface charge, hydrophobicity, binding
properties, etc., that correspond to specie-specific differences in CM sensitivity.
Although validation of CM treatment as an oyster PHP will require more
exhaustive criteria than those presented herein, these results demonstrated that CM
treatment will likely meet the standards for oyster PHP validation. PHP validation
standards described by ISSC (Terzi and Gucukoglu, 2010) require geometric mean
reduction of >3.52 log MPN/g from an initial level of ca. 4.0 log MPN/g to achieve <30
MPN/g following PHP compared to initial levels for three independent trials using 10
replicates of 12 pooled oysters for each trial. The Canadian Food Inspection Agency
(CFIA) recently added total end-product guidelines for raw oysters, limiting
V. parahaemolyticus counts to no more than 1 in 5 samples exceeding 100 total
V. parahaemolyticus per gram and no single sample exceeding 10,000 total
V. parahaemolyticus/g (Arbuckle, 2013). In this study, observed reductions, as
determined by plate count for V. vulnificus in artificially inoculated oysters and in one of
three trials using natural populations, attainted the reductions that met the PHP
validation criteria, and all experiments reached <30 CFU/g by day 2 of treatment.
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Criteria for V. parahaemolyticus based on CFIA were also realized. These results
substantiate the potential for application of CM against Vibrio spp. in oyster PHP,
particularly for reduction of V. vulnificus. This study investigated live oysters, but
applications may also be effective as a hurdle technology for the processed product to
be used in combination with other PHP and for other seafood. Further studies will be
needed to optimize the effects of CM treatments and to determine sensitivity of the
different species and of strains within each species, as well as to explore the capacity
for scaling up the process and to investigate possible changes in the sensory attributes
and shelf life of the resulting product. Validation of CM as a PHP for live oysters or other
shellfish should provide the first available treatment that effectively eliminates potentially
pathogenic Vibrios, while maintaining the viability of the molluscan shellfish.
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Figure 4-1. Effects of CM on growth of Vibrio spp. in broth culture. Vibrio spp. were cultured in LB with a range of CM concentration (0, 0.1, 0.3, and 0.5%) at
37C with shaking as described in Materials and Methods section, and bacterial growth was evaluated by plate count (mean log CFU/ml). Results were the mean of three independent experiments; standard deviations were indicated by error bars.
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Figure 4-2. Effects of CM on survival of Vibrio spp. in ASW. Vibrio spp. were incubated in ASW (20 ppt, pH 7.4) with a
range of CM concentration (0, 0.1, 0.3, and 0.5%) at either 37C, ca. 25C (RT), or 4C. Bacterial survival (mean log CFU/ml) was calculated from three independent experiments; standard deviations are indicated by error bars.
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Figure 4-3. Effects of CM on survival of Vibrio spp. in artificially inoculated oysters.
Oysters (n=3) were inoculated with Vibrio spp. by suspension of bacteria in ASW (20 ppt, pH 7.4, RT), as described in Materials and Methods section. Inoculated oysters were exposed to different concentrations of CM (0, 0.1, 0.3, and 0.5%). Vibrio levels (mean log CFU/g ± standard deviation) in oysters were determined from three independent experiments at 0, 24, and 48 h post treatment on selective agars.
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Table 4-1. Effects of CM on indigenous V. vulnificus in oysters
Experiment CM Treatment
(%)
V. vulnificus levels (mean log CFU/g ± SD)a
Pre-Treatment Day 1 Day 2
1 0.0 Control 4.74 ± 0.16 4.96 ± 0.52 4.64 ± 0.80
0.1 3.34 ± 0.29b
2.69 ± 0.33
0.3 3.00 ± 0.57b
1.60 ± 1.39b
0.5 2.51 ± 0.64b
0.70 ± 1.22c
2 0.0 Control 3.83 ± 0.15 3.80 ± 0.50 3.74 ± 0.13
0.1 2.99 ± 0.40 2.13 ± 0.22
0.3 2.68 ± 0.21b
1.24 ± 1.07b
0.5 2.05 ± 0.21b
0.53 ± 0.92b
3 0.0 Control 4.01 ± 0.38 3.69 ± 0.40 3.02 ± 0.23
0.1 2.74 ± 0.71 1.19 ± 1.00b
0.3 1.34 ± 1.07b
0.90 ± 1.07b
0.5 1.29 ± 1.43b
1.08 ± 1.19b
a Mean log CFU/g ± standard deviation (SD) based on plate counts on mCPC from three independent experiments with three oysters in the first two experiments and six oysters in the third experiment for a total of twelve oysters for each experimental condition and time point.
b Reduction of V. vulnificus from initial level is <3.52 mean log CFU/g but is significantly different from 0.0% control samples as determined by two-tailed, one way ANOVA (p < 0.05).
c Reduction of V. vulnificus from initial level is >3.52 mean log CFU/g and is significantly different from 0.0% control samples, as determined by two-tailed, one way ANOVA (p < 0.05).
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Table 4-2. Effects of CM on indigenous V. parahaemolyticus in oysters
Experiment CM Treatment
(%)
V. parahaemolyticus levels (mean log CFU/g ± SD)a
Pre-Treatment Day 1 Day 2
1 0.0 Control 5.98 ± 0.44 5.63 ± 0.10 4.32 ± 0.66
0.1 3.94 ± 0.77b
2.88 ± 0.40
0.3 3.93 ± 0.65b
2.59 ± 0.38
0.5 3.26 ± 0.44b
1.72 ± 0.46c
2 0.0 Control 3.47 ± 0.46 3.48 ± 0.54 3.46 ± 0.37
0.1 3.33 ± 0.58 2.33 ± 0.55
0.3 2.53 ± 0.67 2.07 ± 0.24
0.5 1.73 ± 0.17b
1.13 ± 0.98b
3 0.0 Control 3.13 ± 0.62 2.58 ± 1.47 2.69 ± 0.52
0.1 1.88 ± 1.15 1.56 ± 1.39
0.3 0.84 ± 0.93 0.69 ± 1.06b
0.5 0.68 ± 0.79 0.76 ± 0.84b
a Mean log CFU/g ± standard deviation (SD) based on plate counts on Vibrio CHROMagar from three independent experiments with three oysters in the first two experiments and six oysters in the third experiment for a total of twelve oysters for each experimental condition and time point.
b Reduction of V. parahaemolyticus from initial level is <3.52 mean log CFU/g but is significantly different from 0.0% control samples as determined by two-tailed, one way ANOVA (p < 0.05).
c Reduction of V. parahaemolyticus from initial level is >3.52 mean log CFU/g and is significantly different from 0.0% control samples, as determined by two-tailed, one way ANOVA (p < 0.05).
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Table 4-3. Effects of CM on heterotrophic aerobic bacteria in naturally infected oysters.
Experiment CM Treatment
(%)
Heterotrophic aerobic bacteria (mean log
CFU/g ± SD)a
Pre-Treatment Day 1 Day 2
1 0.0 Control 6.96 ± 0.13 6.91 ± 0.10 6.11 ± 0.30
0.1 4.68 ± 0.28b
4.33 ± 0.52b
0.3 4.32 ± 0.69b
3.89 ± 0.48b
0.5 4.27 ± 0.41b
2.75 ± 0.67c
2 0.0 Control 5.69 ± 0.30 5.43 ± 0.23 5.31 ± 0.21
0.1 4.28 ± 0.33b
3.89 ± 0.27b
0.3 3.72 ± 0.27b
3.01 ± 0.31b
0.5 3.11 ± 0.43b
2.16 ± 0.30b
3 0.0 Control 5.40 ± 0.47 4.83 ± 0.27 4.90 ± 0.36
0.1 3.93 ± 0.61b
3.66 ± 0.91
0.3 3.97 ± 0.39 3.41 ± 0.52b
0.5 3.63 ± 0.73b
3.49 ± 0.76b
a Mean log CFU/g ± standard deviation (SD) based on plate counts on LA from three independent experiments with three oysters in the first two experiments and six oysters in the third experiment for a total of twelve oysters for each experimental condition and time point.
b Reduction of the heterotrophic aerobic bacterial level from initial level is <3.52 mean log CFU/g but is significantly different from 0.0% control samples as determined by two-tailed, one way ANOVA (p < 0.05). c Reduction of the heterotrophic aerobic bacterial level from initial level is >3.52 mean log CFU/g and is significantly different from 0.0% control samples, as determined by two-tailed, one way ANOVA (p < 0.05).
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CHAPTER 5 ROLE OF CAPSULAR POLYSACCHARIDE IN THE ACTIVITY OF CHITOSAN FOR
VIBRIO VULNIFICUS
Introduction
Chitosan, a polysaccharide biopolymer derived from chitin, displays unique
physicochemical and biological properties that have attracted considerable interest in
various applications. Chitosan has demonstrated efficacy as an antimicrobial agent
against a wide scope of microorganisms, including gram positive and negative bacteria,
fungi, and viruses (Chirkov, 2002; Rabea et al., 2003). For example, chitosan
microparticles (CM), derived from chitosan by cross-linking, successfully reduced
E. coli O157:H7 shedding in cattle as a feed additive (Jeong et al., 2011). Furthermore,
CM also exerts strong antimicrobial activity against Vibrio spp. in seawater and oysters,
which offers promising potential for the application of CM as a PHP treatment in intact
live oysters (Fang et al., 2015). As food safety problems become more complex due to
demanding food production practices, changing dietetic habits, and increased
importation, the antimicrobial activity of chitosan meets the growing consumer demand
for natural preservatives with reduced toxicity and allergenicity. In addition to its
biodegradability and biocompatibility capacity, chitosan and its derivatives have
emerged as a new biomaterial for food preservation purposes and in pharmaceutical
systems (Rhoades and Roller, 2000; Singla and Chawla, 2001).
The commercial usage of CM in the seafood industry will require sufficient
knowledge of its anti-Vibrio activity. Although the exact mechanism(s) has not been fully
elucidated, the interaction of chitosan and its derivatives with bacterial cell surface has
been widely purported. Several studies found that the outer membrane bacterial
components, including negatively charged outer membrane proteins and phospholipids
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(Liu et al., 2004), various lipopolysaccharides (Helander et al., 2001), and amino acids
(Kumar et al., 2005) were involved in interactions that caused modified cell-wall
permeability, leading to leakage of intracellular substances and ultimately to cell death.
A recent study demonstrated that chitosan interacts with OmpA of E. coli at neutral pH
to disrupt cell membrane, suggesting that chitosan may inhibit the growth of other
bacteria in a similar manner by disputing cell surface components (Jeon et al., 2014).
In order to better understand the anti-Vibrio properties related to chitosan, this
study investigated chitosan activity using V. vulnificus as a model organism to
determine the role of capsule polysaccharide (CPS) in these interactions. CPS is a well-
studied extracellular structure that is integral to virulence of this species (Wright et al.,
2001b; Wright et al., 1990; Yoshida et al., 1985). Wild-type V. vulnificus strains show
phase variation in CPS expression that is marked by changes in colony morphology,
whereby opaque colonies (encapsulated) revert to translucent colony type (reduced
encapsulation) or vice versa (Chatzidaki-Livanis et al., 2006). Furthermore, CPS has
been shown to enhance survival of V. vulnificus in seawater and oysters, and the rate of
phase variation to the opaque morphotype greatly increases during oyster colonization,
which may explain why most oyster isolates exhibit opaque colonies (Srivastava et al.,
2009).
A previously described mutant strain (Srivastava et al., 2009), lacking CPS
expression, was examined under various conditions and compared to wile-type strains
for their responses to CM exposure. This present study may serve as a paradigm for
better understanding the complex mechanisms underlying the anti-Vibrio activity of CM
and perhaps contribute to maximizing the potential for commercial utilization.
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Materials and Methods
Bacterial Strains and Culture Conditions
V. vulnificus strains and mutants in this study are summarized in Table 5-1.
Bacteria were stored in Luria broth (LBN; 1.0% tryptone, 0.5% yeast extract, 1.0% NaCl
in deionized water, pH 8.4) with 50% glycerol at -80C. For each experiment, strains
were retrieved from the frozen stock and streaked for isolation on LB with 1.5% agar
(LA).
Effects of Chitosan Microparticles (CM) Treatment on Survival of Individual V. vulnificus Strains
To evaluate the survival of individual V. vulnificus strains, bacteria from frozen
stock cultures were streaked onto LA for isolation, and the plates were incubated at
37C overnight. Inoculum for each species was retrieved from LA plates and cultured
separately overnight (18 to 23 h) in LBN broth at 37C with shaking (100 rpm). Cultures
(1ml) were harvested by centrifugation at 8,000 x g for 3 min and re-suspended in equal
volumes of sterile artificial seawater (ASW; 20 ppt, pH 7.4) to remove nutrients. Washed
cells were separately inoculated into 20 ml ASW sterile conical tubes to prepare a
culture suspension of approximate 106 CFU/ml, and incubated with different CM
concentrations (0.0, 0.05, 0.1, and 0.3%, wt/vol) with shaking at 37C. The survival of
each strain was determined by plate counts on LA at 0, 1, and 10 days post-inoculation.
Effects of CM Treatment on Competitive Survival of V. vulnificus Strains
For competitive survival studies, inocula were prepared following the same
protocol as described above, except inocula for ASW consisted approximately equal
concentrations of both MO6-24/O and MO6-24/Δwzb. ASW cultures (20 ml) were
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incubated with shaking at 37C, and the number of CFU/ml of each strains was
determined by plate counts of opaque (MO6-24/O) vs. translucent (MO6-24/Δwzb) on
LA after 24 h incubation. The results for competitive survival studies were reported as
the mean log ratio of MO6-24/Δwzb: MO6-24/O based on CFU/ml ± standard deviation
from three independent experiments with three replicate tubes for each experiment.
Phase Variation
It is plausible that if CPS played a role in resistance to chitosan, the rate of phase
variation might be also altered by chitosan treatment. For the phase variation study, the
same protocol was followed as described for individual inoculations above except the
number of both translucent and opaque colonies was recorded for each strain on LA at
0, 1, and 10 days post CM exposure.
Statistical Analyses
Results of microbiological tests were transferred to log values for statistical
analysis. Significant differences in culture density between treated and untreated
samples were determined by student’s t-test (Excel, Microsoft, Redmond, WA).
Analyses of variance (ANOVA) were performed to test the null hypotheses that there
are no effects of tested variables on CM treatment of V. vulnificus strains. If a null
hypothesis is rejected, a student’s t-test was used to identify differences. Tests were
established at p<0.05 in respect to significant differences between means of treatments
using JMP pro (version 11) software (Cary, NC).
Results
Strain Variation in the Sensitivity of V. vulnificus to CM Activity
To investigate the role of CPS in the anti-Vibrio activity of CM, this study
evaluated the effects of different CM concentrations (0, 0.05, 0.1, 0.3% w/v) on the
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survival of two different V. vulnificus strains, namely CMCP6 (opaque, clinical wild-type)
and MO6-24/O (opaque, clinical wild-type). As shown in Figure 5-1, levels of untreated
controls did not change significantly throughout the study. The levels of both
V. vulnificus strains became non-detectable in CM with 0.3% concentration by day 1
post-treatment. Levels for 0.05% declined somewhat but were statistically the same for
both strains. It is noteworthy that CMCP6 was successfully reduced by CM at 0.1%
concentration to non-detectable level after only 1 day incubation, while numbers for
MO6-24/O leveled off to about 103 CFU/ml on day 1 and remained statistically the same
through day 10 after exposure of 0.1% CM.
Effects of CM on Survival of V. vulnificus as a Function of CPS Expression.
To investigate the role of CPS in anti-Vibrio activity of CM, the survival of
V. vulnificus MO6-24/O (clinical wild-type) was compared to V. vulnificus MO6-24/Δwzb
(mutant defective in CPS expression). The CPS mutant strain was generally more
resistant to the CM antimicrobial action than the parent encapsulated strain of MO6-
24/O or the other wild-type strain CMCP6 (Figure 5-1). Reductions of MO6-24/Δwzb
with respect to the initial level on day 1 in 0.05% and 0.1% CM were only 0.8 and 1.6
log CFU/ml, respectively, and represent significantly lower reductions compared to wild-
type (p<0.05). Treatments sustained their activity through day 10, and MO6-24/O levels
remained significantly lower than of MO6-24/Δwzb (p<0.03).
Effects of CM Treatment on Competitive Survival of V. vulnificus Strains.
To further examine the role of CPS in the efficacy of CM activity on V. vulnificus,
encapsulated wild type strain MO6-24/O and its CPS mutant MO6-24/Δwzb were co-
inoculated into different CM concentrations (0, 0.05, 0.1, and 0.3%), and their
responses were evaluated with respect to the persistence of opaque (wild-type) vs.
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translucent (mutant) colonies in co-cultures. Both wild type and mutant strains were
completely eliminated in exposure to 0.3% CM, which was consistent with the
observations above (Figure 5-2). Competitive indices revealed that the distribution of
mutant translucent colonies was significantly higher than wild-type opaque following 1
day exposure to lower concentrations (0.1% and 0.05%) of CM at 37C after 24 hours
(p<0.001).
Effects of CM Treatment on CPS Phase Variation of V. vulnificus Strains.
In the study above using individual inocula, phenotypic switching (opaque to
translucent) was observed during CM treatment. This finding indicated that changes in
the rate of phase variation may also occur in response to CM activity. To determine the
role of phase variation in the susceptibility of V. vulnificus to CM treatment, changes in
colony morphology were recorded following the same protocol as described for
individually inoculated strains. As expected, the deletion mutant strain MO6-24/Δwzb
maintained its translucent phenotype throughout, and no switching to opaque
morphology was observed during the study (Figure 5-3). Furthermore, reductions were
similar to those observed in prior experiments with this strain. In addition, no phase
variation or significant reductions were observed on day 1 for wild-type strains in
untreated controls; however, both wild type strains showed significant switching from
opaque to translucent by day 10 without chitosan treatment, while overall levels of
bacteria remained unchanged in these strains. The translucent morphology was
observed in 6 or 4% of population for CMCP6 and MO6-24/O, respectively, for
untreated controls.
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On the other hand, both wild-type encapsulated strains showed reductions in
overall levels and altered colony morphology at days 1 and 10 following CM treatment
(Figure 5-3). In agreement with previous experiments both strains were no longer
detectable in 0.3% CM, while MO6-24/O was significantly reduced (2.5 log CFU/ml
compared to initial levels) and CMCP6 was no longer culturable in 0.1% CM at day 1.
Furthermore, overall populations declined by 4.5 and 2.0 log CFU/ml for CMCP6 and
MO6-24/O, respectively, following 0.05% CM treatment. Both strains showed opaque to
translucent variation in 0.05% CM; however, overall proportions of O:T ratios differed,
primarily due to differences in levels of opaque colonies. Interestingly, both strains
showed 0.8 log CFU/ml for translucent colonies under these conditions, while levels of
opaque populations were 4.9 log CFU/ml for MO6/O as compared to 2.4 log CFU/ml for
CMCP6. These differences were further exacerbated by extended incubation at 10
days. Thus, both extended incubation in seawater and exposure to CM function to
facilitate the phase variation response of V. vulnificus.
Discussion
In the study, the anti-Vibrio activity of CM was evaluated in the context of CPS
expression and phase variation, using V. vulnificus as a model organism. CPS is a
primary virulence factor of V. vulnificus, and protects the bacterium against
phagocytosis. Encapsulated cells are generally more virulent and have an opaque
colony type that can be easily distinguished from the more translucent morphotypes,
which are avirulent due to the lack CPS expression (Wright et al., 1990; Yoshida et al.,
1985). These results demonstrated strain differences in the response to CM, as CMCP6
was significantly more sensitive than MO6-24/O. These strains differ in their CPS
composition, which may contribute to these differences (Bush et al., 1997; Neiman et
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al., 2011). Furthermore, the expression of CPS on the surface appeared to increase
sensitivity to CM, as the unencapsulated mutant strain had increased capacity to
survive after CM treatment in seawater. These results were confirmed by competitive
studies that also showed increased survival in CM treatment of translucent compared to
opaque strains and by the observation that phase variation to translucent phenotype
also increased in response to CM.
It is plausible that CM may kill V. vulnificus by disrupting cell membranes. Using
BacLight staining for viability, it was observed that all surviving cells emitted green
fluorescence (stained with SYTO 9), indicating that sensitive cells were lysed
presumably due to massive loss of membrane integrity (data not shown). Additional
studies are needed to elucidate details of this mechanism, but apparently, CPS
functions to increase CM activity. Bacterial surface charge and polarity are dependent
upon the compositions of outer membrane structures, including proteins, LPS, and
CPS, as well as various phosphate and pyrophosphate groups, that might influence a
net negative charge that could affect the binding of positively charged CM. Significant
strain differences in response to CM treatment were observed between the two strains
examined, suggesting that CM may exert different anti-Vibrio effect as a consequence
of the different cell surface characteristics of individual strains. V. vulnificus have been
shown to exhibit very diverse CPS composition and structure (Hayat et al., 1993).
Perhaps, differences in CPS structure for CMCP6 strain compared to MO6-24/O strain
may account for the observed strain differences in CM sensitivity.
Phenotypic phase variation of opaque to translucent morphotype was observed
at a relatively high frequency in response to sub-lethal CM concentrations. Chitin in an
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intracellular signal for biofilm formation and chitinase production. Chitosan is a
degradation product of chitosan and may perhaps signal induction of translucent phase
variation as a survival response. CPS has been shown to inhibit biofilm in V. vulnificus,
and translucent strains exhibit greater biofilm formation compared to translucent
(Joseph and Wright, 2004). Another possible advantage of reducing CPS expression
would be that cells require less energy to produce the capsule and perhaps phase
variation also redirects that energy to enhance survival. Another report showed that
chitosan treatment affected gene expression, including some essential genes involved
in RNA and protein synthesis and membrane bioenergetics (Raafat et al., 2008).
Taken together, these findings illustrated the contribution of CPS to anti-
V. vulnificus activity of chitosan. However, it is likely that CPS is not the primary target
for CM antibacterial activity, as the acapsular strain was still sensitive to CM, and a
subpopulation of encapsulated strains survived treatment at low concentrations. Further
work should be aimed at clarifying the molecular details of the complex mechanisms
involved in these interactions.
109
Table 5-1. Summary of V. vulnificus strains used in this study.
Strain Description
CMCP6 Encapsulated, virulent, clinical isolate (Kim et al., 2003)
MO6-24/O Encapsulated, virulent, clinical isolate (Wright et al., 2001a)
MO6-24/Δwzb Deletion of wzb in CPS operon, no CPS surface expression (Chatzidaki-Livanis et al., 2006)
110
Figure 5-1. Effects of CMs on survival of individual V. vulnificus strains in ASW with CM. V. vulnificus strains CMCP6 (A), MO6-24/O (B), and MO6-24/Δwzb (C) were incubated in ASW (20 ppt, pH 7.4) with different concentrations of CM (0,
0.05, 0.1, and 0.3% wt:vol) at 37C. Bacterial survival (mean log CFU/ml) was calculated from three independent experiments; standard deviations are indicated by error bar.
111
Figure 5-2. Competitive survival of V. vulnificus MO6-24/O wild-type strain vs. CPS mutant. Mean log ratios ± standard deviation (SD) of CPS deletion mutant (MO6-24/Δwzb) versus wild-type V. vulnificus (MO6-24/O) in ASW are shown. Ratios were based on colony morphology (opaque vs. translucent) of plate counts, as described in text after CM treatment at 0 and 24 h on LA, and reflect the mean of triplicate experiments.
112
Figure 5-3. Comparison of phase variation in colony morphology of V. vulnificus strains. Log ratio of opaque vs. translucent colonies were determined for different concentrations of CM (0, 0.05, 0.1, and 0.3% wt:vol) after A) 1 day and B) 10 days post inoculation at 37°C by plate counts. Bacterial survival (number of mean log CFU/ml) was calculated from three independent experiments; standard deviations are indicated by error bars. ND = No bacteria were recovered on LA plates.
113
CHAPTER 6 SUMMARY AND CONCLUSIONS
One of the primary goals of this research was to provide an analysis of risks
associated with Vibrio species in Apalachicola Bay, as oyster beds in this bay are the
primary source of harvest for the state of Florida. Although several studies have
investigated the distribution of V. vulnificus in this region (Johnson et al., 2010; Tamplin
and Capers, 1992), the prevalence and abundance of other pathogenic species, namely
V. parahaemolyticus and V. cholerae, are essentially unknown, while disease from
contaminated oysters harvested from this bay has been attributed to all three species in
recent surveys (Turner et al., 2014). This research does not provide comprehensive
analysis on the distribution of Vibrios in Apalachicola Bay, but seasonal sampling over
three years for 17 sites yielded positive results for all three species for every time point
and every site examined; however, the three species differed in their association with
different sampling sites, as well as the type of sample examined. Overall, V. vulnificus
and V. parahaemolyticus were prevalent in 100% of oysters throughout the bay in 93
and 76% of water samples, respectively, and also were seen in approximately 70% of
fish samples. In contrast, V. cholerae showed significantly lower levels in oysters and
water and was never detected in fish samples. Furthermore, it was not as widely
distributed throughout Apalachicola Bay as the other species, instead it was mostly
associated with near shore sites with lower salinity and conductivity in both seawater
and oyster samples. Conversely, V. vulnificus and V. parahaemolyticus tended to
persist throughout the Bay, and more positive samples were detected in off shore sites
compared to near shore sites.
114
As a consequence of this differential distribution, both occurrence and
abundance of V. cholerae in oysters and seawater showed a strong inverse correlation
with conductivity and salinity, which is in agreement with previous studies where
V. cholerae was more frequently detected at low salinity (Jiang, 2001; Louis et al.,
2003). In contrast, no significant relationship was found between the presence of V.
vulnificus or V. parahaemolyticus and any environmental factor examined, which was
not consistent with previous larger scale studies in the Gulf of Mexico (Kaspar and
Tamplin, 1993; Kelly, 1982; Randa et al., 2004; CDC, 2009; Daniels et al., 2000;
DePaola et al., 2003). Dissolved oxygen was also shown to have a significant
relationship with V. cholerae presence and occurrence in the oysters but not the water
column. This may reflect the patchiness of the data or potentially, a relationship of V.
cholerae with zooplankton environments, as dissolved oxygen content is vital for algae
and aquatic animal growth. This study offered insight into conductivity and agreed with a
previous study on environmental variables influencing a cholera outbreak in
Bangladesh, which showed that not only water temperature and salinity, but also
rainfall, conductivity, and copepod counts correlated with prevalence of this bacterium
(Huq et al., 1990). Finally, it should be noted that the differential distribution V. cholerae
to near shore sites could also be an indication of human impact from non-point source
fecal contamination, as a consequence of persons with carriage of V. cholerae at sub-
clinical levels. Future studies are needed in order to generate the necessary number of
samples from each sample site and time point for valid statistical comparisons of
species distribution and source-tracking.
115
Investigations into virulence potential of V. cholerae isolates collected from
Apalachicola Bay were conducted in response to multiple reports of cases and one
recent outbreak of disease associated with oysters harvested from the region. Results
revealed a diverse population of strains with some showing close relationship to O75
outbreak strain, while others were closer to O1/O139 pandemic strains. Oyster strains
from this bay were more clonal than either seawater strains or the two isolates from
Tampa Bay, and all oyster strains were closer to O75 than to O1/O139 strains. The
absence of ctxA/B and tcpA genes associated with pandemic strains suggests limited
virulence potential and human health risks associated with these strains; however, the
ubiquitous dissemination of other virulence-associated genes (rtx, ompU, hlyA, toxR) in
Florida isolates implies that these factors play an important role in both environmental
and human host-related survival. The presence of multiple antibiotic resistance genes
provided evidence of the anthropogenic impact to environmental bacteria, as diverse
resistance to different classes of antibiotics was observed with five strains. These data
provide a better understanding of the complex and dynamic factors affecting the
distribution of toxigenic V. cholerae, but further study is warranted to monitor and
characterize genetic structure and virulence potential of these strains in the
environment.
This environmental survey of Vibrios in Apalachicola Bay indicated that efforts to
control vibriosis are essential to sustaining the shellfish industry in Gulf Coast states. To
facilitate public health actions to prevent and control vibriosis from raw shellfish
consumption, the feasibility of chitosan as a PHP treatment for seafood industry was
investigated. Chitosan in the form of microparticles exerted strong antimicrobial activity
116
against Vibrio spp. in both seawater and oysters and offers promising potential for the
application of an environmentally friendly process for intact live oysters. Commercial
usage of CM in the seafood industry will require optimization and greater knowledge of
the mechanisms for anti-Vibrio activity. This study also demonstrated that both capsular
polysaccharide (CPS) expression and phase variation influenced vibriocidal activity,
using V. vulnificus as a model organism. The expression of CPS on the surface
increased sensitivity to CM, and it is plausible that variations in CPS composition may
also contribute to CM sensitivity, as strain differences were observed. Furthermore,
V. vulnificus CPS is commonly composed of uronic acid sugars, which would increase
the negative charge and hydrophilicity (Hayat et al., 1993), while V. cholerae EPS is
primarily composed of neutral sugars glucose and galactose (Yildiz and Schoolnik,
1999). Thus, the more acidic CPS composition of V. vulnificus relative to V. cholerae
CPS may also account for the observed differences in sensitivity between these two
species. A previous study elucidated that CM has greater positive charge at acidic pH;
perhaps charge differential on the surface of these bacteria influences binding and/or
activity of CM (Jeon et al., 2014). This speculation may also explain why V. vulnificus
was more sensitive to CM treatment LB, as compared to seawater. However, it is likely
that CPS is not the primary target for CM antibacterial activity, as the acapsular strain
was still sensitive to CM at higher concentrations, and a subpopulation of encapsulated
strains survived treatment at low concentrations. Taken together, these findings
illustrated the contribution of CPS to anti-V. vulnificus activity of chitosan. Further work
should be aimed at clarifying the molecular details of the complex mechanisms involved
in these interactions.
117
Overall, these studies provide much needed information on the potential risk of
Vibrios in Florida and lay the foundation for development of a novel solution to a
problem that continues to threaten to seafood industry. Recent reports of disease
associated with V. vulnificus in Florida are also a threat to tourism, as cases involving
wound infections are generally not related to oyster consumption and result in warnings
about the safety of recreational waters. There is also growing evidence that climate
change is rapidly increasing the risk of emerging diseases with Vibrios on the forefront
of this trend. The true risk posed by the presence of these species in the Gulf of Mexico,
as well as the sustained public health safety, will only be elucidated through more
rigorous surveillance and better understanding of the evolution of virulence potential.
118
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BIOGRAPHICAL SKETCH
Lei Fang was born in the beautiful capital city of Hangzhou, China, which is
famous for its rare combination of gorgeous natural scenery and prosperous civilization.
Lei attended Xuejun High school and received her bachelor’s degree in food science
and engineering from Zhejiang Gongshang University. Before completing her
undergraduate study as an outstanding student, she was exchanged to Virginia
Polytechnic Institute and State University in 2010, where she learned general lab
techniques and was exposed to several food microbiological projects. In 2011, Lei was
awarded the Alumni Fellowship, one of the most prestigious scholarships at the
University of Florida, and started her Ph.D. degree program in Food Science and
Human Nutrition Department under the direction of her advisor Dr. Anita Wright. In her
spare time, Lei enjoys baking, fishing, swimming, and spending leisure times with her
friends and family. Upon graduation, Lei is devoting herself to a career of food safety
and quality assurance in food industry or government regulation.