Vibrios associated with marine samples from the Northern Gulf of Mexico: 
implications for human health 
 
by 
 
Zhen Tao 
 
 
 
 
A dissertation submitted to the Graduate Faculty of 
Auburn University 
in partial fulfillment of the 
requirements for the Degree of 
 Doctor of Philosophy 
 
Auburn, Alabama 
August 3, 2013 
 
 
 
 
Keywords: recreational fishing, Vibrio vulnificus, public health 
 
 
Copyright 2013 by Zhen Tao 
 
 
Approved by 
 
Covadonga R. Arias, Chair, Full Professor of Fisheries and Allied Aquacultures 
 Thomas A. McCaskey, Professor of Animal Science  
Calvin M. Johnson, Professor of Pathology 
Stephen A. Bullard, Assistant Professor of Fisheries and Allied Aquacultures  
 
 
 
 
ii 
 
Abstract 
 
 
 In this dissertation, I investigated the distribution and prevalence of two human-
pathogenic Vibrio species (V. vulnificus and V. parahaemolyticus) in non-shellfish 
samples including fish, bait shrimp, water, sand and crude oil material released by the 
Deepwater Horizon oil spill along the Northern Gulf of Mexico (GoM) coast.  
 In my study, the Vibrio counts were enumerated in samples by using the most 
probable number procedure or by direct plate counting. In general, V. vulnificus isolates 
recovered from different samples were genotyped based on the polymorphism present in 
16S rRNA or the vcg (virulence correlated gene) locus. Amplified fragment length 
polymorphism (AFLP) was used to resolve the genetic diversity within V. vulnificus 
population isolated from fish. PCR analysis was used to screen for virulence factor genes 
(trh and tdh) in V. parahaemolyticus isolates yielded from bait shrimp. A series of 
laboratory microcosm experiments and an allele-specific quantitative PCR (ASqPCR) 
technique were designed and utilized to reveal the relationship between two V. vulnificus 
16S rRNA types and environmental factors (temperature and salinity). 
 In summary, research data showed that the human pathogen V. vulnificus is 
commonly found in non-shellfish samples (fish, bait shrimp and tar ball) of the Northern 
GoM coast. Moreover, I discovered a higher percentage of strains of great virulence 
potential in fish and shrimp than those previously reported in oysters. I proved that 16S 
type B strains outcompete type A strains at warmer temperatures explaining why more 
ii 
 
cases of vibriosis due to this pathogen occur at the end of summer. Finally, the effects of 
the Deepwater Horizon oil spill significantly increased the presence of V. vulnificus in 
beach samples. Overall, my research shows that recreational activities conducted in the 
Northern GoM coast have an intrinsic risk of exposure to V. vulnificus. 
 
iii 
 
Acknowledgments 
 
 
 First, I would like to express the deepest appreciation to my committee chair 
Professor Dr. Cova R. Arias. Without her guidance and persistent help, this dissertation 
would not have been possible.  I would also like to thank my committee members, Dr. 
Thomas A. McCaskey, Dr. Calvin M. Johnson, and Dr. Stephen A. Bullard for their time 
and expertise. I thank all my lab mates I worked with throughout my time here: Ryan 
Wood, Matt Lewis, ?scar Olivares-Fuster, Stacey LaFrentz, Andrea Larsen, Wenlong 
Cai, Haitham Hussien Mohammed and Jeff Holder. My research would not have been 
possible without their helps. I would also like thank the administration staff and 
colleagues in the Department of Fisheries and Allied Aquacultures for the help over the 
years. I would like to express my gratitude to the Chinese Scholarship Council for their 
financial support through my four-year PhD study. Finally, I would like thank to my 
parents and elder brother. They were always supporting me and encouraging me with 
their best wishes. 
 
 
 
 
 
 
 
 
 
 
iv 
 
Table of Contents 
 
 
Abstract ............................................................................................................................... ii 
Acknowledgments.............................................................................................................. iv  
List of Tables .................................................................................................................... vii 
List of Figures .................................................................................................................... ix 
List of Abbreviations ......................................................................................................... xi 
Chapter 1. Literature review   ..............................................................................................1 
Chapter 2. Objectives   .......................................................................................................36 
Chapter 3. Prevalence and population structure of Vibrio vulnificus on fishes from 
 the northern Gulf of Mexico .................................................................................38 
Abstract   ................................................................................................................38 
Introduction   ..........................................................................................................39 
Material and methods   ...........................................................................................41 
Results   ..................................................................................................................46 
Discussion   ............................................................................................................50 
References  .............................................................................................................56 
Chapter 4. Occurrence of Vibrio vulnificus and V. parahaemolyticus in bait shrimp  
from Alabama and Mississippi ..............................................................................73 
Abstract   ................................................................................................................73 
Introduction   ..........................................................................................................73 
v 
 
Material and methods   ...........................................................................................74 
Results and discussion  ..........................................................................................77 
References  .............................................................................................................81 
Chapter 5. High numbers of Vibrio vulnificus in tar balls collected from oiled areas  
of the north-central Gulf of Mexico following the 2010 BP Deepwater  
Horizon Oil Spill ....................................................................................................90 
Abstract   ................................................................................................................90 
Introduction   ..........................................................................................................91 
Material and methods   ...........................................................................................92 
Results and discussion  ..........................................................................................94 
References  .............................................................................................................96 
Chapter 6. Effect of temperature and salinity on the dynamics of Vibrio vulnificus  
16S rRNA genotypes revealed by allele-specific quantitative PCR ....................101 
Abstract   ..............................................................................................................101 
Introduction   ........................................................................................................102 
Material and methods   .........................................................................................104 
Results ..................................................................................................................112 
Discussion  ...........................................................................................................114 
References  ...........................................................................................................119 
Chapter 7. Summaries and conclusion .............................................................................136 
Appendix 1. Diversity of culturable bacteria from blood vascular system of lesser  
electric rays (Narcine bancroftii) captured in the northern Gulf of Mexico  
off Alabama   .......................................................................................................139 
vi 
 
List of Tables 
 
 
Chapter 3  
Table 3-1 ................................................................................................................61 
Table 3-2  ...............................................................................................................62 
Table 3-3  ...............................................................................................................63 
Table 3-4  ...............................................................................................................65 
Chapter 4 
Table 4-1  ...............................................................................................................85 
Table 4-2 ................................................................................................................87 
Table 4-3 ................................................................................................................88 
Table 4-4 ................................................................................................................89 
Chapter 5 
Table 5-1  ...............................................................................................................98 
Table 5-2 ................................................................................................................99 
Chapter 6 
Table 6-1  .............................................................................................................123 
Table 6-2 ..............................................................................................................124 
Table 6-3  .............................................................................................................125 
Table 6-4  .............................................................................................................126 
Chapter 6 
vii 
 
Table AP-1  ..........................................................................................................158 
 
  
viii 
 
List of Figures 
 
 
Chapter 3 
Figure 3-1  ..............................................................................................................66 
Figure 3-2  ..............................................................................................................67 
Figure 3-3 ...............................................................................................................68 
Figure 3-4 ...............................................................................................................69 
Figure 3-5 ...............................................................................................................70 
Figure 3-6 ...............................................................................................................71 
Chapter 5 
Figure 5-1  ............................................................................................................100 
Chapter 6 
Figure 6-1  ............................................................................................................127 
Figure 6-2  ............................................................................................................129 
Figure 6-3 .............................................................................................................130 
Figure 6-4 .............................................................................................................131 
Figure 6-5 .............................................................................................................132 
Figure 6-6 .............................................................................................................133 
Figure 6-7 .............................................................................................................134 
Figure 6-8 .............................................................................................................135 
Appendix 1 
ix 
 
Figure AP-1  .........................................................................................................161 
Figure AP-2  .........................................................................................................162 
Figure AP-3  .........................................................................................................163 
Figure AP-4 ..........................................................................................................165 
Figure AP-5 ..........................................................................................................167 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
x 
 
List of Abbreviations 
 
 
AFLP Amplified fragment length polymorphism 
ASqPCR Allele specific quantitative PCR 
GoM Gulf of Mexico 
mCPC modified Colistin Polymixin-B Cellobiose agar 
MDS Multidimensional scaling 
RFLP Restriction fragment length polymorphism 
rrn  16S rRNA operon 
TCBS thiosulfate-citrate-bile salts-sucrose agar  
TDH thermostable direct hemolysin 
TRH TDH-related hemolysin 
1 
 
 
 
 
 
 
CHAPTER 1. LITERATURE REVIEW 
 
 
Gulf of Mexico Ecosystem 
 The GoM is a semi-enclosed sea and is the ninth largest body of water in the 
world. It is bordered by the United States to the north (Florida, Alabama, Mississippi, 
Louisiana, Texas), five Mexican states to the west (Tamaulipas, Veracruz, Tabasco, 
Campeche, Yucatan), and the island of Cuba to the southeast. The GoM region covers 
more than 1,942,500 km2 including open water areas and coastal wetlands with input 
from 33 major river systems (1). In total, there are 207 significant estuarine systems, and 
extensive barrier-islands with coastal lagoons, both in the United States and Mexico. 
Every year ca. 60% rainfall in continental American and about 40% in Mexico discharge 
into the Gulf (1). GoM habitats include coastal wetlands, submerged vegetation, upland 
areas, and marine and offshore areas. Due to its location, the GoM is transitional between 
temperate and tropical climate and comprises both elements. The diverse marine habitats 
and climate conditions teems a high biodiversity in the GoM.  A recent inventory 
compiled by Felder and Camp (2) describes about 15,419 eukaryotic species which are 
distributed among over 40 phyla. This number is estimated to cover roughly 80% to 85% 
of the GoM eukaryotic taxa.  However, the number of microbial species is not determined 
in the inventory.  Whitman et al. (3) estimated that the number of prokaryotic cells in 
ocean and oceanic subsurface are 1.2 ?1029 and 3.5 ? 1030, respectively. These 
2 
 
microorganisms play a wide range of ecological and biogeochemical roles (i.e. carbon 
recycling ) in the GoM ecosystems (4). In addition to these roles, several groups of 
microbes can be pathogenic to marine animals and humans. Among those, ,members of 
the family of Vibrionaceae have a long history of being associated with human diseases 
(2). However, the distribution of pathogenic species of vibrios in the GoM environment is 
poorly study, in particular, their associations with fish is largely unknown.   
The genus Vibrio 
The genus Vibrio is a major taxa of culturable heterotrophic bacteria commonly 
found in marine and estuarine environments (5, 6). This genus was first described by the 
Italian physician Filippo Pacini in 1854 while he was studying cholera as a medical 
student. This genus is classified into family Vibrionaceae, which, as of 2012, is the only 
family in the order Vibrionales. The family includes the genus Vibrio (111 species) and 
other 10 genera: Aliivibrio (6 species), Allomonas (1 species), Beneckea (11 species), 
Catenococcus (1species), Enterovibrio (4 species), Grimontia (1species ) Listonella (3 
species ), Lucibacterium ( 1species),  Photobacterium ( 24 species)  and  Salinivibrio (4 
species) (7). According to the Bergey?s Manual of Systematic Bacteriology (8), Vibrio 
spp. are characterized by being small (0.5-0.8 ? 1.4-2.6 ?m), comma-shaped rods with 
polar flagella enclosed in a sheath. They are motile, facultative anaerobes, and oxidase 
positive (except for 2 species). All Vibrio species can ferment D-glucose without 
producing gas and are able to reduce nitrate to nitrite.  
Pathogenic Vibrio spp. Within the genus Vibrio, there are several pathogenic 
species that can cause disease in humans and marine animals (8, 9). Historically, Vibrio 
3 
 
cholera, V. parahaemolyticus and V. vulnificus has been recognized as three most serious 
human pathogens in the genus (10-12) but other species including V. alginolyticus, V. 
fluvialis, V. furnissii, V. mimicus, V. metschnikovii, and V. cincinnatiensis have also been 
reported as the cause of illness in humans (13-19) . Regarding aquatic animals, vibriosis 
is one of the most serious disease affecting marine invertebrate and vertebrates. Some of 
the Vibrio species that cause significant economic losses in finfish, shellfish, and shrimp 
marine culture are Listonella (formerly Vibrio) anguillarum, V. ordalii, V. salmonicida, 
V. tapetis, V. harveyi, and V. vulnificus biotype 2 (20-23).  Other less common Vibrio 
species that cause disease in marine animals are V. splendidus, V. neptunius, V. 
alginolyticus, V. parahaemolyticus, and V. penaeicida. In addition, V. coralliilyticus has 
been associated with coral (Pocillopora damicornis) bleaching, (24-29). As more studies 
on aquatic animal health are published, the list of Vibrio species recognized as putative or 
opportunistic pathogens continues to increase.  
Host-Vibrio interactions. Many species of Vibrio form symbiotic and commensal 
relationships with aquatic animals.  As a part of the natural microbiota of estuarine and 
marine ecosystems, Vibrio species frequently encounter a wide-range of hosts such as 
zooplankton, shellfish, crustacean and fishes. These interactions can range from 
mutualistic symbiosis to pathogenic relationships. 
Vibrio cholerae, the bacterium that causes cholera, can attach to chitinaceous 
zooplankton, particularly copepods (30).  This kind of association with zooplankton is 
proven to be a key factor in deciphering the global nature of cholera epidemics, because 
V. cholerae can use zooplankton as a vector and colonize to new environments (31, 32). 
4 
 
In addition, zooplankton (live or dead) can support the growth of some Vibrio species 
such as V. parahaemolyticus, suggesting that zooplankton can act as possible reservoir 
for this pathogen in the environment (33). Furthermore, a more recent study conducted by 
Turner et al. (34) demonstrated that plankton composition modulates the presence of 
culturable Vibrio spp. thus, indicating the potential complex interactions between 
zooplankton and Vibrio species.  Finally, Senderovich et al. (35) reported that V. cholerae 
can be found in the gastrointestinal tract of fish to where arrives through the food chain 
thus, suggesting that fish can serve as a reservoir for the bacterium.  
Vibrio species are frequently found in shellfish as well. Marine bivalves such as 
oysters and mussels that are filter feeders accumulate microbiota including vibrios from 
surrounding waters (36). For example, high concentrations (105 CFU?g-1) of V. vulnificus 
are commonly found in GoM oysters (37). Vibrio vulnificus is recognized as the most 
invasive and rapidly lethal Vibrio species for humans (38) but it is a mere commensal to 
the oysters. Oysters harvested from waters in where the bacterium is present, become 
passive carriers for this opportunistic human pathogen. Interestingly, an earlier study 
showed V. vulnificus is more persistent in oyster tissues than fecal coliforms (e.g. 
Escherichia coli) and therefore more difficult to remove during post- harvest processes of 
shellfish, including depuration (39).  A study conducted by Paranjpye et al. showed that 
V. vulnificus mutants that lack pili had a significantly reduced ability to persist in oyster 
(Crassostrea virginica) tissues (40).  This finding suggested that type IV pili expressed 
by V. vulnificus are involved in attachment and colonization of oysters.  
5 
 
In crustaceans,  Vibrio spp. accumulate in the digestive tract and the 
hepatopancreas where they can reach high numbers (5). Oxley et al. analyzed the gut 
microbiota of banana prawn (Penaeus merguiensis), showing that Vibrio species 
(including V. parahaemolyticus) were the dominant group in the gut of healthy cultured 
prawns (41).  Another study also found high concentration (from 104  to 105 CFU?g-1 
tissue) of  Vibrio spp. in the hepatopancreas of healthy shrimp (Penaeus vannamei) 
juveniles (42). 
Vibrio communities are often part of the normal microbiota associated with fishes. 
In an early survey, Grimes et al.  recovered Vibrio spp. from blood, liver and other 
internal organs of healthy sharks (43, 44). This finding was further confirmed by routine 
surveys of captive elasmobranches in which bacteria were isolated from the blood of 
healthy animals (45). It has been hypothesized that the bacteria found in the blood of 
elasmobranches could play a role in osmoregulation as most of the recovered isolates can 
hydrolyze urea (46, 47).  In addition to sharks, Vibrio spp. have been reported from 
internal organs of healthy farmed turbot (Scophthalmus maximus ) (48). However, the 
role that these bacteria play in host health and homeostasis is unclear and needs further 
investigation.  
Vibrio vulnificus 
Vibrio vulnificus was first isolated by the US Centers for Disease Control (CDC) 
in 1964 but misidentified as a virulent strain of V. parahaemolyticus. In 1976, it was 
referred to as ?lactose fermenting vibrio?, which was one of main characteristics that 
distinguished this organism from two other closely related species: V. parahaemolyticus 
6 
 
and V. alginolyticus (49).  The bacterium was formally described as Vibrio vulnificus by 
Farmer et al. in 1979 (50). The epithet ?vulnificus? derives from the Latin ?wound?, as the 
first cases on infections caused by V. vulnificus involved wound infections. 
Ecology. Vibrio vulnificus is an autochthonous estuarine and marine bacterium 
that can be found in temperate and tropical climates worldwide (51).  In the United 
States, it has been isolated from the Atlantic, Pacific, and GoM coasts including estuaries 
in those areas (52-56).  In the natural environment, V. vulnificus is generally a free-living 
bacterium that is commonly found in water or sediment (57). In addition, it has been 
associated with shellfish (ie. oyster and clams) (52, 58), intestine of fishes (59), plankton 
(54), and seaweeds (60). Notably, the prevalence and abundance of V. vulnificus is 
governed by environmental factors, particularly by temperature and salinity (55, 61-63).  
Several studies have showed that the occurrence of V. vulnificus usually undergoes a 
seasonal pattern that is dependent on temperature.  When water temperature is above 
20?C, V. vulnificus can be easily recovered from seawater and shellfish as long as the 
salinity remains within moderate values (15 ppt ~ 25 ppt). Consequently, in our latitude, 
V. vulnificus is abundant during the summer months while remains in low to non-
detectable numbers in winter. During the colder months, the low concentration of V. 
vulnificus numbers in the water column led to hypothesize that the bacterium entered the 
?viable but not culturable? state in response to cold conditions (64, 65).  The term ?viable 
but not culturable? (VBNC) for bacterial cells was first coined by Roszak and Colwell 
(66) which refers to a life stage characterized by cells that still retain metabolic function 
but are unable to grow on culturable media under laboratory conditions. Although unable 
7 
 
to grow on media, V. vulnificus cells in this state retain their viability and, most 
important, their virulence (67).  Under in vitro conditions, it was possible to resuscitated 
V. vulnificus cells that had entered the VBNC state by temperature upshift (68).  
However, the ?resuscitation? of the VBNC cells in the environment has not been 
demonstrated and the true relevance of VBNC forms remains a subject of controversy 
(69).   
An alternative explanation for the decrease in V. vulnificus numbers in the water 
column during the winter months is that the vectors/hosts carrying this bacterium also 
decrease in winter. A recent study using culture independent methods provided evidence 
that supports the second hypothesis. The authors showed that the significant reduction of 
population size in winter is the primary factor for the observed decline in V. vulnificus 
colony counts and not the bacteria entering the VBNC state (53). According to this 
theory, the sediment will act as main reservoir for V. vulnificus during the winter months.  
In addition to temperature, the dynamics of V. vulnificus is strongly correlated to 
the salinity of the marine environment.  Although this bacterium has been isolated from 
waters with salinities ranging from 1 ppt to 34 ppt, most isolates are recovered from 
salinities between 15 ppt to 25 ppt (63, 70). The growth of V. vulnificus is negatively 
influenced by salinities greater than 25 ppt (63, 71). For instance, the low incidence of V. 
vulnificus among Vibrio isolates from seawater of the Mediterranean coast has been 
correlated to the constant high salinity (above 30.7 ppt ) typical from the area (72). The 
northern GoM coast, where low to medium salinities (5 ppt ~ 25 ppt facilitates the 
abundance of V. vulnificus, whereas estuaries in similar latitudes on the Atlantic coast are 
8 
 
generally too saline (>28 ppt) to accommodate for this organism (63).  It was found that 
the concentration of V. vulnificus was influenced primarily by water temperature in 
coastal waters of the northern GoM, although V. vulnificus numbers are significantly 
reduced in salinities higher than 26 ppt.  Reversely, salinity is the main factor when 
moderate temperatures occur year round (55). In general, the interrelationship of 
temperature and salinity plays a complex role that controls the distribution and 
fluctuation of this organism in the aquatic environments. Other factors such as dissolved 
oxygen (73), turbidity (74), chlorophyll (75), plankton (34)  and phages (76) have also 
been found related with the ecology of  V. vulnificus. 
Biotypes. The species V. vulnificus is divided into three biotypes which are 
characterized by specific biochemical properties (19, 77).  Table 1 summarizes the 
discriminative biochemical tests between all three biotypes. Biotype 1 strains are positive 
for indole production and ornithine decarboxylation.  Biotype 1 the predominant  biotype 
in the USA and responsible for all human infections reported in this country (10, 78).  
Biotype 2 strains are negative for indole and ornithine decarboxylation, and have been 
associated with severe outbreaks of vibriosis in eels (77, 79). However, at least one 
biotype 2 strain has been isolated from a wound infection in the USA (80). Biotype 2 
strains are immunologically identical, sharing a common lipopolysaccharide profile that 
differs from that of biotype 1 strains (81). In some studies, biotype 2 is referred to as 
serovar E (from eel) although a few non-serovar E biotype 2 strains have also been 
reported (82).  Biotype 3 was first reported by Bisharat et al. in 1999 (83).  Isolates in this 
subgroup were associated with wound infection in fishermen that have been in contact 
9 
 
with tilapia infected with V. vulnificus.  Biotype 3 strains differ phenotypically from 
biotype 1 and 2 strains in five phenotypic characteristics: they are negative for citrate, 
and o-nitrophenyl-?-D-galactopyranoside tests, and also negative for salicin, cellobiose 
and lactose fermentation. A genetic study using multilocus sequence typing in where 10 
housekeeping genes were analyzed, indicated that biotype 3 is a hybrid of biotypes 1 and 
2 (84). 
Genotypes. V. vulnificus is a heterogenetically diverse species. Environmental 
isolates of V. vulnificus, even those isolated from a single oyster, display a remarkable 
degree of genetic variation (85). Conversely, clinical isolates are more similar to each 
other. Early studies indicated that only a small fraction of the strains isolated from 
shellfish caused infections in humans but the search for a specific virulence-marker 
continues these days (86).  The main difficulty for finding a virulence-associated marker 
relies on the fact that the both clinical and environmental isolates presented all the known 
putative virulence factors described for biotype 1 (70). For the past 15 years, several 
research groups have been trying to find molecular markers that could separate the 
putative more virulent isolates from the rest. The two most common single locus 
genotyping methods used for the genotyping of V. vulnificus are herein discussed.  
In one genetic analysis on V. vulnificus, Warner et al. employed randomly 
amplified polymorphic DNA (RAPD) PCR to characterize the strains at the subspecies 
level (87).  This study showed that all of the clinical strains produced an additional band 
that was rarely amplified in environmental strains.  In a follow-up study, this fragment of 
genomic DNA (named as virulence correlated gene or vcg) was sequenced and analyzed, 
10 
 
and the consistent sequence variations were highly correlated with the isolation source, 
where clinical and environmental isolates were significantly associated with vcgC and 
vcgE type, respectively  (88).  The other widely-used subtyping scheme also separated 
the V. vulnificus strains into two subspecies groups, designated as type A and type B 
according to the 16S rRNA sequence polymorphisms (89). In total, there are 17 different 
nucleotides in the 16S rRNA gene that can differentiate type A and B. Originally, this 
polymorphism was identified in the type strain of the species V. vulnificus strain 
ATCC27562 and in a clinical isolate C7184 (89) (GenBank accession number X76333 
and X 76334, respectively).  In a later study by Nilsson et al. (90), a good correlation was 
found between 16S genotypes and source of isolation in a large collection of V. vulnificus 
isolates. In that study, 94% of nonclinical isolates were type A (31 of 33), while type B 
comprised the majority (76%) of human clinical strains (26 of 34).  Using real-time PCR 
for the rapid determination of 16S rRNA types, Vickery et al. also proved the existence 
of a hybrid type AB that contains both A-type and B-type 16S rRNA gene alleles (91). 
The AB-type strains were associated with nonclinical sources, and shared higher genetic 
similarity with A-type strains than with B-type (92, 93).  Both typing methods concur in 
defining two major genotypes within V. vulnificus that exhibit a strong correlation with 
source of isolation. However, a recent study demonstrated that strains of 16S rRNA type 
A, AB and vcgE could be highly virulent in a mouse model, indicating that genotypes are 
correlated with but do not equate to virulence potential in the species V. vulnificus (94). 
Hence, a specific virulence marker is still lacking for this species. 
11 
 
Virulence and Pathogenesis. The virulence of V. vulnificus has been inferred to 
be multifactorial (95-97). An arsenal of virulence factors have been identified in V. 
vulnificus including capsular polysaccharide, the ability to acquire iron, 
lipopolysaccharides, pili, and the RtxA1 toxin (encoded by the rtxA1 gene). In addition, 
other putative virulence factors like extracellular enzymes metalloprotease (encoded by 
vvpE) and hemolysin-cytosin (encoded by vvhA gene) have been proposed (98). 
Nonetheless, these enzymes showed no definitive role in the pathogenesis of V. vulnificus 
in mammalian models (99). To date, all the identified virulence factors can only partially 
explained the extreme tissue damage and high mortality observed in both primary 
septicemia and wound infections caused by V. vulnificus. The full pathogenesis of this 
bacterium is still not fully understood.  
One of the best studied virulence factors in V. vulnificus is the capsular 
polysaccharide (CPS) (100, 101). Early morphological observations identified 
encapsulated strains that formed opaque colonies and translucent colonies that had little 
or no capsule (100).  A positive correlation has been found between the presence and 
amount of CPS in V. vulnificus isolates with virulence in mice (101).  The expression of 
capsular polysaccharides contributes to the virulence in V. vulnificus with two roles.  
First, the presence of CPS on the cell surface helps the bacterium eluding the host 
defenses since capsule products confer V. vulnificus  resistance against the bactericidal 
effects of serum (102). The presence of capsule also provides resistance against 
opsoization by complement and thus avoidance of phagocytosis by macrophages (103). 
12 
 
Second, CPS partially contributes to septic shock formation through the production of 
inflammation-associated cytokines (108).  
The ability to acquire iron from the host?s transferrin is another important 
virulence factor present in V. vulnificus. In human serum, most of the iron is bound to 
transferring making it unavailable to infectious organisms.  In order to establish a 
successful infection, V. vulnificus developed systems to overcome this iron limitation. 
Indeed, multiple systems for iron acquisition (or siderophores) have been observed in V. 
vulnificus (104). The bacteria produce two types of siderophores: a catechol and 
hydroxymate siderophores, which are primarily involved in iron acquisition (105).  
Lipopolysaccharide (LPS) is the putative virulence factor that likely causes the 
septic shock observed in V. vulnificus septicemia cases through the induction of host 
pyrogenic responses (106).  Others also suggested that the some symptoms such as fever, 
tissue edema, and hypotension in response to LPS (107).  
In bacteria, the adherence to hosts is often mediated by pili. Gander et al. reported 
that clinical V. vulnificus isolates from blood or wounds of infected individuals averaged 
higher numbers of individual pilus fibers per cell than environmental isolates (108). 
While another study indicates that type IV pili are conserved in both clinical and 
environmental strains, and has a role in adherence to human epithelial cells as well in 
biofilm forming (109). It was demonstrated that the decreased virulence is also detected 
in a mutant strain that is lacking of a type IV prepilin peptidase (109). 
The ability of V. vulni?cus to cause disease has also been associated with a 
cytotoxin called RtxA1 toxin (110-112).  RtxA1 toxin causes aggregation of actin fibers 
13 
 
and plasma membrane blebs which culminated in a necrotic cell death in host epithelial 
cells.  As a result, it allows V. vulnificus cells to invade the bloodstream by crossing the 
intestine epithelium cells. Furthermore, RtxA1-mediated cytotoxicity would be contact-
dependent and toxin production was introduced by host cell contact (113). Overall, 
RtxA1 toxin is an important virulence factor by the intragastric route of infection in 
animal models, and likely plays a significant role in the food-borne infection related to V. 
vulnificus.  
In addition to virulence factors described above, V. vulnificus cells produce 
multiple enzymatic factors like such as metalloprotease, hemolycin-cytolysin, chitinase, 
chondroitinase, collagenase, deoxyribonucleic, elastase, gelatinase, lecithinase, mucinase 
and phospholipase. These extracellular proteins may play an important role in taking 
advantage of the host throughout the course of infection.  
Vibrio vulnificus infections and Public Health  
In the United States, surveillance for Vibriosis has been conducted for more than 
two decades. There are two main surveillance data bases maintained by Center for 
Disease Control and Prevention (CDC) that record V. vulnificus infections. COVIS was 
initiated by CDC, FDA, and the GoM Coast states (Alabama, Florida, Louisiana, 
Mississippi, and Texas) in 1988.  By 1997, nearly all states were voluntarily reporting 
into COVIS. In 2007, vibriosis became nationally notifiable and all 50 states are 
mandated to report any vibriosis case to the CDC. The second data base is the 10-state 
Foodborne Diseases Active Surveillance Network (FoodNet).  COVIS is a passive 
surveillance, while FoodNet is an active surveillance in sentinel populations for 
14 
 
laboratory confirmed Vibrio infections.  Through COVIS and FoodNet, Newton et al. 
examined all cases of vibriosis reported to the CDC from 1996 to 2010 and found that the 
incidence of vibriosis increased from 1996 to 2010. In this study they also found that V. 
vulnificus was the most commonly reported species (114). Unfortunately, this finding 
indicates that the current prevention efforts have not been effective to decrease vibriosis.  
Epidemiology. Geographically, cases of human infections caused by V. vulnificus 
have been reported from most locations where the bacterium was found in the 
environment (51, 115-118).  In the United States, vibriosis associated with V. vulnificus 
occur most commonly in the GoM Coast region due to the warmer water temperature and 
consumption of raw shellfish (17, 119). The incidence of reported infections according 
community-based data in coastal regions is ca. 0.5 cases/100,000 population/year (120). 
Using data from two CDC surveillance systems, an analysis showed an increased in the 
incidence of infections from 1996 to 2010 (the peaked was observed in 2010) (114).  This 
report reviewed all 1446 cases of V. vulnificus infections reported to the Cholera and 
Other Vibrio Illness and Surveillance (COVIS) and all 193 cases reported to the 
Foodborne Diseases Active Surveillance Network (FoodNet). The hospitalization rate of 
infections with V. vulnificus was high (86%, COVIS) and the overall mortality in the US 
was higher (ca. 30%) than that occurred in vibriosis caused by other Vibrio spp. The 
demographic analysis shows that the illnesses typically occurred in males (68% of total 
cases) of 40-49 years of age (19% COVIS, 21% in FoodNet).  Most infections occurred 
primarily during the warmer months of the year and peaked in July and August. This 
results is not surprising since V. vulnificus proliferates in warmer water with temperature 
15 
 
above 20?C. Vibrio vulnificus infections can be acquired through two routes: i) infection 
following ingestion of raw or undercooked seafood, particularly oysters, containing this 
pathogen (121);  ii) infection of preexisting wounds or those incurred during seawater-
associated activities, for instance, a penetrating injury by fish fin or cut obtained in 
shucking oyster (51).   
The most common clinical manifestations associated with V. vulnificus illness are 
gastroenteritis, primary septicemia and wound infection. Gastroenteritis can be 
characterized by enteric symptoms in which a stool culture yielded V. vulnificus, and 
clinical presentations of wound infection and septicemia are excluded.  Retrospective 
studies showed that symptoms in patients with this type gastroenteritis encompass 
predominantly diarrhea, followed by abdominal cramps, nausea, vomiting, fever, and 
bloody stools (17, 122).  Gastroenteritis induced by V. vulnificus is usually self-limiting 
and not life-threatening and is therefore thought to be largely underreported.   
Primary septicemia is defined as a systemic illness characterized by fever and 
shock and in which V. vulnificus was isolated from blood or other sites typically sterile, 
but no wound infection preceding illness was reported (70). Nearly all the cases of 
primary septicemia were followed by consumption of raw or undercooked oyster or other 
raw seafood (119, 121). Main symptoms include fever, chills, nausea and hypertension 
(systolic blood pressure < 85 mm Hg). Notably, secondary lesions develop on either 
extremities or the trunk as a direct result of sepsis in approximately two thirds of the 
cases (123). The time for developing symptoms after ingesting oyster is typically less 
than 36 hours, ranging from 7 hours to 10 days (78). The reported mortality rate with this 
16 
 
illness is between 40% and 60% (123). However, it can reach up to 90% for those who 
become hypotensive within 12 hours of initial presentation (120).   
Wound infections are characterized by the inflammation at the wound site where 
V. vulnificus is directly isolated.  They differ from primary septicemia only in the 
presence of a cutaneous portal of entry. Symptoms of wound infections include pain, 
erythema and edema at wound site where the pathogen was isolated, which can rapidly 
progress to cellulitis, bullous lesions and necrosis, and it could even develop into 
secondary bacteremia in some patients (78). Symptoms began within 4 h to 4 days in 
general although the onset of the disease typically occurs around 24 hours post-exposure 
(51, 124). The fatality rate for V. vulnificus wound infection is approximately 20% (120, 
125). 
Epidemiological studies showed that patient with infections caused by V. 
vulnificus were probably predisposed by certain underlying medical conditions (13, 17, 
119, 126).  Typical conditions found in patients suffering from V. vulnificus infections 
include liver diseases, immune disorders (i.e. AIDS) and some chronic conditions (i.e. 
diabetes mellitus). Preexisting conditions are more prevalent among primary septicemia 
patients than in those with either gastroenteritis or wound infections. Liver disease was 
the most common preexisting condition in all primary septicemia patients (> 80%). Why 
patients with underlying diseases are more susceptible to V. vulnificus infections? It is 
speculated that the elevated iron level in serum associated with those diseases favors 
pathogen growth while suppressing the host immune system (127, 128). People with 
chronic a liver disease or compromised immune system have been described to be up 80 
17 
 
times more likely to develop primary sepsis than healthy individuals. By contrast, V. 
vulnificus-induced gastroenteritis and wound infections do not seem to be favored by 
preexisting conditions and can occur in healthy individuals (119). However, severe 
outcomes like necrotizing fasciitis or secondary septicemia are complications of wound 
infections that are more common in those who belong to the previously mentioned high-
risk group.   
Interestingly, males markedly outnumbered females in terms of V. vulnificus-
related wound infections and primary septicemia (51, 129, 130), and one single study 
found it was due to the protection that estrogen provides to host against V. vulnificus 
(131).  Other risk factors may also contribute to Vibrio vulnificus infections, i.e. alcohol 
abuse (132) and advance age (129, 133). Given all those potential predictors for host 
susceptibility, a report shows that the susceptible individuals represent between 7% and 
16% of the adult population in the USA (134). However, the V. vulnificus infections are 
rare and it is estimated that less than one case of V. vulnificus-induced illness occurs per 
10,000 meals of raw oysters served (134).  There are some explanations as to why only a 
few cases of primary septicemia are reported annually. One possible explanation is that 
not all V. vulnificus strains are equally pathogenic and only a subset is able to cause 
disease in humans. Another explanation could be that only a subset of the susceptible risk 
population is indeed a true-high risk group (135). Currently, it is necessary to 
characterize all risk factors in susceptible hosts and the virulence factors of V. vulnificus 
strains before we can assess the true infection risks.  
18 
 
Foodborne infections. As an etiologic agent for vibriosis, V. vulnificus is both a 
foodborne and a nonfoodborne pathogen. As foodborne pathogen, it is largely associated 
with the consumption of raw oysters and primary septicemia. Oysters are filter feeders 
that concentrate V. vulnificus from water column into their digestive tracts. Since V. 
vulnificus is a natural component of the GoM coast aquatic inhabits, elimination of this 
bacterium from oyster tissues is difficult unless they are processed. In fact, high 
concentrations of  V. vulnificus (>103 per g) are typically found in GoM oysters during 
the summer months (63). On the other hand, oysters are popular seafood in US and it is 
estimated that about 20 million Americans consume raw oysters annually (136). From a 
public health point of view, raw oysters harvested from the GoM should undergo 
postharvest processing, especially during summer months, to ensure the safety of the 
product. For instance, the state of California has banned the sale of untreated raw oysters 
from the GoM states during summer months starting spring 2003. In 2009, the U.S. Food 
and Drug Administration (FDA) was planning to impose a ban on live oyster sales in the 
Gulf region during the warmer months to eliminate deaths related to the consumption raw 
oyster containing V. vulnificus (137). The proposed regulation would require all oysters 
from the GoM region harvested between May and October to be processed post-harvest.  
However, the plan was not implemented due to unclear impact to the oyster industries in 
the GoM region which supports local community economy. Indeed, the GoM is the major 
harvesting region for eastern oyster (Crassostrea virginica) producing about 90% of all 
the eastern oysters in the US and accounting for $ 64,244,826 annually (138). Eventually, 
it is believed that federal regulatory mandates and market constraints would be set up 
19 
 
according to different states to mandate the post-harvest processing for raw oyster 
production in the GoM.   
Nonfoodborne infections. In addition to foodborne infections, V. vulnificus is 
also an important nonfoodborne pathogen in US. Using data from COVIS, Dechet et al. 
reviewed all the nonfoodborne Vibrio infections (NFVI) from 1997 to 2006 in the US, 
showing that V. vulnificus infections constituted 35% of those cases, and 72% of which 
was reported from residents of GoM Coast states (129).  Herein, the NFVI mostly refer to 
wound infections with documented direct contact with salt water, marine wildlife, raw 
seafood, or seafood drippings (not including consumption of raw seafood). Among those 
with a pre-existing wound or a cut acquired on site, up to 70% of exposures were related 
with marine recreational water activities, including boating, surfing, swimming, and 
shore walking. Reversely, wound infections caused by V. vulnificus were more frequently 
followed by handling or cleaning seafood (58% of all cases) than by recreational 
activities (129). Notably, nonfoodborne V. vulnificus infections has also been linked to 
fishing and handling fishing equipment. Indeed, recreational fishing is a favorite outdoor 
activity in US very popular along the GoM coast. The northern GoM is one of top 
destinations for recreational anglers where an estimated >2.8 million anglers participate 
in more than 7 million fishing trips annually (139). Although many people engage in 
recreational activities in marine and estuarine waters and handle fish or bait shrimp there, 
little information is available regarding the prevalence and distribution of V. vulnificus in 
GoM fishes and bait shrimp. This highlights the need to identify the risk factors 
20 
 
associated with wound infections caused by V. vulnificus, particularly those related to 
fishing. 
Vibrio sp. infections after environmental disasters. The risk for illness caused 
by vibrios after natural disasters along the GoM Coast is also a public health concern. For 
instance, during the aftermath of hurricane Katrina, August and September 2005, 22 
cases of Vibrio-wound infection resulting in 5 deaths were reported. People become in 
contact with warm and low salinity water while suffering injuries caused by debris (140).  
More recently, the man-made disaster -the 2010 BP Deepwater Horizon oil spill (DHOS) 
potentially changed the natural environment of V. vulnificus in some areas along the GoM 
coast. The spill released approximately 4.9 million barrels (779 million L) of Louisiana 
light sweet crude oil (from the MC-252 Macondo well) into the north-central GoM over a 
period from 20 April 2010 through 15 July 2010 (141). It is not clear the exact effect that 
this amount of allochthonous carbon had on vibrios living in the affected areas. 
Specifically, the effect that that carbon had on human pathogens is unknown.  Natural or 
man-made disasters often offer fertile grounds to address basic and applied research 
questions. Understanding the ecosystem resilience during and after catastrophic events 
will help developing better response efforts including preparation for infectious diseases 
and possible outbreaks.   
 In summary, V. vulnificus is an autochthonous inhabitant of the GoM coast and a 
potential threat to public health. Humans can be easily exposed to this bacterium by 
eating raw oysters during the summer months and by performing recreational activities 
in, on, or around seawater.  Thus, a better understanding of the ecology of V. vulnificus is 
21 
 
critical to fully delineate a comprehensive risk assessment plan that can be used by public 
health authorities to educate the public and reduce the number of vibrio cases in the 
population.       
22 
 
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_Full_Hq-Print_111110.Pdf Accessed June 12,  2011. 
 
 
  
36 
 
 
 
 
 
 
CHAPTER 2. OBJECTIVES 
 
 
 The overarching goal of my research was to help public health specialists to 
prevention and control human vibriosis in the United States by studying the zoonotic 
bacterial agents V. vulnificus and V. parahaemolyticus. I focused on understanding the 
distribution and prevalence of these two pathogenic bacteria in non-shellfish samples 
including fish, bait shrimp, water, sand and crude oil material released by the Deepwater 
Horizon oil spill. The null hypothesis (H0) for my dissertation research was that 
recreational activities performed at the GoM coast are not associated with a higher risk of 
contracting vibriosis.  To test the above hypothesis, I proposed the following objectives:  
1. Estimate the prevalence of V. vulnificus on skin and mucus of fishes.  
a. If samples yielded V. vulnificus isolates, those will be typified at the 
subspecies level to determine any correlation between V. vulnificus types 
and environmental or biological factors. 
2. Enumerate V. vulnificus and V. parahaemolyticus in bait shrimp.  
a. Genotype recovered V. vulnificus isolates and screen for virulence-factor 
genes in V. parahaemolyticus isolates. 
3. Enumerate the V. vulnificus in water, sand and tarball collected from GoM 
Coastal region and statistically analyze the difference of concentration between 
different type of samples. 
37 
 
4. Assess the adaptation of V. vulnificus strains of 16S type A and type B to various 
environmental conditions by using laboratory microcosms combined with allele 
specific quantitative PCR technique.  
  
38 
 
 
 
 
 
 
CHAPTER 3. PREVALENCE AND POPULATION STRUCTURE OF VIBRIO 
VULNIFICUS ON FISHES FROM THE NORTHERN GULF OF MEXICO 
 
 
Abstract 
 The prevalence of Vibrio vulnificus on the external surfaces of fish from the 
northern GoM was determined in this study. A collection of 244 fish comprising 20 
species was analyzed during the course of 12 sampling trips over a 16-month period. The 
prevalence of V. vulnificus was 37% but increased up to 69% in summer. A positive 
correlation was found between the percentages of V. vulnificus-positive fish (Vv+) and 
water temperatures, while salinity and Vv+ prevalence were inversely correlated. A 
general lineal model (Vv+% = 0.5930 - 0.02818 ? salinity+ 0.01406 ? water temperature) 
was applied to best fit the data. Analysis of population structure was carried out using 
244 isolates recovered from fish. Ascription to 16S rRNA gene types indicated that 157 
isolates were type A (62%), 72 (29%) were type B and 22 (9%) were type AB. The 
percentage of type B isolates, considered to have greater virulence potential, was higher 
than previously reported in oyster samples from the northern GoM. Amplified fragment 
length polymorphism (AFLP) was used to resolve the genetic diversity within the 
species. One hundred and twenty one unique AFLP profiles were found among all 
analyzed isolates resulting in a calculated Simpson?s index of diversity of 0.991. AFLP 
profiles were not grouped based on collection date, fish species, temperature or salinity, 
but isolates were clustered into 2 main groups that correlated precisely with 16S type. 
39 
 
The population of V. vulnificus associated with fishes from the northern GoM is 
heterogeneous and includes strains of great virulence potential. 
Introduction 
Vibrio vulnificus is a Gram-negative bacterium commonly found in estuarine and 
coastal habitats throughout the northern GoM (1, 2). This species is an opportunistic 
human pathogen that can cause primary septicemia, wound infection, and gastroenteritis 
in susceptible individuals (3).  Gastroenteritis is the more benign but less common 
clinical syndrome associated with V. vulnificus infections that typically courses as a self-
limited illness. Conversely, primary septicemia is the most common and severe 
manifestation of V. vulnificus-associated illnesses, having a mortality rate of more than 
50% (3-5). Both gastroenteritis and primary septicemia are associated with the 
consumption of raw shellfish harboring the pathogen, particularly the Eastern oyster 
(Crassostrea virginica) (3). In addition, V. vulnificus can produce severe skin and soft-
tissue infections in patients with preexisting wounds who come in contact with the 
bacterium via seawater or by handling seafood or who sustain an injury while exposed to 
those sources (6).  
Ecological studies have shown a seasonal pattern wherein the number of V. 
vulnificus in oysters, seawater and sediments increased with warmer temperatures (7-10). 
Predictably, the incidence of wound infections has been found to be positively correlated 
with warm temperatures (11, 12). According to the Cholera and Other Vibrio Illnesses 
Surveillance (COVIS), a Vibrio spp. wound infection is recorded as such when the 
pathogen is cultured from wound and the patient is reported to have sustained a wound or 
40 
 
having a pre-existing one while exposed to marine or estuarine water or by physical 
contact with marine wildlife in the seven days prior illness onset (13).  In most clinical 
cases, patients reported that they had handled seafood prior to the onset of the disease but 
the data do not specify the kind of handled seafood (e.g., shellfish, crustaceans, or fish) (4, 
11, 14). However, the only two documented outbreaks of V. vulnificus involving wound 
infections were attributed to handling farm raised fish in Israel (15) or from injuries 
sustained prior to or during a fishing contest in Texas (12), resulting in 62 and 5 cases, 
respectively. 
Recreational fishing is a main service industry for the US, generating large 
revenues for local coastal communities (16). The northern GoM is a top destination for 
recreational anglers where an estimated >2.8 million anglers participate in more than 7 
million fishing trips annually (17). Although many people recreate in marine and 
estuarine waters and handle fish there, little information is available regarding the 
prevalence and distribution of V. vulnificus in GoM fishes. DePaola et al. (18) 
enumerated the density of V. vulnificus in the intestine of estuarine fishes of Mississippi 
and Alabama, reporting higher levels (105- 108 CFU/g) in fish intestine than in the 
surrounding seawater and sediments; suggesting that fish may be reservoirs for V. 
vulnificus. However, it is noteworthy that those authors did not analyze the external 
surfaces of those fish. Anglers may sustain puncture wounds, lacerations, or bites from 
live fish or they may be incidentally punctured or cut by dead fish during routine 
recreational angling activities (e.g., de-hooking fish, filleting) (19). During these handling 
events, anglers may be exposed to bacteria present in the skin and mucus of their catch. 
41 
 
The purpose of this study was (i) to document the prevalence of V. vulnificus on 
the body surface of a group of estuarine fishes commonly caught by recreational 
fishermen in the GoM and (ii) to characterize the population structure of V. vulnificus in 
those fishes.  
Material and methods 
Sample collection. Sampling began during November 2009 and continued 
through March 2011 at regular intervals except during December 2009 and January and 
February 2010. Sampling sites were selected based on accessibility and considered to be 
representative of public fishing piers in Alabama and Mississippi. Locations included 
Dauphin Island and Gulf Shores in Alabama and Ocean Springs in Mississippi (Figure 3-
1). Table 3-1 summarizes collection dates, locations and numbers of fish analyzed per 
collection event. Seawater surface temperature (at 1 m depth) was measured in situ using 
a mercury-in-glass thermometer (SargentWelch, USA). Salinities were measured with a 
handheld refractometer (Vital Sine? Model SR-6). Fishing efforts lasted between 4 and 
8 h. Fish were captured using standard baited hooks and standard 20 pound-test 
monofilament fishing line on standard spinning reels. Hooked fish were deliberately 
exhausted in ambient water before being raised from the water, secured and suspended in 
air by the angler grasping the leader base or hook shaft, and then touched only by a 
second worker donning sterile surgical gloves and equipped with flamed and ethanol-
rinsed, heavy-gauge scissors. In coordination with raising the exhausted, immobilized 
fish from the water, the second worker approached and immediately excised a portion 
(about 1 cm2) of the dorsal fin and placed the excised tissue in the tube containing 10 ml 
42 
 
of APW. Hence, no sampled fish was placed on any surface or touched by a second 
person before each sample was collected by the surgical-gloved worker. Each sample was 
enriched overnight in APW at room temperature (approximately 25?C). All fish were 
identified according to Carpenter (20), ordinal classification of fishes follows Nelson (21) 
and common names for fishes follows Eschmeyer (22). 
Bacteriological analysis. Upon arrival to the laboratory, 100 ?l of APW cultures 
were plated onto modified cellobiose-polymyxin B-colistin (mCPC) (23) and thiosulfate 
citrate bile salts sucrose (TCBS) (BD, Becton, Dickinson & Co., Franklin Lakes, NJ) agar 
plates and incubated overnight at 30?C. Three colonies displaying the typical V. 
vulnificus morphology (24) were randomly selected from each selective media and re-
isolated on Marine Agar (MA) (BD). Putative isolates recovered from TCBS and mCPC 
agar were subjected to colony dot-blot hybridization according to the protocol described 
by Wright et al. (25). Briefly, putative isolates were cultured in Marine Broth (MB) (BD) 
overnight in a 96-well microtiter plate and approximately 5 ?l of each culture was 
transferred to mCPC using a multiple channel replicator and allowed to grow overnight at 
35 ?C.  Colonies were lifted onto Whatman? 541 filter papers, followed by hybridization 
using an alkaline phosphate-conjugated oligonucleotide (5?-GAGCTGTCACGGCAGT- 
TGGAACCA-3?) (DNATechnology A/S, Risskov, Denmark) that recognizes a specific 
sequence in the V. vulnificus hemolysin gene. Positive isolates were stored at -80 ?C as 
glycerol stocks (MB supplemented with 20% glycerol) for further testing.  
Ascription of V. vulnificus isolates to biotypes. A total of 251 V. vulnificus 
isolates recovered from fish and 8 reference strains (Table 3-2) were included in the 
43 
 
genetic analysis. DNA was extracted from all isolates using standard protocols (26). All 
V. vulnificus isolates were subjected to a multiplex PCR assay for biotype ascription 
according to Sanju?n et al. (2007) (27).  In short, PCR was performed in a 25 ?l reaction 
volume containing 1? PCR buffer, 1.5 mM MgCl2, 200 ?M of each deoxynucleoside, 0.1 
?M of primer vvhA-F (5?-CGCCACCCACTTTCGGGCC-3?) and vvhA-R (5?-CCGC- 
GGTACAGGTTGGCGC-3?), 0.2 ?M of primer Bt2-F (5?-AGAGATGGAAGAAACA- 
GGCG-3?) and Bt2-R (5?-GGACAGATATAAGGGCAAATGG-3?), 1.5 U GoTaq DNA 
polymerase and 1 ?l DNA template (20 ng), and dH2O up to 25 ?l. Unless stated 
otherwise, all molecular reagents were purchased from Promega (Madison, WI, USA). 
PCR reaction was carried out on a Bio-Rad PTC-0200 DNA Engine Cycler (Bio-Rad, 
Valencia, CA) with cycling profile as following: an initial denaturation step 94?C for 10 
min, 35 cycles of 94?C for 30 sec, 60?C for 45 sec and 72?C for 1 min, and a final 
extension step of 72?C for 10 min. The PCR products were analyzed by electrophoresis 
on a 2% agarose gel and visualized with UV light by staining with ethidium bromide. 
Ascription to biotype 1-3 or biotype 2 was based on amplicon(s) size (the method does 
not discriminate between biotypes 1 and 3). 
16S-Restriction Fragment Length Polymorphism (RFLP) typing. 16S rrn 
genotype was determined by RFLP according to Nilsson et al. (28). A 492 bp region of 
16S rRNA gene of V. vulnificus was amplified using primers UFUL (5?-GCCTAACAC- 
ATGCAAGTCGA-3?) and URUL (5?-CGTATTACCGCGGCTGCTGG-3?). PCR 
reaction was performed as described above with the only difference being the annealing 
temperature of 57 ?C.  PCR products were verified by 2% agarose gel electrophoresis. 
44 
 
Restriction endonuclease digestion of amplified product was performed in 20 ?l reaction 
including 10 ?l of amplicon, 2 ?l of 10? buffer B, 0.2 ?l of acetylated BSA (10 ?g/ml), 
0.5 ?l of AluI (10U/ ?l) and sterile dH2O up to 20 ?l. Digestion was carried at 37?C for 2 
hr after which DNA fragments were separated by electrophoresis on 4% agarose gel. 16S 
rrn types were ascribed based on profiles described by Nilsson et al. (29) 
Amplified Fragment Length Polymorphism (AFLP). AFLP reactions were 
carried out as described by Arias et al. (30). Briefly, 100 ng of RNase-treated genomic 
DNA was double digested with TaqI and HindIII. Following digestion, specific TaqI and 
HindIII adaptors were ligated to the restriction fragments and subsequently amplified by 
PCR using primers T000 (5?CGATGAGTCCTGACCGAA-3?) and H00A (5?-
GAACTGCGTACCAGCTTA-3?), selective bases at the3?end are underlined.  HindIII 
primer (H00A) was labeled with an IR700 fluorochrome from LI-COR (LI-COR, 
Lincoln, NE, USA). PCR amplifications were performed with the following cycle profile: 
cycle 1, 60 s at 94?C, 30 s 65?C, and 60 s at 72?C; cycles 2 to 12, 30 s at 94?C, 30 s at 
annealing temperatures 0.7?C lower than that used for each previous cycle, starting at 
64.3?C, and 60 s at 72?C; cycles 13 to 24, 30 s at 94?C, 30 s at 56?C, and 60 s at 72?C. 
After completion of the cycling program, 5 ?l of AFLP Blue Stop Solution (LI-COR) 
was added to the reaction mixtures. Prior to gel loading, the samples were heated for 5 m 
at 94?C then rapidly cooled on ice to prevent reannealing. The PCR products were 
electrophoresed on the NEN Global Edition IR2 DNA Analyzer (LI-COR) following 
manufacturer?s instructions.  
45 
 
Data analysis. The BioNumerics 6.6 software suite (Applied Maths, Saint 
Martens-Latem, Belgium) was used for AFLP data analysis. Pairwise similarities were 
calculated using Pearson correlation coefficient with a 0.5% optimization. Using the 
similarity matrix as an input, a dendrogram was constructed with arithmetic averages 
algorithm (UPGMA). The Jackknife group separation method (based on maximal 
similarities between isolates) was used to assess the fidelity of the clustering analysis. 
Only bands within the range of 100-530 bp and with at least 8% of minimum profiling 
were considered in the analysis. Transversal clustering was performed based on the 
swapped data matrix of profile similarities (fingerprint patterns, horizontal cluster) and 
characters (band classes, vertical cluster). In transversal clustering, the isolates are 
grouped by fingerprint patterns (similarities based on band position and intensity, Pearson 
correlation coefficient), while the character are sorted by means of value of band classes 
(0 absent and 1 present, Jaccard coefficient). The diversity of the V. vulnificus isolates 
from each fish species was compared by generating rarefaction curves using the software 
Diversity version 1.4 (Hunt Mountain Software, Athens, GA). 
The SAS Software 9.2 version (SAS Institute, Cary, N.C.) was applied to analyze 
the relationship between environmental factors and the percentages of fish harboring 
Vibrio vulnificus (Vv+%). The percentage, namely, is the frequency of fish fin clips that 
yielded confirmed V. vulnificus isolates and was calculated for each sampling event. 
Pearson correlation coefficient was used to estimate the relationship between percentages 
of Vv+ fish and environmental factors (water temperature and salinity). A general linear 
46 
 
regression analysis was used to quantify the trends observed between Vv+ fish and 
variation in salinity and water temperature.  
Results 
Prevalence of V. vulnificus in fish. A collection of 244 fish were randomly 
sampled during 12 sampling trips. Overall, 90 (37%) individual fish (each represented by 
an excised dorsal fin sample) yielded V. vulnificus isolates, and therefore were considered 
positive (Vv+) for harboring the bacterium (Table 3-1). The prevalence of V. vulnificus in 
fish varied between sampling events from 0-78%. Vibrio vulnificus was recovered from a 
large diversity of fish species (Table 3-3) with prevalence ranging from 0% to 58%. 
However, the numbers of analyzed fish were not equal across all species due to variable 
capture success. Because of this limitation, comparing the prevalence of V. vulnificus 
among fish species was not statistically feasible. Vibrio vulnificus was documented from 
any fish species wherein more than 3 individuals of that species were sampled. The order 
Perciformes was best represented in our sampling, with 63 of 171 (37%) individual fish 
harboring V. vulnificus. Among fishes wherein >10 individuals were analyzed, Atlantic 
croaker and southern flounder had the highest prevalence at 58% and 53%, respectively, 
while striped mullet had the lowest at 25%. 
Environmental parameters fluctuated as expected during the study, with water 
temperatures ranging from 16?C to 31 ?C and salinity from 8 ppt to 33 ppt  (Figure 3-2). 
Percentage of Vv+ fish and salinity were inversely correlated (Pearson?s correlation 
coefficient r = -0.91077, n=12, p<0.0001). By contrast, a positive correlation was found 
between Vv+ fish and water temperature (Pearson?s correlation coefficient r = 0.62481, 
47 
 
n=12, p=0.0298). General linear models were applied to describe the changes in 
percentages of Vv+ fish and the environmental factors (salinity and water temperature) 
analyzed in the study. The linear regression model identified these two environmental 
factors as having significant effect on the percentage of Vv+ fish (Vv+%). The model 
with two significant factors that best fits the data is as follows: Vv+% = 0.5930 - 0.02818 
? salinity+ 0.01406 ? water temperature. Parameter estimate statistics are included in 
Table 3-4 and the surface plot for the model equation is shown in Figure 3-3. The 
coefficient of determination (r2) for the model was 0.9041, which denotes that 90% of the 
observed variation is explained by the independent variables.  When an interaction term 
between salinity and water temperature was introduced to the model, the p-value 
indicated the interaction term was not significant (p-value= 0.9571). Essentially, this 
indicates that the number of Vv+ fish was independently affected by salinity and 
temperature but that both parameters were not linked to each other. In addition, and to 
test if the collections sites influenced the percentages of Vv+ fish, two dummy variables 
(S1 and S2) accounting for two of the collection areas (Dauphin Island and Ocean 
Springs) were brought into the model. The result of t-test on parameters S1 (p-value= 
0.6413) and S2 (p-value= 0.9199) indicated that site was not significant. 
Population structure of V. vulnificus on fish. Out of more than 500 putative V. 
vulnificus colonies recovered from selective media, 270 isolates were positive by colony 
dot-blot hybridization and, out of those, 251 were positive by vvhA-specific PCR. None 
of 251 isolates were Biotype 2 based on multiplex PCR (27). 16S-RFLP typing classified 
the isolates in 3 types (16S type A, B and AB) as previously reported (29). According to 
48 
 
this classification method, 157 isolates (62%) were 16S type A, 72 (29%) were type B 
and 22 (9%) were type AB. Representatives of 16S type A and B were recovered in every 
sampling, except for March-2010 where only 16S type A isolates were obtained. Figure 
3-4 shows the temporal distribution of 16S types. The percentage of 16S type B isolates 
varied from 0% to 50% but no clear correlation between salinity, water temperature or 
fish species and 16S type could be inferred from our data. 
Eight additional strains, proved to be virulent in a mouse model (31), were 
included as references in the AFLP analysis. All but 12 fish isolates were typeable by 
AFLP (untypeable isolates consistently produced profiles of weak intensity). AFLP 
profiles were highly informative with an average 95 bands per profile ranging from 50 to 
700 bp. In order to test reproducibility of the AFLP method, we performed 3 independent 
AFLP experiments using 10 fish isolates (data not shown). Based on the variability 
observed, we selected 90% as our threshold for considering an AFLP type unique as 
previously described (32). Profiles displaying more than 90% similarity were ascribed to 
the same AFLP type while similarities lower than 90% indicate different AFLP profiles. 
A total of 121 unique AFLP types were defined within the 239 isolates of V. vulnificus 
(Table 3-3). Rarefaction curves were generated for each fish species in where 10 or more 
isolates from the same fish species had been typed (data not shown). In all cases, the 
slope of the curves was similar and the number of AFLP types observed linearly 
increased with the total number of V. vulnificus analyzed. These results suggest that the 
genetic diversity of the isolates was similar in all fishes.  
49 
 
Figure 3-5 displays a multiscaling dimensional (MSD) analysis of the similarity 
among all isolates derived from the AFLP cluster analysis. Two non-overlapping clusters 
could be clearly delineated that correlated with 16S type ascription. Cluster I grouped all 
16S type B isolates at 70% similarity (see Figure 3-6), while cluster II consisted of 148 
isolates including all 16S type A and type AB at 71% similarity. All V. vulnificus isolates 
shared a similarity of 65% or higher. Cluster I and cluster II were comprised of 36 and 85 
unique AFLP types, respectively. Group separation statistics based on Jackknife analysis 
confirmed the statistically significant correlation between AFLP clusters I and II and 16 
types B and A-AB. Resampling of the data showed a 100% agreement between assigned 
16S type B and randomly selected subgroups (for types A and AB the agreement was 96% 
and 83%, respectively). 
Overall, isolates did not cluster based on collection date, geographic location or 
fish species (data not shown). Moreover, V. vulnificus isolates recovered from the same 
individual fish were not more related to each other than to those isolates from other fish. 
As an example of how V. vulnificus types did not show any specificity for fish species, 
Atlantic croaker isolates (the fish species that yielded the most V. vulnificus isolates) 
were distributed across the entire AFLP-based dendrogram (Figure 3-6).  
 The band matching analysis revealed a total of 119 polymorphic bands 
responsible for AFLP cluster ascription out of 123 total observed bands. Transversal 
clustering identified the AFLP bands, or markers, characteristic of 16S type B isolates. In 
Figure 3-6, the isolates were clustered based on band profiles, while the character was 
grouped according to values in the band Table 3-which reflected band classes of 
50 
 
individual band profiles (1 present, 0 absent). In the two-dimensional clustering (data not 
shown), the isolates and band classes were arranged according to their relatedness. For 
example, band classes (labeled by band length) 116 bp, 156 bp, 200 bp, 253 bp, 343 bp, 
386 bp, and 491 bp were more abundant in Cluster I (16S type B isolates). These band 
classes are AFLP markers for 16S type B isolates (highlighted area in Figure 3-6).  
Discussion 
Few studies have documented prevalence and distribution of V. vulnificus in wild 
fish (18), despite the fact that wound infections caused by this bacterium have been 
reported in fishermen worldwide  (11, 15, 33, 34). In the US, wound infections attributed 
to V. vulnificus have been reported from US GoM Coast states since the species was first 
described in 1976 (5, 14, 35, 36). Approximately 30 confirmed cases occurred annually 
in the GoM coast region (13), and most of them purportedly linked to exposure to 
seawater or seafood (11). However, it is not possible to account for how many cases of 
wound infections were acquired by direct contact with fish based on epidemiological 
studies. Our data showed that V. vulnificus occurs on the fins of fishes from the northern 
GoM, with an overall prevalence of 37% although prevalence increased to 69% during 
the warmer months (June-September). It is noteworthy that during these months, V. 
vulnificus was recovered at all sampling events. This pathogen was isolated from a wide 
taxonomic and phylogenetic spectrum of fishes, including 16 species representing 15 
genera, 8 families, and 6 orders (Table 3-3). Interestingly, the highest prevalence of V. 
vulnificus was found in two bottom feeder fish (Atlantic croaker and southern flounder), 
which have been shown to have high densities of this bacterium in their gut (18). 
51 
 
However, our findings suggest that V. vulnificus is a transient member of the microbiota 
(37) associated with fish fins surfaces because it was not found on each fish even when 
several individuals from the same species were collected at the same time.  
The occurrence of V. vulnificus on fish fins may be higher than that reported here 
since we used a culture-based method to estimate prevalence. The success rate of 
recovering environmental bacteria with culture-based methods varies depending on 
several factors, but it tends to underestimate original densities. For instance, both APW 
and mCPC have been reported to favor the growth of V. vulnificus biotype 1 over biotype 
2 (38). Similarly, Chase and Harwood (39) showed than biotype 1 grows better than 
biotypes 2 and 3 under identical conditions. These factors may have negatively 
influenced the recovery of biotype 2 in our study. In addition, culture methods disallow 
for recovery of viable but not culturable (VBNC) forms of V. vulnificus, a well-described 
life stage of this bacterium (40) although the abundance of VBNC forms in the 
environment is not well known (9). The protocol used herein (enrichment in APW 
followed by plating onto mCPC) could have preferentially enriched for V. vulnificus 
biotype 1 but, overall, it provided a simple and inexpensive methodology for successfully 
recovering V. vulnificus from fish.  
The frequency of detection of V. vulnificus in fish was a factor of salinity and 
temperature. These environmental factors are known to influence V. vulnificus abundance 
and distribution in the marine environment (2, 7, 41).  Temperature is positively 
correlated with V. vulnificus presence while salinity is negatively correlated. However, it 
has been suggested that salinity becomes the main factor influencing the abundance of V. 
52 
 
vulnificus in oysters when salinities exceed 30 ppt, irrespectively of water temperature 
(42). Our field data support this hypothesis since we failed to recover V. vulnificus from 
fish when salinities were higher than 29 ppt even when water temperatures were above 
18?C.  Based on our data, prevalence of V. vulnificus in fish fins was more affected by 
salinity than by water temperature, and this was particularly noticeable from June through 
September in where water temperatures remained over 25?C. Our model predicts that 
salinity accounted for 83% of the variability of Vv+ fish; whereas, temperature explained 
only 7% of the variability. Nevertheless, this model may not be accurate when water 
temperatures and salinities are outside the ranges observed during our study.  
Epidemiological data suggest that most V. vulnificus strains must present little risk 
to the susceptible population because of its abundance in marine samples and the small 
number of clinical cases observed per year (3, 43, 44). The first marker to be used as an 
indicator for virulence was a polymorphism present in the 16S rRNA gene that classifies 
the isolates as 16S type A and 16S type B; in addition, some strains can present both 16S 
alleles and are classified as 16S type AB. Based on epidemiological data, up to 75% of 
clinical V. vulnificus isolates are 16S type B (this percentage increased up to 94% when 
clinical fatalities were considered) (29). However, Thiaville et al. (31) showed that V. 
vulnificus genotypes were correlated with but did not predict virulence in mice using a 
skin-infection model. Isolates of V. vulnificus are typically referred to as of ?clinical 
origin? (including primary septicemia and wound infection) or ?environmental origin? 
making it difficult to determine what percentage of wound infections are caused by 16S 
type B (29, 31, 45). The majority (62%) of the fish isolates recovered in this study were 
53 
 
classified as 16S type A (up to 71% if we include type AB) but at 29%, the number of 
type B isolates was higher than expected based on previous studies from the same area 
(29, 46, 47). Although significant correlations were not noted with any environmental 
parameter, the highest proportion of type B relative to A and AB was observed during the 
warmest months, which is consistent with similar observations of V. vulnificus 
populations in Galveston Bay, TX (48).  
AFLP revealed two main genetic groups within the species that correlated well 
with 16S type ascription. The division of the species into two main genetic groups has 
been shown previously using up to 14 different loci including markers for virulence (44, 
49, 50) and prompted Rosche et al. (2010) to hypothesize that this pattern is indicative of 
speciation. Our results, derived from a whole-genome fingerprinting method, are in 
agreement with this hypothesis. In terms of strain characterization, AFLP provided a high 
level of resolution and confirmed the high genetic diversity present in the species with a 
calculated Simpson?s index of diversity of 0.991 (51).  This value is in range with that 
previously reported by Arias et al. (52) when isolates from a broader geographic area 
were characterized by AFLP. In some cases, isolates recovered from different fish species 
(i.e., red drum and Atlantic croaker) and collected from different locations at different 
times, shared highly similar AFPL profiles (at 95% similarity), while in other examples 
isolates recovered from the same fish (i.e., Atlantic croaker) clustered in distinct AFLP 
types (Table 3-3). Vibrio vulnificus isolates associated with fish were highly 
heterogeneous, and we could not determine spatiotemporal- or fish-specific patterns that 
linked V. vulnificus types with environmental parameters or fish species. The inclusion of 
54 
 
reference strains in the AFLP analysis further confirmed the lack of uniqueness among 
fish isolates since some of them shared high similarities with reference strains including 
those of clinical origin. This indicates that fish from the nearshore waters of the northern 
GoM harbor V. vulnificus genotypes of great virulent potential for humans. 
In summary, this study presents new data on the prevalence of V. vulnificus on 
fish and presents a novel statistical model for predicting its occurrence. Our data indicate 
that this bacterium has a broad, seasonally-dependent distribution among fishes ranging 
in coastal areas of the north-central GoM. The high genetic heterogeneity found among V. 
vulnificus isolates from fish resembles those found in other samples from the same area 
(including Eastern oysters), but the population structure differs as the percentage of 16S 
type B from fish was higher than expected. Vibrio vulnificus appears to be a transient 
member of the fish surface microbiota as it was affected by changes in environmental 
parameters, mainly salinity. Potentially pathogenic strains of V. vulnificus were recovered 
from fish, based on 16S ascription and AFLP similarity with clinical strains. Although 
reported cases of wound infections caused by this pathogen are clearly low proportional 
to the numbers of anglers, fish captured, and fishing trips taken in the GoM, our 
distributional data show that fishermen could likely be inoculated by V. vulnificus during 
fish handling and processing if punctured, cut, or abraded, particularly if the fish was 
captured in an estuary.  
Acknowledgements 
This work was funded by the National Oceanic and Atmospheric Administration 
(NOAA-NA08NMF4720545), Marine Environmental Science Consortium, GoM 
55 
 
Research Institute, and National Science Foundation (NSF DEB-1048523). Zhen Tao is 
the recipient of graduate research fellowship funded by the Chinese Scholarship Council 
and Ocean University of China. 
  
56 
 
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61 
 
Table 3-1. Temporal and spatial distribution of fishing efforts summarizing number of 
fish analyzed and number of fish positive for Vibrio vulnificus (Vv+). 
Date (mm/dd/yy) Sites No. of fish No. of Vv+ fish 
11-17-09 OSa 19 9 
03-26-10 OS 30 10 
06-02-10 DIb 20 12 
06-16-10 DI 25 18 
07-18-10 OS 23 18 
08-18-10 DI 23 6 
09/18/10 OS 18 11 
09/18/10 DI 10 4 
10-16-10 GSc 25 0 
11-12-10 OS 18 0 
11-28-2010 OS 25 2 
3-17-2011 GS 8 0 
Total  244 90 
a Ocean Springs, Mississippi 
b Dauphin Island, Alabama 
c Gulf Shores, Alabama 
  
62 
 
Table 3-2.  Reference strains. 
In this 
study 
Straina Source Origin Date 16Sd vcge Virulencef 
R-1 CDC 9060-96 Clinicalb TX 1996 B C Virulent 
R-2 CDC 9070-96 Clinicalc TX 1996 B C Virulent 
R-3 ATL-9824 Clinicalc TX 1994 B C Virulent 
R-4 98-640-DP-E9 Oyster LA 1998 A E Virulent 
R-5 99-625 DP-D8 Oyster TX 1999 AB E Virulent 
R-6 246-0058  Clinical FL - A E Virulent 
R-7 99-609 DP-A4 Oyster OR 1999 A E Virulent 
R-8 99-537 DP-G7 Oyster MD 1999 A E Virulent 
a For a full description of these strains see Thiaville et al. (2011) 
b Fatal outcome. 
c patient recovered from infection. 
d Typed according to Nilsson et al. (2003) 
e Typed according to Rosche et al. (2005)  
f Based on Thiaville et al. (2011)  
 
 
 
 
 
 
63 
 
Table 3-3. Occurrence of Vibrio vulnificus in fish collected during the study  
Fish species (order: family), common name No. Vv+ fish/  No. total fish 
No. of  
recovered  
 isolates 
AFLP Typea 
Dasyatis sabina ( Myliobatiformes: Dasyatidae ), Atlantic 
stingray 1/3 3  55 
Elops saurus (Elopiformes: Elopidae), ladyfish 3/9 11  6, 36, 62, 64, 79, 95, 100 
Brevoortia patronus (Clupeiformes: Clupeidae), Gulf 
menhaden 0/1 0  - 
Dorosoma petenense (Clupeiformes: Clupeidae), threadfin shad 0/1 0  - 
Bagre marinus (Siluriformes: Ariidae ), gafftopsail sea catfish  2/2 2  2 
Ariopsis felis (Siluriformes: Ariidae ), hardhead sea catfish  6/9 16  7, 11, 12, 14, 26, 37, 43, 59, 94, 98 
Opsanus beta (Batrachoidiformes:  Batrachoididae), Gulf 
toadfish  0/1 0  - 
Mugil cephalus (Mugiliformes: Mugilidae), flathead grey 
mullet  6/24 16  5,  82,  84,  86,  111,  117,  118 
Strongylura marina (Beloniformes: Belonidae), Atlantic 
needlefish 0/2 0  - 
Prionotus tribulus (Scorpaeniformes: Triglidae), bighead 
searobin  0/2 0  - 
Echeneis naucrates (Perciformes: Echeneidae), live 
sharksucker 0/2 0  - 
Caranx sp. (Perciformes: Carangidae) 0/1 0  - 
Selene vomer (Perciformes: Carangidae), lookdown  0/1 0  - 
Orthopristis chrysoptera (Perciformes: Haemulidae), pigfish  2/7 9  21, 80 
Archosargus probatocephalus (Perciformes: Sparidae ), 
sheepshead  1/6 2  116 
Lagodon rhomboides  (Perciformes: Sparidae ), pinfish  7/23 18  1, 4, 25, 28, 39, 47, 69, 73, 82, 97, 102, 103 
a only 239 out of 251 isolates were typeable by AFLP. 
  
 
64 
 
Table 3-3. Continued  
Fish species (order: family), common name No. Vv+ fish/  No. total fish 
No. of  
recovered  
 isolates 
AFLP Typea 
Bairdiella chrysoura (Perciformes: Sciaenidae), silver perch  5/24 13  60, 81, 85, 94, 99, 100, 106 
Cynoscion arenarius (Perciformes: Sciaenidae), sand weakfish 11/28 32  13, 17, 22, 26, 29, 43, 44, 50, 52,  57, 63, 75, 79, 89, 94, 96, 117, 120 
Cynoscion nebulosus (Perciformes: Sciaenidae), spotted 
weakfish 7/16 23  10, 15, 17, 33, 34, 35, 45, 66, 90, 91, 121 
Leiostomus xanthurus (Perciformes: Sciaenidae), spot croaker  0/3 0  - 
Menticirrhus sp. (Perciformes: Sciaenidae) 2/9 8  36, 51, 109 
Micropogonias undulatus (Perciformes: Sciaenidae), Atlantic 
croaker 22/38 67  
3, 8, 9, 10, 11, 15, 16, 19, 20, 23, 24, 38, 
40,  
41, 46, 48, 50, 54, 58, 61, 65, 67, 68, 70, 
72,  
77, 78, 82, 87, 92, 101, 104, 105, 108, 113 
Pogonias cromis (Perciformes: Sciaenidae), black drum  0/1 0  - 
Sciaenops ocellatus (Perciformes: Sciaenidae), red drum  3/5 7  15, 18, 64, 85, 112 
Chaetodipterus faber (Perciformes: Ephippidae), Atlantic 
spadefish  3/3 8  27, 28, 49, 71, 114 
Scomberomorus cavalla (Perciformes: Scombridae), king 
mackerel 0/1 0  - 
Scomberomorus maculatus (Perciformes: Scombridae), 
Atlantic Spanish mackerel 0/3 0  - 
Paralichthys lethostigma (Pleuronectiformes: Paralichthyidae), 
southern flounder 9/17 16  12, 31, 74, 76, 81, 83, 88, 93 
 Total 90/242 251   
a only 239 out of 251 isolates were typeable by AFLP.
 
65 
 
Table 3-4. Statistical values for the multiple linear regression model. 
Parameter Estimate Standard Error t -value Pr>|t| 
Intercept 0.5930 0.1786 3.3200 0.0089 
Salinity -0.0282 0.0041 -6.9500 <.0001 
Water Temperature 0.0141 0.0053 2.6500 0.0266 
 
  
 
66 
 
 
Figure 3-1. North-central GoM showing collecting sites: DI, Dauphin Island, GS, Gulf 
Shores, and OS, Ocean Springs. MS, Mississippi; AL, Alabama.  
  
 
67 
 
 
Figure 3-2. Distribution of Vibrio vulnificus-positive fish throughout the study (bars) in 
relationship to salinity and water temperature. 
 
  
 
68 
 
 
Figure 3-3. Surface plot for model equation: (Vv+)% = 0.5930 - 0.02818 ? salinity + 
0.01406 ? water temperature 
  
 
69 
 
 
 
Figure 3-4. Distribution of Vibrio vulnificus 16S type A, AB, and B across the study. 
  
 
70 
 
 
Figure 3-5. Multidimensional Scaling (MDS) representation of the similarity matrix 
generated by AFLP cluster analysis. Each of 251 V. vulnificus isolates is represented by a 
dot and the distance between dots represents relatedness obtained from the similarity 
matrix. Isolates are colored based on origin (fish family or reference strains). Dotted lines 
highlight the two main clusters observed in the analysis. Cluster II groups all V. vulnificus 
16S type A and AB and cluster I groups 16S type B. 
  
 
71 
 
 
 
72 
 
Figure 3-6. Clustering analysis based on AFLP fingerprint analysis. Composite matrix of 
band classes based on transversal clustering of V. vulnificus AFLP fingerprints. Scale 
represent percent of similarity based on Person?s product moment correlation coefficient. 
The two main observed clusters (I and II) are delineated at 70% similarity. AFLP markers 
identified by transversal clustering and used for cluster delineation are framed. Reference 
strains are indicated with an asterisk. Vibrio vulnificus strains isolated from Atlantic 
croaker are marked with a dot. 
  
 
73 
 
 
 
 
 
 
CHAPTER 4. OCCURRENCE OF VIBRIO VULNIFICUS AND V. 
PARAHAEMOLYTICUS IN BAIT SHRIMP FROM ALABAMA AND MISSISSIPPI 
 
 
Abstract 
 The prevalence of the human pathogens Vibrio parahaemolyticus and V. vulnificus in 
bait shrimp species was analyzed in this study. Shrimps were obtained from commercial bait 
shops along the coast of Alabama and Mississippi from March 2010 to January 2011. The 
levels of both Vibrio species were under the detection limit in November and March, and 
reached the highest level during the summer months (>1.1?105 MPN per gram shrimp tissue 
for both Vibrio species). Randomly selected isolates of both species were type using 
virulence-related typing schemes. Out of 26 V. vulnificus isolates analyzed, 69% were 16S 
type B and vcgC type indicating the virulence potential of the isolates. By contrast, none of 
the 33 V. parahaemolyticus isolates was positive for tdh and trh hemolysin genes. This study 
confirms bait shrimp as potential vector for V. vulnificus infections. 
Introduction 
 Bacteria of the genus Vibrio are key constituents of the microbiota of estuarine and 
coastal environments of the GoM in where they (1). Although most Vibrio species have 
never been associated with clinical cases, approximately a dozen Vibrio species have been 
demonstrated to be pathogenic to animals or humans (2).  In the United States, according to 
CDC data from 1996 to 2010, Vibrio parahaemolyticus and V. vulnificus are the leading 
cause of Vibrio infections in humans (3). There are clinical manifestations of illness caused 
 
74 
 
by V. parahaemolyticus and V. vulnificus: gastroenteritis, wound infections, and septicemia 
(4). Septicemia, the most severe illness, and gastroenteritis are typically contracted by 
ingestion of raw or poorly cooked seafood while wound infections occur when compromised 
skin is exposed to seafood or seawater (5).  
 Many studies have focused on the importance of V. parahaemolyticus and V. 
vulnificus as foodborne pathogens in the US (6, 7) and risk assessments plans have been 
developed for both species (8-12). In addition, they also play a substantial role in 
nonfoodborne infections associated with activities such as seafood handling, swimming, 
wading and recreational fishing (13). However, less information is known about the risk 
factors of nonfoodborne Vibrio infections involved in recreational activities, including 
fishing. In present study, we investigated the distribution of V. vulnificus and V. 
parahaemolyticus in bait shrimp (Penaeus spp.) sold at coastal bait shops in Alabama and 
Mississippi. To enumerate these bacteria in shrimp, we used the most probable number 
(MPN) procedure according to Bacterial Analytical Manual (14).  Furthermore, we examined 
the virulence potential of V. vulnificus and V. parahaemolyticus isolates recovered from bait 
shrimp using current typing schemes (9, 15, 16). 
Material and methods 
 Sample collection. From March 2010 to January 2011 (except April, May and 
December in 2010), bait shrimps were purchased monthly from local baits shops, one located 
in Dauphin Island , Alabama (DI, AL) and one in Ocean Spring , Mississippi (OS, MS). Both 
facilities were on the water and shrimp were kept in holding tanks under flow-through 
conditions (pumps were used to circulate water continuously from Mobile bay (DI, AL) or 
 
75 
 
the intracoastal water way OS, MS. The shop owners decline to disclose the amount of time 
they kept the shrimp in those systems but they confirmed they were caught offshore using 
traditional trawling methods (17). One additional sample, July 2010, was collected from a 
marsh area in OS by using a castnet. Shrimp samples were transported to the laboratory with 
aeration within 5 hours of collection.  
 Sample analysis.  Samples were analyzed following the 3-tube Most Probable 
Number (MPN) method described in the U.S. Food and Drug Administration?s 
Bacteriological Analytical Manual (BAM) (14). Briefly, 12 shrimps were patted-dried, 
weighted, diluted 1:1 in phosphate-buffered saline (PBS), and homogenized for 90 sec in a 
blender (Osterizer?, USA) at maximum speed. The first 1:10 dilution corresponded to 20 g of 
the homogenate into 80 ml sterile PBS. Samples were subsequently diluted in PBS until 10-5 
g. Samples were incubated at 35?C for 18 h. For V. vulnificus enumeration, an aliquot of each 
tube was streaked onto modified Colistin Polymixin-B Cellobiose (mCPC) agar plates (18) 
and incubated at 35?C for 18 h. Three putative colonies from each plate were transferred to a 
V. vulnificus Agar (VVA), and incubated at 35?C for 18 h. Colonies on VVA were lifted onto 
Whatman 541 filter papers, and confirmed by alkaline phosphate-conjugated oligonucleotide 
probe (5?-GAGC TGTCACGGCAGTTGGAACCA-3?) (DNATechnology A/S, Risskov, 
Denmark) that recognizes a specific sequence in the V. vulnificus hemolysin gene (vvhA) 
(19).  MPN of V. vulnificus MPN?g?1 was subsequently determined using the chart located in 
the BAM to interpret the results (20). The enumeration of V. parahaemolyticus was 
performed following similar protocols (14) with the exception of selective media and specific 
probe. For isolation of V. parahaemolyticus, the selective medium used was thiosulfate-
 
76 
 
citrate-bile salts-sucrose agar (TCBS) (BD, Becton, Dickinson & Co., Franklin Lakes, NJ). 
Putative V. parahaemolyticus colonies were subsequently incubated on T1N3 agar (1% 
tryptone, 3% NaCl, and 1.5% agar). A hybridization probe (5?-AAAGCGGAT- 
TATGCAGAAGCACT-3?) targeting the thermolabile hemolysin (tlh) gene was used to 
confirm the V. parahaemolyticus isolates (21). 
 Isolate selection.  A subset of confirmed V. vulnificus and V. parahaemolyticus 
isolates were arbitrarily selected for genetic typing.  A total of 26 V. vulnificus and 36 V. 
parahaemolyticus isolates were stored in semisolid marine agar (marine broth [BD] with 
0.3% agar) at room temperature until further use. Template DNA was prepared using a rapid 
boiling method as follows. Five colonies from each isolate  were selected from a 24-hour 
culture on Marine Agar (BD) plates, and resuspended in a centrifuge tube with 100 ?l sterile 
distilled H2O. Proteinase K was added to the cell suspension to a final concentration of 30 
unit/?l. After 20 min digestion at 55 ?C, the lysate was heated to 100 ?C for 15 min, and spun 
down at 15,000 g for 5 min. The supernatant was transferred to a new tube and used as 
template DNA. Species-specific PCR reactions using primers listed in Table 4- 1 were 
applied to selected isolates to further confirmed their ascription to species.  
 Vibrio vulnificus typing. All V. vulnificus isolates were ascribed to biotype as per by 
Sanju?n et al. (22). Assignment to the virulence correlated gene types vcgE (environmental 
type) and vcgC (clinical type) were performed as described by Rosche et al. (16). 16S rRNA 
genotyping was performed by amplifying a conserved fragment of the 16S rRNA gene 
followed by restriction fragment length polymorphisms (RFLP) analysis according to Nilsson 
et al (15).   
 
77 
 
 Vibrio parahaemolyticus PCR analysis. All V. parahaemolyticus isolates were 
screened for tdh and trh genes in a multiplex PCR assay. PCR was performed in a 25-?l 
reaction volume containing 1? PCR buffer, 1.5 mM MgCl2, 200 ?M each deoxynucleoside, 
0.2 ?M primers Bt2-F  and Bt2-R , 1.5 U GoTaq DNA polymerase, 1 ?l DNA template, and 
distilled H2O (dH2O) up to 25 ?l. Primer concentrations varied depending on the reaction 
(Table 4- 1). Unless stated otherwise, all molecular reagents were purchased from Promega 
(Madison, WI).  PCR was carried out on a Bio-Rad PTC-0200 DNA engine cycler (Bio-Rad, 
Valencia, CA) with the following cycling profile: initial denaturation at 94?C for 3 min, 
followed by 35 cycles as shown in Table 4- 1, and ended with a final extension at 72? for 10 
min. PCR products was confirmed by determining the presence and sizes of products using 
agarose gel electrophoresis. DNA in gels was stained with ethidium bromide and visualized 
with UV light in a UVP BioSpectrum 300 Imaging system (city, state).  An additional step 
was involved in RFLP for the V. vulnificus 16S rRNA genotyping. Restriction endonuclease 
digestion of amplified product was performed in a 20?l reaction volume including 10 ?l of 
amplicon, 2 ?l of 10? restriction buffer B (Promega), 0.2 ?l of acetylated bovine serum 
albumin (10 ?g?ml-1), 0.5 ?l of AluI (10 U??l-1), and sterile dH2O up to 20 ?l. Digestion was 
carried at 37?C for 2 h, after which DNA fragments were separated by electrophoresis on a 
4% agarose gel. 
Results and discussion 
 Recreational fishing is one major industry in GoM States and generate large revenues 
for local communities (23). For this region,  2.7 million fishermen participated in the 
recreational fishing and took nearly 22 million fishing trips in 2010 (24). Live shrimps are 
 
78 
 
popular bait along the US GoM coast where recreational fishermen typically purchased them 
at their local bait shops. For instance, recreational anglers aiming at spotted seatrout 
(Cynoscion nebulosus), red drum (Sciaenops ocellata) and flounder (Paralichthys spp.) 
prefer to use live bait shrimp. Therefore, handling live bait shrimp is common for a large 
number of saltwater fishermen. Epidemiologically, anglers may be incidentally punctured or 
cut by bait shrimp while handling or hooking the shrimp (25). During these handling events, 
anglers may be exposed to bacteria present in the external or internal organs of the shrimp. 
Thus, the objective of this study was to assess if bait shrimp could act as vector for 
pathogenic vibrios. 
 Our results showed that both species of pathogenic vibrios occur at high levels in bait 
shrimp. Eight out of 13 and 9 out of 13 samples tested contained >104 MPN/g of V. vulnificus 
and V. parahaemolyticus, respectively. Only 4 samples had non-detectable (<3 MPN/g) of 
either pathogen. Not surprisingly, negative samples occurred during cold temperature months 
(January and March) when levels of V. vulnificus and V. parahaemolyticus are low in water, 
sediments, and shellfish (8, 26).Conversely, high levels (>105 MPN/g) were obtained in all 
samples tested from June through August. However, levels of both species remained high 
(>104 MPN/g) during transition months (October and November).   
 All randomly selected positive V. vulnificus isolates (n=26) were confirmed by 
specific PCR assay. All these isolates were assigned to biotype1or 3 strains (Table 4- 3).  
Thirty one percent of V. vulnificus isolates were ascribed to 16S type A, and 69 % to 16S 
type B. Results of vcg genotyping were in accordance with 16S typing as all vcgC isolates 
were 16S type B and all vcgE isolates were identified as 16S type A. In this study, a high 
 
79 
 
percent (69%) of V. vulnificus isolates recovered from shrimp were classified as 16S type 
B/vcgC. The species V. vulnificus can be divided into distinct genetic groups according to 
two typing schemes. The first method uses a polymorphism present in the 16S rRNA gene 
that divides the species into 16S A, B and the hybrid AB types (15, 27, 28). The second 
methods, uses a marker referred to as ?virulence correlated gene? that classifies the isolates 
into two types: vcgC type and vcgE type (16, 29).  In general, 16S type A and AB isolates are 
vcgE type while 16S type B strain are vcgC type (30).  The justification for these typing 
schemes derives from epidemiological data that showed a strong correlation between clinical 
isolates and 16S and vcg types. While most (>80%) of clinical isolates (>95% when only 
fatality cases are considered) are 16S type B (vcgC) the majority of the environmental 
isolates are 16S type A and AB (vcgE ) (15, 29, 31). Our data contradicts previous study in 
that most of our isolates were 16S type B (vcgC), which denotes a risk of contracting V. 
vulnificus wound infection by handling bait shrimp particularly due to the elevated numbers 
of this bacterium present in the shrimp (except during the winter months). However, due to 
the arbitrarily selection and the low number of isolates tested, it is possible that our isolates 
may not be accurately represent the V. vulnificus population in bait shrimp. 
 All 33 V. parahaemolyticus isolates that tested positive with the tlh probe were 
verified by specific. All isolates were negative for the tdh and trh genes (Table 4- 4). In V. 
parahaemolyticus, thermostable direct hemolysin (TDH, encoded by tdh) and TDH-related 
hemolysin (TRH, encoded by trh) are considered virulence markers because they cause the 
hemolytic activity known as the Kanagawa phenomenon (32-34). The pathogenicity of the 
bacterium has been strongly related with these two hemolysins and the genes codifying for 
 
80 
 
them are commonly found in clinical strains rather than in environmental isolates (35).  
Epidemiological data (reviewed by Nishibuchi et al.) have shown that that only 1% or 2% of 
nonclinical strains carry the tdh gene (36). Therefore, the absence of these two hemolysin 
genes among the V. parahaemolyticus isolates obtained from shrimp isolates indicated that 
these isolates were not potentially pathogenic for humans.  
 In summary, I found a high prevalence of V. vulnificus and V. parahaemolyticus in 
bait shrimp. Bacterial density was high throughout the year with the exception of the winter 
months. Interestingly, the percentage of potentially virulent V. vulnificus was higher than 
those reported from shellfish and other marine samples. Conversely, all of the V. 
parahaemolyticus isolates were negative for virulence markers. Our data suggest that 
handling bait shrimp has an intrinsic risk of exposure to pathogenic V. vulnificus. GoM 
fishermen should be aware of this risk and seek immediate medical attention if wounds 
inflicted while handling bait shrimp become infected.  
  
 
81 
 
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85 
 
Table 4-1. DNA regions targeted for PCR amplification in this study; primer names, sequences, and values for the 
amplification reaction conditions that varied among primer sets; amplicon size; and literature sources of the oligonucleotide 
primers used. 
 
PCR assay* Primer name and sequence (5'?3')  PCR condition (temp., time) Primer  con. (?M) Amplicon  size (bp)  References 
VV          
 vvhA Vvh-785F (CCGCGGTACAGGTTGGCGC) 94 ?C, 30s?57 ?C, 45s?72?C, 60s 0.1 519 a 
  Vvh-1303R (CGCCACCCACTTTCGGGCC)  0.1   
 Bt-2 Bt2-F (GGACAGATATAAGGGCAAATGG) 94 ?C, 60s?60 ?C, 45s?72?C, 60s 0.2 344 b 
  Bt2-R (AGAGATGGAAGAAACAGGCG)  0.2   
 vcgC P1 (ACCTGCCGATAGCGATCT) 94 ?C, 20s?50 ?C, 20s?72?C, 20s 0.2 277 c 
  P3 (CGCTTAGGATGATCGGTG)  0.2   
 vcgE P2 (CTCAATTGACAATGATCT) 94 ?C, 20s?50 ?C, 20s?72?C, 20s 0.2 277 c 
  P3 (CGCTTAGGATGATCGGTG)  0.2   
 rrn typing UFUL (GCCTAACACATGCAAGTCGA) 94 ?C, 30s?57?C, 30s?72?C, 30s 0.2 492 d 
  URUL (CGTATTACCGCGGCTGCTGG)  0.2   
VP      
 tlh L-TL (AAAGCGGATTATGCAGAAGCACTG) 94 ?C, 60s?58?C, 60s?72?C, 120s 0.5 450 e 
  R-TL (GCTACTTTCTAGCATTTTCTCTGC)  0.5   
 tdh VPTDH-L (GTAAAGGTCTCTGACTTTTGGAC) 94 ?C, 60s?55?C, 60s?72?C, 60s 0.5 424 e 
  VPTDH-R (TGGAATAGAACCTTCATCTTCACC)  0.5   
 trh VPTRH-L (TTGGCTTCGATATTTTCAGTATCT) 94 ?C, 60s?55?C, 60s?72?C, 120s 0.5 500 e 
    VPTRH-R (CATAACAAACATATGCCCATTTCCG)        
 
86 
 
* VV refers to V. vulnificus and VP to V. parahaemolyticus. 
Table 4-1 literature references: 
a. Hill WE, Keasler S, Trucksess M, Feng P, Kaysner C, Lampel K. 1991. 
Polymerase chain reaction identification of Vibrio vulnificus in artificially contaminated 
oysters. Applied and environmental microbiology 57:707-711. 
 
b. Lee CT, Amaro C, Sanjuan E, Hor LI. 2005. Identification of DNA sequences 
specific for Vibrio vulnificus biotype 2 strains by suppression subtractive hybridization. 
Applied and environmental microbiology 71:5593. 
 
c. Rosche T, Yano Y, Oliver J. 2005. A rapid and simple PCR analysis indicates 
there are two subgroups of Vibrio vulnificus which correlate with clinical or 
environmental isolation. Microbiology and immunology 49:381-389. 
 
d. Nilsson WB, Strom MS. 2002. Detection and identification of bacterial 
pathogens of fish in kidney tissue using terminal restriction fragment length 
polymorphism (T-RFLP) analysis of 16S rRNA genes. Diseases of aquatic organisms 
48:175-185. 
 
e. Bej AK, Patterson DP, Brasher CW, Vickery MCL, Jones DD, Kaysner CA. 
1999. Detection of total and hemolysin-producing Vibrio parahaemolyticus in shellfish 
using multiplex PCR amplification of tlh, tdh and trh. Journal of microbiological methods 
36:215-225. 
 
 
87 
 
Table 4-2. Density of Vibrio vulnificus and V. parahaemolyticus in bait shrimp. 
Date V.  vulnificus density (MPN?g-1) V. parahaemolyticus density (MPN?g-1) 
(mm/dd/yy) DI, ALa OS, MSb DI, AL OS, MS 
03/26/10 <3 <3 <3 <3 
06/02/10 >1.1?105 >1.1?105 1.1?105 >1.1?105 
07/17/10 NDc >1.1?105 ND >1.1?105 
08/19/10 >1.1?105 ND >1.1?105 ND 
09/18/10 2.8?103 >1.1?105 >1.1?105 >1.1?105 
10/18/10 1.1?105 ND >1.1?105 ND 
11/12/10 1.1?105 2.4?104 4.6?104 >1.1?105 
01/29/11 <3 <3 <3 <3 
a Dauphin Island, Alabama 
bOcean Springs, Mississippi 
c Not determined.  
  
 
88 
 
Table 4-3. Result of analyzed V. vulnificus isolates for PCR analysis and for rrn and vcg 
types 
Isolate code Site Date VvhA Bt-2 16S-type Vcg C Vcg E 
DC-2 DI,  AL 06/03/10 + - B + - 
SC-2 OS, MS  06/03/10 + - A - + 
SC-1 OS, MS  06/03/10 + - B + - 
SC-4 OS, MS   07/18/10 + - A - + 
SC-7 OS, MS   07/18/10 + - A - + 
SC-8 OS, MS   07/18/10 + - A - + 
SC-9 OS, MS   07/18/10 + - A - + 
SC-11 OS, MS   07/18/10 + - A - + 
SC-21 OS, MS   07/18/10 + - A - + 
SC-19 OS, MS   07/18/10 + - A - + 
SC-13 OS, MS   07/18/10 + - B + - 
SC-3 OS, MS   07/18/10 + - B + - 
SC-5 OS, MS   07/18/10 + - B + - 
SC-10 OS, MS   07/18/10 + - B + - 
SC-15 OS, MS   07/18/10 + - B + - 
SC-16 OS, MS   07/18/10 + - B + - 
SC-18 OS, MS   07/18/10 + - B + - 
SC-23 OS, MS   07/18/10 + - B + - 
DC-12 DI,  AL 08/19/10 + - B + - 
DC-3 DI,  AL 08/19/10 + - B + - 
DC-4 DI,  AL 08/19/10 + - B + - 
DC-7 DI,  AL 08/19/10 + - B + - 
DC-9 DI,  AL 08/19/10 + - B + - 
DC-16 DI,  AL 08/19/10 + - B + - 
DC-21 DI,  AL 08/19/10 + - B + - 
DC-22 DI,  AL 08/19/10 + - B + - 
 
  
 
89 
 
Table 4-4. Result of V. parahaemolyticus isolates for PCR analysis on virulence factor 
genes 
Isolate code Site Date tlh tdh trh 
ST-1 DI, AL 06/02/10 + - - 
ST-2 DI, AL 06/02/10 + - - 
ST-1 OS, MS  06/02/10 + - - 
ST-2 OS, MS  06/02/10 + - - 
ST-4 OS, MS 07/18/10 + - - 
ST-5 OS, MS 07/18/10 + - - 
ST-6 OS, MS 07/18/10 + - - 
ST-7 OS, MS 07/18/10 + - - 
ST-8 OS, MS 07/18/10 + - - 
ST-9 OS, MS 07/18/10 + - - 
ST-10 OS, MS 07/18/10 + - - 
ST-12 OS, MS 07/18/10 + - - 
ST-13 OS, MS 07/18/10 + - - 
ST-14 OS, MS 07/18/10 + - - 
ST-15 OS, MS 07/18/10 + - - 
ST-3 DI, AL 08/19/10 + - - 
ST-7 DI, AL 08/19/10 + - - 
ST-9 DI, AL 08/19/10 + - - 
ST-10 DI, AL 08/19/10 + - - 
ST-11 DI, AL 08/19/10 + - - 
ST-13 DI, AL 08/19/10 + - - 
ST-14 DI, AL 08/19/10 + - - 
ST-15 DI, AL 08/19/10 + - - 
ST-16 DI, AL 08/19/10 + - - 
ST-17 DI, AL 08/19/10 + - - 
ST-18 DI, AL 08/19/10 + - - 
ST-19 DI, AL 08/19/10 + - - 
ST-20 DI, AL 08/19/10 + - - 
ST-24 DI, AL 09/18/10 + - - 
ST-26 DI, AL 09/18/10 + - - 
ST-16 OS, MS 09/18/10 + - - 
ST-18 OS, MS 09/18/10 + - - 
 
 
90 
 
 
 
 
 
 
CHAPTER 5. HIGH NUMBERS OF VIBRIO VULNIFICUS IN TAR BALLS 
COLLECTED FROM OILED AREAS OF THE NORTH-CENTRAL GULF OF 
MEXICO FOLLOWING THE 2010 BP DEEPWATER HORIZON OIL SPILL 
 
 
Abstract 
The Deepwater Horizon Oil Spill was the largest oil spill in USA history 
releasing approximately 4.9 million barrels of crude oil into the Gulf of Mexico. Soon 
after the spill started, tar balls and other forms of weathered oil appeared in large 
numbers on beaches in Mississippi and Alabama. In this study, we analyzed tar balls for 
total aerobic bacterial counts and also for the presence of Vibrio vulnificus, a human 
pathogen known to be abundant in the GoM Coast environment and capable of causing 
severe wound infections by contact with contaminated surfaces. Our results showed that 
total aerobic bacterial counts were significantly higher in tar balls than in sand and 
seawater collected at the same location. In addition, V. vulnificus numbers were 10? 
higher in tar balls than in sand and up to 100? higher than in seawater. Densities of V. 
vulnificus were higher than 105 colony forming units/g of tar ball in all samples analyzed. 
Our data suggest that tar balls can act as reservoirs for bacteria including human 
pathogens. 
Introduction 
The 2010 BP Deepwater Horizon Oil Spill (DHOS) released approximately 4.9 
million barrels (779 million L) of Louisiana light sweet crude oil (from the MC-252 
 
91 
 
Macondo well) into the north-central Gulf of Mexico over a period from 20 April 2010 
through 15 July 2010 (FISG, 2010). An estimated 1.1 million barrels (22%) of DHOS oil 
may still exist in the Gulf of Mexico basin as (i) surface oil (light sheen), (ii) shoreline tar 
balls, and (iii) sediment oil or submerged ?oil mats? associated with the benthos (1). 
Because microbes utilize myriad organic compounds as sources of carbon and energy, 
soon after the DHOS event began it was widely publicized that endemic gamma-
proteobacteria could significantly affect the environmental fate and ecological impacts of 
the spilled oil (2). Subsequently, and as expected, the large-scale influx of DHOS 
allochthonous carbon markedly increased the number of heterotrophic microbes in the 
water column and a high rate of hydrocarbon biodegradation was documented from 
within the DHOS oil plume (3). The few existing DHOS-related microbial studies to date 
have focused on the interactions between the oil plume and pelagic microbes (3), but no 
published study has analyzed bacteria on the large amounts of shore-cast weathered oil 
(e.g., tar balls) from within DHOS-effected coastal zone habitats (i.e., marshes and 
beaches).  
Among marine gamma-proteobacteria, Vibrio spp. are well-studied 
microorganisms because they (i) include human pathogens, (ii) are abundant in coastal 
ecosystems, (iii) are critical to the carbon cycle, and (iv) are highly effective at degrading 
varied carbon sources including some polycyclic aromatic hydrocarbons (PAH) (4). The 
human pathogen Vibrio vulnificus is of special concern for public health authorities in 
coastal areas of the southeastern United States because it is the leading cause of seafood-
borne fatalities nationwide (5). In addition to primary septicemia after ingestion of 
 
92 
 
contaminated seafood, cutaneous exposure to seawater, fish, shellfish, or fishing gear 
contaminated with this bacterium can cause severe wound infections. Signs of infection 
include inflammation and necrotizing fasciitis plus cellulitis that can lead to secondary 
septicemia. The fatality rate for patients contracting wound infections caused by V. 
vulnificus is 20?30% (6). 
Since the DHOS, many citizens have encountered tar balls by stepping on them or 
inspecting them while recreating on beaches; therefore, quantifying bacteria in tar ball is 
epidemiologically relevant since very little information is available regarding weathered 
oil as source of bacterial contamination. The objective of this study was to quantify total 
bacterial numbers in tar balls that appeared in great numbers on Mississippi and Alabama 
beaches after the DHOS. In addition, and because of the epidemiological relevance of V. 
vulnificus in the Gulf Coast, the presence of this human pathogen was also evaluated.  
Material and methods 
Sample collection. During July through October 2010, sand (200 g/collection), 
tar balls (200 g/collection), and seawater (1 L/collection) were collected aseptically from 
the intertidal (swash zone) of three beaches in Alabama and two in Mississippi (Table 5-
1; FigureV-1). Air temperature was measured in situ. Samples were placed in insulated 
coolers with ice packs immediately after collection and during transport to the Aquatic 
Microbiology Laboratory, Auburn University (internal temperature of the coolers 
remained between 15-20?C).  
Enumeration of V. vulnificus. Upon arrival to the laboratory, seawater (500 ml) 
was centrifuged at 5,000 g at 20 ?C for 30 min. Pellets were re-suspended in 10 ml (50:1) 
 
93 
 
phosphate buffered saline (PBS), and shaken vigorously to dislodge bacteria from 
suspended aggregates. Cell suspensions were 10-fold diluted in PBS to the 10-7 dilution. 
Sand (15 g of wet sand) was mixed with 15 ml of PBS and vortexed for 2 min to detach 
bacterial cells from sand grains. Large particles were allowed to be settled by gravity and 
2 ml of the supernatant was diluted into 8 ml of PBS. Ten-fold serial dilutions in PBS 
were carried out to the 10-7 dilution. Tar balls were prepared similar to sand samples, 
except that tar ball material (15 g) was broken down using a sterile inoculation loop prior 
to emulsification in PBS and a longer shaking time was employed (15 min). One hundred 
?l of each dilution were plated onto T1N1 agar (1% tryptone, 1% NaCl, 1.5% agar) in 
triplicate and incubated for 48 h at 30 ?C. Total aerobic bacterial (TAB) counts were 
calculated as colony forming units (CFU) per ml (seawater) or gram (sand and tar ball) of 
undiluted sample. To quantify V. vulnificus, colonies were transferred to a 90-mm 
Whatman no. 541 ?lter paper (Whatman, USA), followed by colony-dot-blot DNA 
hybridization using a specific V. vulnificus probe (7) as described in Food and Drug 
Administration Bacteriological Analytical Manual (FDA-BAM) (8). Positive dots were 
recorded from each filter paper as V. vulnificus counts from that dilution. 
Statistical analysis. Using Statistical Analysis System v. 9.2 (SAS Institute Inc., 
Cary, NC), both TAB and V. vulnificus counts were converted to base 10 logarithms to fit 
the model assumption of normal distribution. One-way analysis of variance (ANOVA) 
was used to determine the differences in V. vulnificus counts and Welch?s ANOVA 
(allowing for unequal variance) in TAB in all samples. If either ANOVA or Welch?s 
ANOVA was statistically significant, Tukey?s method and Scheffe's method were applied 
 
94 
 
to perform post hoc, pair-wise comparisons at ?=0.05 for the means of log V. vulnificus 
counts or the Dunnett's T3 test (allowing unequal variance) as post hoc, pair-wise 
comparisons for TAB at ?=0.05. 
Results and discussion 
Shoreline tar ball density and abundance is used to estimate the amount of spilled 
oil within an area (9) as part of the shoreline cleanup assessment team  process (10). In 
our collection sites, tar ball density was in the range of 20-40 tar balls per m2, with an 
average size of 3 cm. Tar balls are considered more of a nuisance than a public health 
concern with cutaneous allergic reactions due to hydrocarbons as primary hazard (11). 
Tar balls have not reportedly been identified as a source of infectious disease, but in this 
study we found TAB counts significantly higher in tar ball samples (5.1x106 to 8.3x106 
CFU/g) than in seawater (3.5x103 to 3.1x104 CFU/ml) and sand (1.9x105 to 4.2x105 
CFU/g) (Welch?s ANOVA p <0.0001; Dunnett's T3 p<0.05) (Table 5-2). The leves of 
TAB we described are on the upper range of those previously reported (3x104 to 3x106 
CFU/g) from tar ball samples collected in Nigeria (12), one of the few studies in where 
bacterial counts in tar balls have been measured. Interestingly, counts of the human 
pathogen V. vulnificus were significantly higher in tar ball samples than in any other 
sample analyzed (ANOVA p <0.0001, Tukey?s HSD p<0.05). Numbers of V. vulnificus 
in tar balls were >101 higher than in sand and >102 higher than in seawater. Vibrio 
vulnificus numbers in tar balls were >105 CFU/g in all cases, reaching >106 CFU/g in 4 
out of 5 samples analyzed (Table 5-2). 
To test if V. vulnificus biodegraded tar balls, we attempted to culture several 
 
95 
 
strains of this bacterium on tar ball-enriched seawater agar (12) (data not shown). No 
visible growth was observed on culture plates after 7 days of incubation. It is plausible 
that V. vulnificus, although probably not actively consuming organic compounds directly 
from the tar balls could benefit from byproducts of the microbes that do degrade 
weathered oil. 
Our values for V. vulnificus in seawater and sand were comparable to those 
previously reported in Gulf of Mexico seawater, sediment, and oysters (13). However, the 
high number of V. vulnificus recovered from tar balls is noteworthy because 
environmental samples rarely contain more than 105 CFU of V. vulnificus per gram (14). 
Therefore, the high number of V. vulnificus (>106 CFU/g) we document in tar balls from 
Alabama and Mississippi beaches has clear public health implications. Tar balls are 
sticky, especially during the warmer months, difficult to remove (11), and upon contact 
with skin abrasions may vector V. vulnificus and lead to severe wound infections. Persons 
who have immunocompromising conditions (such as liver disease) are particularly at risk 
for V. vulnificus infections (6) and should avoid contact with contaminated sources. 
  
 
96 
 
References 
 
1. Federal Interagency Solutions Group. 2010. Oil budget calculator: Deepwater 
Horizon. 
http://Www.Restorethegulf.Gov/Sites/Default/Files/Documents/Pdf/Oilbudgetcalc
_Full_Hq-Print_111110.Pdf Accessed June 12,  2011. 
2. Voosen P. 2010. Will bacterial plague follow crude oil spill along Gulf coast?, 
The New York Times. Arthur Ochs Suizberger, Js., New York. 
3. Hazen TC, Dubinsky EA, DeSantis TZ, Andersen GL, Piceno YM, Singh N, 
Jansson JK, Probst A, Borglin SE, Fortney JL, Stringfellow WT, Bill M, 
Conrad ME, Tom LM, Chavarria KL, Alusi TR, Lamendella R, Joyner DC, 
Spier C, Baelum J, Auer M, Zelma ML, Chakraborty R, Sonnenthal EL, 
D'haeseleer P, Holman N, Osman S, Lu Z, Van Nostrand J, Deng Y, Zhou J, 
Mason OU. 2010. Deep-sea oil plume enriches indigenous oil-degrading bacteria. 
Science 330:204-208. 
4. Thompson FL, Iida T, Swings J. 2004. Biodiversity of Vibrios. Microbiology 
and Molecular Biology Reviews 68:403-431. 
5. Mead PS, Slutsker L, Dietz V, McGaig LF, Bressee JS, Shapiro C, Griffin 
PM, Tauxe RV. 1999. Food-related illness and death in the United States. 
Emergent Infectious Diseases 5:607-625. 
6. Strom MS, Paranjpye RN. 2000. Epidemiology and pathogenesis of Vibrio 
vulnificus. Microbes and infection 2:177-188. 
7. Wright AC, Miceli GA, Landry WL, Christy JB, Watkins WD, Morris JG. 
1993. Rapid identification of Vibrio vulnificus on nonselective media with an 
alkaline phosphatase-labeled oligonucleotide probe. Applied and Environmental 
Microbiology 59:541-546. 
8. FDA 2002, posting date. MPN procedure for the enumeration of Vibrio vulnificus 
using gene probe for identification. [Online.] 
9. NOAA. 2000. Shoreline assessment manual (version 3.0). Hazardous Materials 
Response Division, National Oceanic and Atmospheric Administration, Seattle, 
WA. 
 
97 
 
10. Goodman R. 2003. Tar balls: the end state. Spill Science & Technology Bulletin 
8:117-121. 
11. NOAA. 2010. Understanding tar balls. NOAA's oil spill response. 
12. Itah AY, Essien JP. 2005. Growth profile and hydrocarbonoclastic potential of 
microorganisms isolated from tarballs in the Bight of Bonny, Nigeria. World 
Journal of Microbiology & Biotechnology 21:1317-1322. 
13. Motes ML, DePaola A, Cook DW, Veazey JE, Hunsucker JC, Garthright 
WE, Blodgett RJ, Chirtel SJ. 1998. Influence of water temperature and salinity 
on Vibrio vulnificus in Northern Gulf and Atlantic Coast oysters (Crassostrea 
virginica). Applied and Environmental Microbiology 64:1459-1465. 
14. DePaola A, Capers GM, Alexander D. 1994. Densities of Vibrio vulnificus in 
the intestines of fish from the U.S. Gulf Coast. Applied and Environmental 
Microbiology 60:984-988. 
 
 
  
98 
 
Table 5-1. Sample collection data. 
Collection date 
(dd/mm/yyyy) Location Geographic coordinates 
Air temperature 
(?C) 
27/07/2010 Dauphin Island, AL 30?14'54''N; 88?07'40''W 28.5 
30/07/2010 Ship Island, AL 30?12'46''N; 88?58'08''W 30.7 
06/09/2010 Gulfport, MS 30?22'07''N; 89?04'50''W 27.6 
26/09/2010 Fort Morgan, AL 30?13'30''N; 88?00'33''W 26.5 
17/10/2010 Gulf Shores, AL 30?14'55''N; 87?40'05''W 23.5 
 
  
 
99 
 
Table 5-2. Total aerobic bacterial counts (TAB) and Vibrio vulnificus counts (VVC). 
Values represent average ? standard deviation of repeat plate counts of each sample. 
Significantly different means (P<0.05) within each sampling date are noted with 
superscripts a, b, and c.  
Date Location Sample* 
Total bacterial 
count (CFU/ml or 
CFU/g) 
V. vulnificus 
count (CFU/ml or 
CFU/g) 
27-07-2010 Dauphin Island, AL SW 3.1?0.4?104a 1.0?0.0?103a 
  S 4.2?0.9?105b 3.0?1.4?104b 
  TB 5.1?1.2?106c 2.8?1.0?106c 
30-07-2010 Ship Island, MS SW 4.3?0.4?103a 5.0?2.8?102a 
  S ND? ND 
  TB 8.3?0.2?106b 3.5?0.7?105b 
06-09-2010 Gulfport, MS SW 5.4?1.9?103a 5.5?2.1?102a 
  S 3.7?0.5?105b 1.6?0.4?105b 
  TB 7.1?1.2?106c 2.8?0.4?106c 
26-09-2010 Fort Morgan, AL SW 3.5?0.3?103a 6.7?5.8?10a 
  S 3.8?0.2?105b 1.0?1.0?104b 
  TB 5.4?0.2?106c 2.7?0.6?105c 
17-10-2010 Gulf Shores, AL SW 1.9?0.7?104a 8.3?0.6?103a 
  S 1.9?0.7?104b 1.3?0.6?103b 
  TB 8.0?1.5?106c 3.3?0.5?106c 
* SW stands for sea water;  S, sand and TB, tar ball. 
? ND, not determined. 
 
100 
 
 
Figure 5-1. North-central Gulf of Mexico showing collecting sites: S1, Dauphin Island; 
S2, Ship Island; S3, Gulfport; S4, Fort Morgan; S5, Gulf Shores. Triangle indicates 
position of BP Deepwater Horizon wellsite, Mississippi Canyon Block 252 of the Gulf of 
Mexico.  
 
  
 
 101 
 
 
 
 
 
 
CHAPTER 6. EFFECT OF TEMPERATURE AND SALINITY ON THE DYNAMICS OF 
VIBRIO VULNIFICUS 16S rRNA GENOTYPES REVEALED BY ALLELE-SPECIFIC 
QUANTIFICATION PCR 
 
 
Abstract 
 Previous studies suggested the existence of two ecotypes within the species Vibrio 
vulnificus as indicated by molecular markers using the dimorphism on the virulence correlated 
gene (vcg) and the16S rDNA loci. To test whether 16S rRNA types are induced under different 
environmental conditions and thus could explain the ecotypes observed in the environment a 
series of experiments were carried out in this study. First, I investigated 16S type A and type B 
strains exhibited the same growth patterns when growing individually than when they were co-
cultured (A-type: B-type strain, 50:50) under microcosms conditions. Different temperatures and 
salinities were assayed to determine the growth patterns of 16S type A and B strains.  An allele-
specific quantitative PCR (ASqPCR) was developed to quantify the proportion shift of the two 
genotypes (A and B) in the mixed culture. Results showed that the B-type strain was better fit 
than the A-type strain at high temperature (37?C and 42?C), while the opposite was true at low 
temperatures (15?C, 10.5?C) and at a low salinity (5 ppt).  ASqPCR was also used to quantify the 
expression levels of the two 16S rRNA transcripts (A and B) in a hybrid (type AB) strain in 
response to temperature shift and different osmotic conditions. Results from this experiment 
showed no significant difference in gene expression between 16S rRNA type A and B within the 
same cells, indicating that 16S type B gene was not differentially regulated than 16S type A 
gene.   
 
 102 
 
Introduction 
Vibrio vulnificus is a Gram negative, motile, halophilic bacterium commonly found in 
estuarine and marine environments around the world (1). Although it can be isolated through a 
broad range of temperatures and salinities, this bacterium prefers warm waters (?20?C) of 
moderate salinities (5? to 25?) (2, 3). Vibrio vulnificus is an opportunistic human pathogen 
that can cause severe illnesses such as primary septicemia and wound infections (4-7). 
Consumption of raw oysters is primarily linked to septicemia cases while wound infections can 
occur by direct exposure to contaminated seafood or seawater (for a review see Shapiro et al., 
2002). Overall, the incidence of V. vulnificus infections in the USA is low, with an average of 
about 100 cases per year (8), but their severity and associated high mortality have significant 
implications for public health specialists (5, 9-11). Epidemiological studies showed that not 
everyone presented the same risk of contracting a septicemia caused by V. vulnificus (7) since 
more than 95% of septicemia patients suffered from underlying diseases, primarily liver 
disorders (4, 11) . Nevertheless, and based on the number of raw oysters consumed in the USA 
annually, the estimated number of V. vulnificus cases affecting the ?at risk population? is 
significantly higher than the observed number of cases. This paradox could be explained by an 
unequal distribution of virulence within V. vulnificus isolates (4, 12).  
To investigate if clinical isolates represented a subset of the total V. vulnificus population, 
molecular epidemiology studies were carried out to identify virulence-associated loci.  A study 
by Nilsson et al. (2003) was the first one to establish a correlation between clinical origin and 
genotyping (13). The authors used a dimorphism present in 16S rRNA gene sequence of V. 
vulnificus that have been previously described by Azanr et al. (14). Using this typing scheme, V. 
vulnificus isolates can be classified as 16S rRNA gene type A, and B. In addition, some strains 
 
 103 
 
present both alleles and are classified as 16S type AB (15). While most (76.4%) clinical isolates 
are type B, the majority of environmental isolates are type A or AB (16, 17). Similar and 
additional marker was found in a region referred to as ?virulence correlate gene? (18, 19). 
Following this classification scheme, most clinical isolates have the allele vcgC while 
environmental strains are primarily classified as vcgE (19).  Subsequent studies showed that both 
genotyping methods are generally in agreement (16, 17). Clinical strains classified as 16S type B 
present the vcgC allele while 16S type A/AB isolates are vcgE.   
Both rrn and vcg typing systems have been used in to follow the instraspecies dynamics 
of V. vulnificus in the natural environment. Lin et al. showed that the distribution of 16S type 
A/AB and type B in the water column of an estuarine in the GoM was mediated by temperature 
(20). The 16S type A strains dominated the V. vulnificus populations in the colder months, while 
type B strains peaked in September after a period of high water temperatures. A similar dynamic 
of V. vulnificus subpopulations was described in estuarine waters in North Carolina.  The 
proportion of vcgC strains increased as water warmed up and reached its highest point in August 
(21).  These findings suggested that the distribution of 16S types A and B in the environment 
was not uniform. It is plausible that environmental factors such as temperature and salinity shape 
the distribution of V. vulnificus subpopulations in the marine environment. 
The objective of this study was to assess the influence of temperature and salinity in a co-
culture of V. vulnificus 16S type A and B under controlled conditions. I hypothesized that under 
co-culture conditions the growth patterns of 16S type A and B differ, with one type being favor 
the other. To test this hypothesis, we designed a series of microcosm experiments and designed 
an allele-specific quantitative PCR (ASqPCR) technique to measure the frequency of two 
different 16S rRNA gene alleles in a given sample. In addition, we applied ASqPCR to measure 
 
 104 
 
the gene expression of alleles 16S type A and B in a V. vulnificus type AB strain to investigate if 
environmental factors directly influenced gene expression of 16S rDNA alleles.  
Material and methods 
 Bacterial strains. A total of six V. vulnificus strains were used in the study (Table 6-1). 
The original description of the 16S rRNA gene polymorphism used two clinical strains, the type 
strain ATCC 27562 and stain C7481 (14). In this study, the 16S rRNA gene was amplified, 
cloned and sequenced from strains ATCC 27562 and C7481-T. These sequences were 
considered as the reference sequences for 16S type A (ATTCC 27562) and B (C7481-T). My 
strain selection for 16S reference types was based on the study by Aznar et al. (year) who used 
the same strains to first describe the polymorphism present in the 16S rRNA gene of V. 
vulnificus. C7481-T is a translucent, avirulent morphotype of C7481, kindly provided by Dr. J. 
Oliver while ATCC 27562 is the type strain of the species.  Environmental strain VV3 (16S type 
A) and clinical strain CMCP6 (type B) were used for mixed-culture microcosm experiments. 
Strains VV3 and CMCP6 were preferred over ATCC 27562 and C7481-T since VV3 was 
originally isolated from GoM coast oysters and CMCP6 was a clinical isolate proved to be 
virulent in a mouse model (22)  . ATCC 27562 has shown to have both variants of the 16S rRNA 
gene (23) and, although of clinical origin, is more related to the V. vulnificus 16S type A group 
(24). Strain C7184-T was not chosen for the microcosm experiment due to its lack of capsule 
that might have interfered with its overall fitness. Strain CDC 9030-95 and CN 7501 were used 
in cold and heat-shock experiments; both strains were isolated from GoM coast oysters and are 
Type AB. Prior to use, all strains were confirmed as V. vulnificus using species specific PCR 
(25) and vvhA probe (26). Their genotypes were verified by using RFLP as previously described 
(13).   
 
 105 
 
 Growth curves.  Vibrio vulnificus strain VV3 (16S type A) and CMCP6 (16S type B) 
were individually cultured in 30-ml CAYEG (3% Casamino Acids [Amresco, Solon, OH], 0.3% 
yeast extract [Becton, Dickinson and Co. Spark, MD], 0.2% glucose, 0.05% KH2PO4, pH 8.4 ) 
broth (27) at 22?C overnight.  Bacterial cells were span down for 5 min at 5,000 g and washed 
once in 1? phosphate buffered saline (PBS, pH 7.4).  Then each pellet was resuspended 
individually in PBS and calibrated to an equal OD600 value of 0.5 (3.0?108 CFUs?ml-1) as the 
inoculum for strain VV3 and CMCP6. Modified M9 (mM9) medium was used for the 
inoculation as per Chase et al. (28). mM9 contained 23.2mM Na2HPO4, 11.02 mM KH2PO4, 
9.34mM NH4Cl, 1.0mM MgSO4 and was supplemented with 0.06% casamino acids, 0.006% 
yeast extract and 4.5mM glucose (28).  The base salinity of mM9 was 5 ppt. To obtain the 
different salinities required for the study, mM9 was supplemented with sodium chloride.  
Microcosms were prepared by inoculating 100 ?l of the inoculum into 100 ml of mM9 medium 
at the assayed salinity with an initial bacterial concentration of 3 ?105 CFUs?ml-1. Microcosms 
were incubated with shaking (100 rpm) at the corresponding temperature. Growth curves were 
determined at each of the 5 temperatures assayed (10.5?C?0.5?C, 15?0.5?C, 22?1?C, 37?0.5?C?C 
and 42?0.5?C?C) with a salinity of 20ppt and 3 salinity levels (5 ppt, 20 ppt and 33 ppt) with a 
set temperature of 22?C.  Each microcosm experiment was performed in duplicate. Cell growth 
was quantified by measuring OD600 of the cultures. Growth curves for each strain under each 
condition were generated by plotting mean OD600 values against culture time.  
 Construction of plasmid standards and sequencing analysis. Reference 16S type A 
and type B sequences were generated by inserting the corresponding 16S rRNA gene fragments 
into pCR?4-TOPO? vector (Invitrogen, San Diego, CA), cloned, and verified by sequencing. In 
short, near-complete 16S rRNA genes were amplified using universal primers 63V and 1387R 
 
 106 
 
(Table 6-2). The PCR was performed under a thermocycling profile: initiated with a 5-min 
denaturation at 95 ?C followed by 30 cycles of 94 ?C (45 s), 55 ?C (45 s), and 72 ?C (60 s), with 
a final extension at 72 ?C for 10 min.  PCR products were purified by 1.5% agarose gels, and 
extracted using GEANCLEAN? Turbo Kit (MP Biomedicals, Solon, OH). The purified PCR 
products were cloned using a TOPO TA cloning kit (Invitrogen, San Diego, CA). Cloned 16S 
rRNA genes were sequenced with an ABI 3730xl DNA sequencer and with vector primers T3 
and T7 or primers 537F and CD-R that targeted conserved internal 16S rRNA sequences. The 
sequencing was conducted by Lucigen Corporation (Middleton, WI).   
 Plasmids extraction and copy number determination. Sequence-verified plasmids 
containing standard 16S type A (PL-rrnA) and type B (PL-rrnB) were purified from E. coli cells 
using an Aurum Plasmid mini kit (Bio-Rad Laboratories, Richmond, CA). The concentration of 
plasmid was quantitated on a NanoDrop 1000 Spectrophotometer (NanoDrop, Wilmington, DE). 
Copy number of plasmid per microliter (CMP??l-1) was calculated according to the equation: 
                                                           , where COP is concentration of 
plasmid with unit ng??l-1, plasmid size is the number of nucleotide base pair of dsDNA, and 
Avogadro?s number is equal to 6.022?1023 molecules?mole-1. 
 Allele-specific primer design. 16 rRNA gene sequences of V. vulnificus strain 
ATCC27562, C7184 and CMCP6 were retrieved from GenBank. Two allele-specific forward 
primers and a shared reverse primer were manually designed and tested on primer Express 
software 3.0 (Applied Biosystems, Foster City, CA) and IDT SciTools Oligo Analyzer 3.0 
software (Integrated DNA Technologies, Coralville, IA). A single nucleotide polymorphism was 
placed at the 3 end (position -1 or -2) on two forward primers (Table 6-2). One additional 
mismatch was incorporated in the allele specific primer near the 3 end between position -2 and -
 
 107 
 
4. Primers were designed with melting point between 58?C and 60?C. The specificity of primer 
sequences were assessed by using NCBI online tool Primer-BLAST 
(http://www.ncbi.nlm.nih.gov/tools/primer-blast/).  All the primers were screened based on 
criteria as described by Liu et al. (29)  In short, Ct values of qPCR reaction meet the criteria that 
Ct B -A ? 8 against pure PL-rrnB ? pure PL-rrnA and CtB -A ? 1 against 50:50 PL-rrnB: PL-rrnA 
mixture (Figure 6-1). The standard rrn plasmid template was diluted to 108 copies??l-1. The 
selected primes for this study amplify a fragment of 125 bp from 16S type A strain and of 123 bp 
from 16S type B strain (Figure 6-2). 
 Quantitative Real-Time PCR assay. Quantitative PCR (qPCR) were performed in 25 ?l 
reaction volumes containing 1?l of template, 200 nM of forward and reverse primer, 
respectively, and 1? Power SYBR? Green PCR Master Mix (Applied Biosystems [AB], 
Warrington, UK) and molecular-grade water.  For each sample, reactions for both primer 
ASVvA and ASVvB were performed on ABI PRISMTM  96-well optical reaction plate (AB, 
Forster City, CA).  Each plate included non-template control (NTC) and pure A-type and B-type 
plasmid-construct positive control.  All reactions were executed in triplicate. Amplification and 
florescence were performed on a 7500 Real Time PCR System (Applied Biosystems) using the 
following profile: 10 min at 95?C for polymerase activation, 40 cycles of 15sec at 95?C and 1 
min at 60?C. A panel of annealing temperatures (60?C, 62?C and 64?C) and extension time (35s, 
45s and 60s) were compared during the optimization.  Dissociation curve analyses were 
performed using the instrument's default setting immediately after each PCR run. qPCR products 
from 12 well of each plate were randomly selected and analyzed on a 2% agarose gel to confirm 
a single ampli?ed band of a correct size. Linearity of amplification of each primer pair was 
assessed according to the determination coefficient (R2).  The R2 of the regression curve was 
 
 108 
 
estimated through the plotting of log10 initial template copies (106, 107, 108 and 109 CMP??l-1 of 
the standard plasmids) against Ct values. The PCR efficiency were calculated using the equation 
of ?E= 10(-1/S) ? 1?, where S is the slope of regression curve. The proportion of  16S type B allele 
in a mixed (c)DNA sample was obtained using equation adapted from Greer et al. (30): 
                                           , where ?Ct = (mean Ct of type B specific 
qPCR ? mean Ct of type A allele specific qPCR) and ??Ct = ?Ct ? normalization value.  The 
mean Ct values were produced from three repeats of the template on a plate, and normalization 
value was derived from ?Ct 50:50 ratio in order to correct for unequal amplification efficiency 
or plasmid concentrations (Figure 6-1).  
 Assay verification. A panel of artificial mixtures of PL-rrnA and PL-rrnB was analyzed 
with ASqPCR to assess amplification and quantification accuracy (Table 6-3).  To test the 
reproducibility and repeatability of the assays, three repeated runs of qPCR were carried out. The 
intra-run and inter-run standard deviations of Ct values were then calculated. In addition to using 
plasmid as template DNA, calibrated mixtures of strain VV3 (16S type A) and CMCP6 (16S 
type B) cells were tested. For this assay, we assumed that that both strains had same number of 
repeated 16S rRNA operon copies in their genome based on the genome sequence information 
available for the species V. vulnificus (31). To generate the mixtures, we quantitate the bacterial 
cells by plate count and OD600 value and made a series of 16S type B:16S type A (5:95, 10:90, 
90:10, 95:5) mixtures with a final concentration of 3.0?108 CFUs?ml-1. Genomic DNA was 
extracted by using the rapid boiling protocol (see below).   
 16S type A and type B strain mixed-culture microcosm. The individual inoculum for 
strain VV3 or CMCP6 in PBS (OD = 0.5) was prepared as described above. Equal volume of 
these two inoculums were mixed to produce the mixed inoculum (VV3:CMCP6=50:50). Mixed-
 
 109 
 
culture microcosms were then generated by diluting 100 ?l of the mixed inoculum into 100 ml of 
mM9 medium at the assayed salinity (see below). The initial bacterial concentration of 
microcosms was ca. 3 ?105 CFUs?ml-1. Microcosms were incubated with shaking (100 rpm) at 
the corresponding temperature. Each microcosm experiment was carried out in triplicate. 
 Environmental variables tested. The influence of temperature and salinity on the 
growth dynamics of 16S type A and B mixed cultures was assayed as follows. When temperature 
was the tested variable, salinity was fixed at 20 ppt and four temperatures were compared: 15?C, 
22?C, 37?C and 42?C. mM9 broth was equilibrate to each tested temperature prior inoculation. 
When salinity was the tested variable, three salinities at 5 ppt, 20 ppt and 33 ppt were tested at 
22?C.  When cultures reached the middle of the exponential phase (OD600 value 0.6), samples (1 
ml of culture medium) were collected in duplicate from each replicate (microcosm). For each 
temperature and salinity assayed, a total of 6 samples (3 microcosm replicates ? 2 samples from 
each microcosm) were analyzed. 
 Cell viability at low temperature.  To determine if there was a difference in cell 
viability between 16S type A and type B strains under low temperature, the following study was 
carried out. A mixed inoculum of strain VV3 and CMCP6 50:50 was made as described above. 
Because cells were not expected to grow at the assayed low temperature, instead of inoculating 
100 ?l of the inoculum into 100 ml modified M9 medium as before, I diluted 10 ml of the mixed 
inoculum into 90 ml of mM9 medium (salinity 20 ppt) to obtain sufficient cells for DNA 
extraction. Thus, the initial cell density was 3?107 CFUs?ml-1. Microcosms were incubated at 
10.5?C with shaking (100 rpm). Samples for DNA extraction were taken at 0, 24, and 48 h after 
inoculation as processed as before. In addition, single-strain cell suspensions (3?107 CFUs?ml-1) 
of each individual strain were inoculated in parallel. The viability of each individual strain as 
 
 110 
 
well as the mixed culture was at 10.5 ?C was assessed by using L7012 LIVE/DEAD? BacLight 
Viability kit (Invitrogen) at 24 and 48 h post-inoculation.                      
 DNA extraction. Bacterial cells were immediately pelleted from 1 ml samples by 
centrifugation at 10,000 g for 1min.  Pellets were washed in 1?PBS, resuspended into 350 ?l of 
1? TE buffer (10mM Tris, 1mM EDTA, pH8.0) and subsequently transferred into a sterile 2 ml 
screw-cap tube containing 200 mg 150-212 ?m acid-wash glass beads (Sigma-Aldrich Co. St. 
Louis, MD). Cell disruption was performed by vortexing the tube horizontally at maximum scale 
for 5 min on Vortex-Genie? 2 mixer (Mo Bio Laboratory Inc.).  The lysates in tubes were heated 
at 100?C for 15 min and cooled on ice followed by centrifugation of 16,000 g for 5 min. The 
supernatant was carefully transferred to a new tube and stored at -80?C until used as DNA 
templates.               
 Microcosms with 16S type AB strains. Two 16S type AB strains, CN 7501and CDC 
9030-95, were used to determine if the expression of 16S rRNA gene alleles was differentially 
influenced by temperature or salinity. Cold and heat shock (C/HS) as well as changes in salinity 
were assayed as follows. For each 16S type AB strain, the inoculum was prepared as described 
above.  A 100 ?l of inoculum was inoculated into 100 ml sterile mM9 medium (for C/HS 
treatments salinity was maintained at 20 ppt while for the salinity assays temperature was 
maintained at 22?C).  In C/HS experiments, the culture was initially grown at 22?C to an OD600 
of ca. 0.5 before being transferred to 10.5?C (CS) or 42?C (HS) for 30 min. Samples (1 ml of the 
culture) for RNA extraction were taken from each microcosm in duplicate immediately after 
C/HS. As control, one set of microcosm was incubated at 22?C throughout the experiment.  For 
the salinity test, cultures were sampled when they reached an OD600 of 0.6 at 22?C under 
different salinities (5 ppt, 20 ppt and 33 ppt).  Samples were centrifuged at 10,000 g for 1min and 
 
 111 
 
immediately preserved in RNAlater solution (Ambion, Austin, TX) at 4?C according to the 
manufacturer?s instructions. Samples were taken in duplicate from each microcosm, and RNA 
extractions of two repeats were pooled before reverse transcription. RNA was isolated and 
reversely transcribed to cDNA within 24 hours (see below).  
 Isolation of RNA and preparation of cDNA for 16S type AB strains. Total bacterial 
RNA was isolated using RNeasy? Mini Kit (Qiagen, Inc., Valencia, CA). In brief, 16S type AB 
cells were pelleted from RNAlater suspensions by centrifugation at 10,000 g for 30 sec, and 
resuspended in 700 ?l of  lysis buffer RLT of the RNeasy Kit (added with 1% of ?-
mercaptoethanol) after the removal of the supernatant.   The resuspension was transferred to a 2 
ml screw cap tube filled with 100 mg 150-212 ?m acid-washed glass beads. Cells 
homogenization was performed in a mini-bead beater (Biospec Products, Bartlesville, OK) for 30 
sec twice with a 15 sec on-ice off in between and immediately cooled down on ice. Additional 
steps followed manufacturer?s instructions (32).  DNA was removed from RNA samples by 
using a TURBO DNase-free kit (Ambion, Cambridgeshire, UK) following the protocol for 
rigorous DNase treatment. Treated RNA was confirmed to contain an insignificant amount of 
gDNA as indicated by a Ct value >32 when RNA was directly used as template in ASqPCR. 
RNA concentration was determined with a NanoDrop? ND-1000 spectrophotometer.  Each of 
RNA sample was adjusted to a concentration of 100 ng??l-1 with RNase-free sterile water. One 
?g of RNA was reversely transcribed to cDNA by using the High Capacity cDNA Reverse 
Transcription Kits (Applied Biosystems). cDNA samples were stored at -80?C until further 
analysis.  
 Statistical analysis. The one-way or two-way ANOVA with Tukey?s post hoc analysis 
was used to determine the significance of the frequency of 16S type B strain in mixed-culture 
 
 112 
 
microcosms or B-type 16S rRNA in type AB strain cells under different conditions. The ratio of 
A-type and B-type 16S rDNA allele copies in 16S type AB strain was tested by pairwise t-test. 
The statistical analysis was performed on the SAS Software 9.2 version (SAS Institute, Cary, 
N.C.). 
Results 
 Growth curves of strain VV3 and CMCP6. Growth curves are shown in Fig 3 
(temperature gradient) and Fig 4 (salinity gradient). Both strains displayed similar growth 
patterns at 15 ?C, 22 ?C and 37 ?C while no growth was observed at 10.5 ?C. Interestingly, only 
16S type B CMCP6 strain grew at 42?C while no growth was observed in the 16S type strain 
VV3. Salinity had little effect on the growth curves and both strain produced similar growing 
patterns under the three salinities tested. However, 16S type B strain CMCP6 exhibited a shorter 
lag phase than 16S type A VV3 strain at 20 ppt salinity (Figure 6-4).  
 ASqPCR assay validation.  PL-rrnA construct was amplified with specific primer 
ASVvA and PL-rrnB construct with primer ASVvB.  Dissociation curves generated only a single 
peak for each type amplicon (Tm with a slight difference: ca. 82?C for amplicon A and ca. 83?C 
for amplicon B), and no peaks that indicates the presence of primer-dimers. The ASqPCR 
parameters for A type (ASVvA) and B type (ASVvB) are shown in Table 6-3.  The relatively 
low PCR efficiencies were expected due to the intentional mismatches introduced into the 
primers to balance the specificity. The short length region containing the A/B polymorphism in 
the 16S gene limited option for specific primer design resulting in the two ASqPCR reactions 
having slight different amplification efficiencies. However, both reactions have a good linearity 
indicated by the coefficient of determination R2 >0.99 for the regression curve (Ct value against 
log10 [plasmid copy number]). The average intra-run and inter-run standard deviations of Ct value for A 
 
 113 
 
type and B type reaction were lower than 0.18, indicating a good reproducibility between PCR 
runs. Testing the artificial mixtures of PL-rrnB from 1% to 95% at 108 copies??l-1 displayed a 
good correlation between the expected percentage and the calculated percentage (Table 6-4). To 
test the ASqPCR efficacy when cell mixtures were used we compared the calculated value of 
16S type B percentage with the expected value determined by plate count. The percentage of 16S 
type B cells calculated in the four tested ratios was as follows: 1) for a A:B ratio of 5:95 the 
calculated 16S type percent was 3.5%; 2) for 10:90 was 7.6%; 3) for 90:10 was 90.2%; and 4) 
for 95:5 was 95.4%.  In general, the deviation between the expected value and calculated value 
was small, although it was slightly higher when the percent of 16S B type cells was low. 
 Dynamics of 16S type A and type B strains under co-culture conditions. Figure 6-5 
summarizes the results of the co-culture experiments. At low growth temperature (15?C), my 
results showed that 16S type A VV3 strain constituted 97.8% of the population in the mixed 
culture when the culture reached the late exponential phase.  Conversely, 16S type B CMCP6 
strain outcompeted VV3 at 37?C with 70.4% of cells estimated to be 16S type B. At 42?C, 
practically the whole population was composed of 16S type B cells (99.9%) (Figure 6-5A). 
When 16S type A and B co-cultures were incubated at room temperature the percentage of 16S 
type B cells was close to 50% (49.6%) indicating that this temperature does not play a selective 
role for either 16S type.  Regarding salinity, the 16S type A outcompeted 16S type B cells under 
low-salinity conditions (5ppt) with 16S type B cells remaining below 0.4% at the late 
exponential phase.  Similar percentages (~50%) of both strains were detected under salinities of 
20 ppt and 33 ppt (Figure 6-5B), suggesting these salinities do not favor any 16S type.  
 Survivability at 10.5?C. The effect of cold temperature (lower than the minimum 
temperature require for growth) on both 16S types was assessed during a 48-hour incubation 
 
 114 
 
period at 10.5?C. The results of dual fluoresce staining on each single strain showed that the 
density of viable cells decreased between 24 h and 48 h for both strains (Figure 6-6). However, 
based on qualitative microscopy data, more 16S type A cells remained viable at 24 h and 48 h 
than 16S type B. ASqPCR analysis on the co-cultured microcosm confirmed the results observed 
by microscopy since the percentage of 16S type B strain (both live and dead cells with intact 
genomic DNA) declined from the initial 62.7% to 23.4% at 24 h and to a final 9.9 % after 48 
hours at 10.5?C (Figure 6-7).  Here we have an assumption that dead cells of each genotype 
strain were lysed at same rate. 
 Determination of 16S rRNA alleles copy number in 16S type AB strains. The 
frequency of the 16S type B allele in the genomic DNA extracted from the 16S type AB strain 
was determined by ASqPCR. Results showed that in strain CN 7501 the copy number of 16S 
type B allele was 10.87% ? 0.66% (mean ? SD, measured in 3 independent DNA extractions) in 
strain CN 7501 while in strain CDC 9030-95 was 12.17% ? 1.55%. Statistical analysis using 
pairwise t-test comparison indicated that the ratio of A: B alleles are significantly equal to 9:1 
(p=0.58 for strain CN 7501 and p=0.15 for CDC 9030-95, n=3, ?=0.01). These results were 
taken into account when calculating the gene expression levels in the cold/heat shock 
experiments. 
 Expression of 16S alleles in the 16S type AB strains in response to cold/heat shock 
and salinity. Changes in temperature did not significantly (?=0.01) influence the expression 
level of 16S type B gene in either strain tested.  Levels of 16S type B gene transcript remained 
the same in cold and heat-shocked treated cultures as they were in controls and represented 
approximately 15% of the total detected 16S rRNA gene transcripts. Similarly, no significant 
 
 115 
 
differences were found in 16S allele expression when strains were incubated at different 
salinities (Figure 6-8).   
Discussion 
 The goal of this study was to determine if there was a difference in the growth pattern of 
16S type A and type B of V. vulnificus could explain why 16S type B is more abundant at the 
end of the summer period. The ASqPCR method I designed for this study allowed me to 
determine the proportion of cells belonging to each 16S genotype in a mixed culture. My data 
showed that when both types were cultured individually, their growth curves were similar at 
15?C, 22?C, and 35?C while only 16S type B strain grew at 42?C. However, when strains were 
co-cultured, the increase of 16S type B cells at 35?C was significant, indicating this type 
outcompeted 16S type A at this temperature.  As expected, since 16S type A did not grow at 
42?C, when both strains were co-cultured at this temperature, only 16S type B was detected. 
Conversely, 16S type A outperformed 16S type B cells at 15?C although both strains displayed 
similar individual growth patterns at 15?C. 
 The structural variation of V. vulnificus population in the environment is influenced by 
water temperature. The seasonal dynamic of V. vulnificus 16S type A and type B subpopulations 
in the water column has been observed in the Galveston Bay, TX and in Alligator Bay, NC (20, 
33). In both studies, the percentage of 16S type B (vcgC) strains increased with water 
temperature, and peaked at the end of summer (when seawater temperatures was above 30?C).  
These observations obtained from field surveys are supported by the results of my in vitro study.  
In vitro, the proportion of 16S type B cells increased from ca. 50% to 77% at 37?C and up to > 
99 % at 42?C when the mixed culture reached to an OD of 0.6. Thus, it is likely that warmer 
water temperature acts as a selecting force that increases the proportion of 16S type B strains 
 
 116 
 
during the summer. Moreover, this adaptation to warmer temperatures displayed by 16S type B 
might explain why 16S type B are responsible for the majority of V. vulnificus infections in 
humans (13, 19). It is possible that the human body temperature (37?C) acts as a filter when V. 
vulnificus enters the human host allowing 16S type B cells to outgrow 16S type A cells.  Thus, it 
is plausible that the main difference between 16S type A and B is not based on virulence factors 
but in the adaptation to warmer temperatures displayed by 16S type B. This notion is supported 
by the fact that when tested in mice, a correlation between 16S type B and virulence was not 
found and strains of both types (A and B) were found to cause disease in this mammal model 
(22). 
 My results also suggest that 16S type A strains are better adapted to low temperatures 
than 16S type B. The 16S type A strain increased its percentage in relationship to type B under 
low temperature (15?C) conditions. In addition, more 16S type A type cells remained viable after 
48-hour incubation at below-growth temperature (10.5?C). These findings agree with previous 
results from field studies.  In Lin and Schwarz?s study, both genotypes of V. vulnificus were 
undetectable during the winter months but 16S type A strains were recovered earlier in spring 
than 16S type B.  Similarly, data from the Warner and Oliver?s study demonstrated that a higher 
proportion of 16S type A was present in the water column when the bacterium was first 
recovered in spring before the summer bloom (33).  
Additionally, 16S type A V. vulnificus strains seemed better adapted to low salinities than 
16S type B strains.  My in vitro results showed a competitive advantage of the 16S type A strain 
over the 16S type B strain in the microcosm with a salinity of 5 ppt.  Herein, my study supports 
the idea (34) that two ecotypes are present within the species V. vulnificus and that they are better 
adapted to different environmental conditions that play a significant role in the life cycle of an 
 
 117 
 
estuarine bacteria such are temperature and salinity. It needs to be noted that although the 16S 
marker used targets a very small fragment of the V. vulnificus genome, a perfect correlation 
exists between the two genotypes found within the species using AFLP analysis (whole genome 
typing method) and 16S type A/AB and B (24, 35). Therefore, is expected that many other loci 
besides the 16S rRNA gene are implied in the observed adaptations to temperature and salinity.   
 The presence of both 16S haplotypes in 16S type AB strains posits an interesting 
evolutionary question as duplicate genes tend to share the same sequence through 
homogenization processes analogous to eukaryotic gene conversion. Members of the genus 
Vibrio to carry more than one rrn operon and intragenomic heterogeneity is not uncommon. It 
has been postulated that bacteria with more rrn operons can adapt quicker to changes in the 
environment than those with fewer copies. Vibrio vulnificus contains 9 rrn copies and a study by 
Arias et al. (36) revealed a high degree of interoperonic heterogeneity. Are these different 
operons the result of a recent lateral transfer event and thus have not had time to converge 
through homogenization? Or, do they offer an evolutionary advantage to the cell and thus they 
have been maintained over time?  Recently, is has been demonstrated that different 16S 
haplotypes can be differentially expressed within the same cell depending on environmental 
factors (37). To determine if this was the case in V. vulnificus, I measured the expression level of 
each 16S rDNA allele within the same cell under different environmental conditions. If the 16S 
rRNA variants in the V. vulnificus hybrid strain were functionally specialized, I expected to 
observe changes in the frequency of 16S rRNA type A/B expressed in the cells. My results 
showed that both 16S rDNA variants were expressed in the 16S type AB strain cells. However, 
the proportion of A/B transcripts indicated no selective expression of either variant as levels of 
both types remained statistically identical regardless of the treatment applied.  
 
 118 
 
In summary, under microcosm conditions, I showed that 16S type A and type B of V. 
vulnificus significantly different in their growth response to different temperature and salinity 
regimes. Interestingly, some of those differences were only observed when both types were co-
cultured indicating that V. vulnificus subpopulation dynamics in nature are influenced by 
temperature and salinity. In hybrid 16S type AB strains, the two 16S alleles are not regulated 
differently from each other when exposed to temperature changes and different osmolality, 
which suggest that the better fitness observed in the co-culture experiments was not due to the 
expression of a specific 16S alleles but likely to other loci. This study provided additional 
evidence for the delineation of two V. vulnificus ecotypes. 
 
  
 
 119 
 
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2011. Genotype is correlated with but does not predict virulence of Vibrio vulnificus 
 
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immunity 79:1194-1207. 
23. Arias CR, Olivares-Fuster O, Goris J. 2010. High intragenomic heterogeneity of 16S 
rRNA genes in a subset of Vibrio vulnificus strains from the western Mediterranean coast. 
International microbiology 13:179-188. 
24. Tao Z, Larsen AM, Bullard SA, Wright AC, Arias CR. 2012. Prevalence and 
Population Structure of Vibrio vulnificus on Fishes from the Northern Gulf of Mexico. 
Applied and environmental microbiology 78:7611-7618. 
25. Hill WE, Keasler S, Trucksess M, Feng P, Kaysner C, Lampel K. 1991. Polymerase 
chain reaction identification of Vibrio vulnificus in artificially contaminated oysters. 
Applied and environmental microbiology 57:707-711. 
26. Wright AC MG, Landry WL, Christy JB, Watkins WD, Morris JG Jr. 1993. Rapid 
identification of Vibrio vulnificus on nonselective media with an alkaline phosphatase-
labeled oligonucleotide probe. Applied Environmental Microbiology 59:541-546. 
27. Tamplin ML, Colwell RR. 1986. Effects of microcosm salinity and organic substrate 
concentration on production of Vibrio cholerae enterotoxin. Applied and environmental 
microbiology 52:297-301. 
28. Chase E, Harwood VJ. 2011. Comparison of the Effects of Environmental Parameters 
on Growth Rates of Vibrio vulnificus Biotypes I, II, and III by Culture and Quantitative 
PCR Analysis. Applied and environmental microbiology 77:4200-4207. 
29. Liu CM, Driebe EM, Schupp J, Kelley E, Nguyen JT, McSharry JJ, Weng Q, 
Engelthaler DM, Keim PS. 2010. Rapid quantification of single-nucleotide mutations in 
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virological methods 163:109-115. 
30. Germer S, Holland MJ, Higuchi R. 2000. High-throughput SNP allele-frequency 
determination in pooled DNA samples by kinetic PCR. Genome Research 10:258-266. 
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Progulske-Fox A, Hillman JD, Handfield M, Rhee JH. 2003. Characterization and 
Pathogenic Significance of Vibrio vulnificus Antigens Preferentially Expressed in 
Septicemic Patients. Infection and immunity 71:5461-5471. 
32. Qiagen. 2006. Purification of total RNA from bacteria using the RNeasy? Mini Kit. 
http://www.qiagen.com/literature/render.aspx?id=579 (Accessed on November 10, 2012). 
 
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33. Warner E, Oliver J. 2008. Population structures of two genotypes of Vibrio vulnificus in 
oysters (Crassostrea virginica) and seawater. Applied and environmental microbiology 
74:80. 
34. Rosche TM, Binder EA, Oliver JD. 2010. Vibrio vulnificus genome suggests two 
distinct ecotypes. Environmental Microbiology Reports 2:128-132. 
35. Wood RR, Arias CR. 2012. Distribution and survival of Vibrio vulnificus genotypes in 
postharvest Gulf Coast (USA) oysters under refrigeration. Journal of Applied 
Microbiology 113:172-180. 
36. Tao Z, Bullard S, Arias C. 2011. High numbers of Vibrio vulnificus in tar balls 
collected from oiled areas of the North-Central Gulf of Mexico following the 2010 BP 
deepwater horizon oil spill. EcoHealth 8:507-511. 
37. L?pez-L?pez A, Benlloch S, Bonf? M, Rodr?guez-Valera F, Mira A. 2007. 
Intragenomic 16S rDNA Divergence in Haloarcula marismortui is an adaptation to 
different temperatures. Journal of molecular evolution 65:687-696. 
 
 
  
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Table 6-1. Vibrio vulnificus strains  
Strain ID Source 16S type 
CN 7501 Fish, Alabama, USA AB 
CDC 90 1506 Clinical, Florida, USA AB 
CMCP6 Clinical, South Korean A 
VV3 Oyster, Alabama, USA B 
ATCC 27562 Human blood, Florida, USA A 
C7184 Human blood, USA B 
 
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Table 6-2. Primers for sequencing and ASqPCR. 
Assay Sequence (5'? 3') Reference 
Sequencing primers   
63V CAGGCCTAACACA TGCA AGTC a 
1387R GGGCGGWGTGTACAAGGC a 
533 F GTGCCAGCAGCCGCGGTAA b 
CD-R CTTGTGCGGGCCCCCGTCAATTC c 
ASqPCR primers* 
  Fw.(ASVvA) CGATGGCTAATACCGCATCATA This study 
Fw.(ASVvB) ATGGCTAATACCGCATCATGC This study 
Rev.(Common) AGGGATCGTCGCCTTGGT This study 
*For allele specific qPCR primes, the incorporated mismatch is shown as underlined, and 
the target single-nucleotide mutation locus is shown as bolded.  
Table 6-2 literature references: 
a. Marchesi JR, Sato T, Weightman AJ, Martin TA, Fry JC, Hiom SJ, Wade 
 WG. 1998.  Design and evaluation of useful bacterium-specific PCR primers 
 that amplify genes coding for bacterial 16S rRNA. Applied and environmental 
 microbiology 64:795-799. 
 
b. Weisburg WG, Barns SM, Pelletier DA, Lane DJ. 1991. 16S ribosomal DNA 
 amplification for phylogenetic study. Journal of bacteriology 173:697-703. 
 
c. Rudi K, Skulberg OM, Jakobsen KS. 1998. Evolution of Cyanobacteria by 
 Exchange of Genetic Material among Phyletically Related Strains. Journal of 
 bacteriology 180:3453-3461. 
 
 
 
 
 
 
 
 
 
  
 
 125 
 
Table 6-3. Slopes, Intercepts, R2, efficiencies, intra/inter -run standard deviations of two 
ASqPCR amplifications. 
qPCR reaction Slope Intercept R2 Efficiency  
Ct value intra-
run SD. average 
(range) 
Ct value inter-
run SD. average 
(range) 
A type (ASVvA) 3.814 42.217 >0.99 83% 0.17 (0.03-0.49) 0.14 (0.03-0.25) 
B type (ASVvB) 3.574 40.429 >0.99 90% 0.18 (0.02-0.35) 0.14 (0.03-0.25) 
  
 
  
 
 126 
 
Table 6-4. Relative 16S rRNA allele frequencies using standards of mixed plasmid 
constructs 
Ratio (B:A) ?Ct B-A ?CtB-A (Nor.) Expected % B Calculated % B 
100:0 11.57 11.33 0 0.04 
1:99 7.40 7.16 1 0.69 
10:90 9.48 9.24 10 9.50 
90:10 -2.78 -3.02 90 89.00 
95:5 -3.72 -3.96 95 94.00 
100:0 -9.10 -9.34 100 99.85 
50:50 0.24a 0.00 - - 
 
These assays were evaluated at 108 total copies per reaction.  a The ?Ct B-A for ratio 50:50 
should be equal to zero, where the deviation reflected the differences in amplification 
efficiencies and plasmids copy number in the reaction.  ?CtB-A (Nor.) are the results of 
?Ct B-A subtracted with the normalization value for the assay dynamic range (see the 
text). These normalized values were used to calculate the percentage of 16S genotype B 
plasmid in the mixture. 
  
 
 127 
 
 
 
 128 
 
Figure 6-1. ASqPCR amplification plots of 108 copies of (A) 50:50 standard rrn plasmid 
construct of A-type : B-type, (B) pure A-type plasmid, and (C) pure B-type plasmid.  The 
?CtB-A with A : B (50:50) was the normalization value. The graph shows that this value 
calculated at 108 copies was 0.24. In this study, the average normalization value across 
the entire dynamic range from 106 to 109 copies was generated to use for the ASqPCR 
assay. The average ?Ct was also equals to 0.24. 
  
 
 129 
 
 
 
Figure 6-2. Partial 16S rDNA sequence alignment of V. vulnificus strain C7184 
(GenBank accession no.: X76334), CMCP6 (NR 074889), ATCC 27562 (X76333) and E. 
coli (J01695). Shading indicates the ASqPCR primer positions: P1, P2 and P3 refer to 
primer ASVvA, ASVvB and common reverse primer, respectively. Numbers on top ruler 
denote position in the Escherichia coli reference sequence. 
  
 
 130 
 
 
Figure 6-3. Growth curves of V. vulnificus across 5 temperature gradients (with a salinity 
of 20ppt): (A) strain VV3; (B) strain CMCP6. Data are the average values of two 
replicates. 
  
 
 131 
 
 
Figure 6-4. Growth curves of V. vulnificus across 3 salinity gradient (at 22?C): (A) strain 
CMCP6; (B) strain VV3. Data are the average values of two replicates. 
 
  
 
 132 
 
 
Figure 6-5. Growth dynamic of the co-culture of two genotypes in the mixed-culture 
microcosm at different conditions when the OD reaches to 0.6: (A) across a temperature 
gradient (medium salinity 20ppt); (B) across different salinities (at 22?C). 
  
 
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Figure 6-6. Photographs of V. vulnificus strain CMCP6 and VV3 epiflourescence 
microscopic observations after 24-h and 48-h incaution at 10.5?C. Bacteria were stained 
with L7012 LIVE/DEAD? BacLight Viability kit before the microscopic observation at 
320? magnification.  Cells fluorescing green are considered alive while red fluorescing 
cells are considered dead. 
  
 
 134 
 
 
Figure 6-7. The genotype dynamic incubating at 10.5?C at 0h, 24h and 48h (below 
minimum growth temperature; OD value declined over time, see text). 
  
 
 135 
 
 
Figure 6-8. Expression level of B-type 16S rRNA (rrn) gene relative to the expressed 16S 
rRNA genes (A and B) through cold/heat shock (medium salinity 20ppt); (C) across a 
salinity gradient (at 22?C). Means ? SD are shown. The frequencies of B-type 16S rRNA 
do not vary significantly with cold/heat shock treatments or under different salinities. The 
significance level of ?=0.01was used.   
 
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CHAPTER 7. SUMMARY AND CONCLUSION 
 
 
 In this dissertation, I investigated the distribution and prevalence of two human-
pathogenic Vibrio species (V. vulnificus and V. parahaemolyticus) in non-shellfish 
samples including fish, bait shrimp, water, sand and crude oil material released by the 
Deepwater Horizon oil spill along the Northern GoM coast.  
 Results from the fish investigation showed that 37% of all sampled fish (n=242) 
were positive for V. vulnificus presence on skin and mucus. A total of 244 V. vulnificus 
isolates were recovered from fish and further analyzed to determine the population 
structure of this species in fish. Ascription to 16S rRNA gene type indicated that 157 
isolates were type A (62%), 72 (29%) were type B (significantly correlated with clinical 
cases) and 22 (9%) were type AB. Amplified fragment length polymorphism (AFLP) was 
used to resolve the genetic diversity within the species. One hundred and twenty one 
unique AFLP profiles were found among all analyzed isolates resulting in a calculated 
Simpson?s index of diversity of 0.991. AFLP profiles were not grouped based on 
collection date, fish species, temperature or salinity, but isolates were clustered into 2 
main groups that correlated precisely with 16S type.  
 A second investigation focused on the prevalence of two human pathogens, Vibrio 
parahaemolyticus and V. vulnificus, in bait shrimp obtained from commercial bait shops 
along the coast of Alabama and Mississippi. Both Vibrio species were below the 
 
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detection limit in November and March, and reached the highest level during the summer 
months (>1.1?105 Most Probable Number (MPN) per gram shrimp tissue) in both Vibrio 
species. Randomly selected isolates of both species were typed using virulence-related 
typing schemes. Out of 26 V. vulnificus isolates analyzed, 69% were 16S type B 
indicating the virulence potential of the isolates. By contrast, none of the 33 V. 
parahaemolyticus isolates was positive for tdh and trh hemolysin (virulence factors) 
genes.  
 Soon after Deepwater Horizon Oil Spill in the Gulf in 2010, I investigated if the 
large number of weathered oil (tar ball) present on beaches in Alabama and Mississippi 
acted as a sink for bacteria. My results showed that total aerobic bacterial counts were 
significantly higher in tar balls than in sand and seawater collected at the same location. 
Densities of V. vulnificus were higher than 105 colony forming units (CFU)/g of tar ball 
in all samples analyzed. In addition, V. vulnificus numbers were 10? higher in tar balls 
than in sand and up to 100? higher than in seawater.  
 Lastly, I used microcosm experiments in combination with allelic quantitative 
PCR technique, to demonstrate that V. vulnificus16S type B strain is better fit than type A 
strains under high environmental temperatures (37?C and 42?C). Conversely, type A 
grows or survives better than B-type at low temperature (15?C, 10.5?C) and low salinity 
(5 ppt). These findings further support the notion that posits two different ecotypes (16S 
type A and B) are present within the V. vulnificus species.  
 In summary, research data showed that the human pathogen V. vulnificus is 
commonly found in non-shellfish samples of the Northern GoM coast. Moreover, I 
 
 138 
 
discovered a higher percentage of strains of great virulence potential in fish and shrimp 
than those previously reported in oysters. I proved that 16S type B strains outcompete 
type A strains at warmer temperatures explaining why more cases of vibriosis due to this 
pathogen occur at the end of summer. Finally, the effects of the Deepwater Horizon oil 
spill significantly increased the presence of V. vulnificus in beach samples. Overall, my 
research shows that recreational activities conducted in the Northern GoM coast have an 
intrinsic risk of exposure to V. vulnificus. 
 
  
 
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APPENDIX 1. DIVERSITY OF CULTURABLE BACTERIA FROM BLOOD 
VASCUAR SYSTEM OF LESSER ELECTRIC RAYS (NARCINE BANCROFTII) 
CAPTURED IN THE NORTHERN GULF OF MEXICO OFF ALABAMA 
 
 
Abstract 
The prevalence and taxonomic diversity of culturable bacteria recovered from the 
blood vascular system of healthy, grossly normal and showing no clinical sign of disease, 
lesser electric rays (Narcine bancroftii) captured from open beach habitat in the north-
central GoM are reported herein. The blood of 9 of 10 rays was positive for bacteria, and 
bacterial isolates (n=83) were identified by partial 16S rRNA gene sequences. Vibrio spp. 
comprised 53% of all isolates and was recovered from all blood culture-positive rays. 
Among them, V. harveyi (n=14) and V. campbellii (n=11) were most common, followed 
by a group of unidentified Vibrio sp. (n=10) related to V. nigripulchritudo. Isolates 
representing species of Pseudoaltermonas (n=13), Shewanella (n=5), Anphritea (n=3), 
Nautella (n=3), and Arenibacter (n=1) were also recovered from ray blood. Higher 
bacterial diversity was observed in blood cultured on marine agar relative to blood agar 
but Gram positive bacteria were isolated from the latter only. Partial 16S rRNA gene 
sequences of the ray bacterial isolates were compared phylogenetically to those from 
related type strains. Most isolates were identified to the level of species but some 
clustered independently from reference strains, likely representing new species of Vibrio, 
Amphritea, Shewanella, and Tenacibaculum. The present study is the first record of any 
 
 140 
 
bacterium from this ray species and reveals a taxonomically and phylogenetically diverse 
microbiota associated with its blood. Moreover, these data document that the presence of 
bacteria in elasmobranch blood is not coincident with clinical signs of disease; thereby 
rejecting the paradigm of septicemia indicating a disease condition in aquatic vertebrates. 
Introduction 
Little published information exists on the biodiversity, prevalence, and 
physiological effects of bacteria that infect the blood and other tissues of cartilaginous 
fishes (Chondrichthyes: sharks, skates, rays, and chimaeras) (1, 2) (Table 1). Based on a 
perusal of the published literature, Vibrio spp. comprise a large component of the normal 
microbiota in shark blood (3, 4). Based on this foundational taxonomic work with blood-
borne bacteria in sharks, seemingly, the classical assumption that bacterial presence in 
blood indicates disease is no longer robust: (i) healthy captive elasmobranchs have urea-
hydrolyzing bacteria in their blood vascular system (1) that may aid in osmoregulation, 
(ii) bacteria have been isolated from liver, spleen, kidney, and pancreas of healthy sharks 
and rays (1, 3) (Table 1), and (iii) Mylniczenko et al. (1) reported that 27% (50% when 
only rays were considered) of healthy captive and free-ranging elasmobranchs (n=80) 
yielded positive blood cultures. Yet, other bacteria, including Vibrio spp., recovered from 
sharks are indeed considered opportunistic pathogens (4). Hence, whether the taxonomic 
spectrum of these elasmobranch-associated bacteria comprises opportunistic/obligate 
pathogens, benign commensals, or bona fide tissue-dwelling symbionts that serve a 
critical role in elasmobranch physiology is indeterminate. Nevertheless, documenting 
microbial taxonomic diversity in other elasmobranch lineages is a good first step towards 
 
 141 
 
testing such hypotheses. No such detailed study has been published based on materials 
sampled from non-shark elasmobranchs, i.e., skates, rays, or chimaeras. 
The lesser electric ray, Narcine bancroftii (Griffith and Smith, 1834), 
(Torpediniformes: Narcinidae; syn. Narcine brasiliensis) ranges in shallow waters of 
tropical and sub-tropical continental shelves to 37 m depth, including the GoM, the 
Caribbean Sea, and the islands of the West Indies (5). Three other species of Narcinidae 
have geographic ranges that overlap with N. bancroftii, but N. bancroftii is the only 
narcinid that reportedly ranges in the north-central GoM (= the focus area for the present 
study)(6). This ray is a slow swimming fish that can be seasonally aggregated on 
sandbars and surf zones along open beaches and barrier islands. It can be regionally 
abundant in summer months, during which time pregnant females birth viviparous 
offspring, but then moves to offshore deep waters in winter (7). During Fall (Aug-Oct) in 
the northern GoM, lesser electric rays can be observed commonly by snorkeling in waters 
of 0.2?3.0 m; with the spiracles of the nearly completely buried rays appearing as 
characteristic holes in the sand (SAB, personal observations). Perhaps because this ray 
species is seldom landed by commercial fishermen, has no recreational or commercial 
value, and is typically hidden, nearly completely buried in sand, it is rarely included in 
faunal surveys of beach habitat in the GoM. They seldom are landed by recreational 
fishermen, and few local citizens are aware that they range in the shallow waters (<1 m) 
of open beaches. Perhaps as a result, there is little substantive quantitative information on 
the abundance and population structure of this species throughout its range or in the 
 
 142 
 
northern GoM. Concomitantly, we know little of its general biology, symbionts (e.g., 
parasites and microbiota), and diseases.  
The 2010 BP Deepwater Horizon oil spill has focused a torrent of media attention 
on the littoral zone of the GoM, including estuarine, seagrass bed, and beach 
communities, with the implication being that the health of aquatic vertebrates and 
invertebrates in those areas are a relative metric from which oil impacts can be measured. 
As such, the interplay between bacterial communities, fish health in the coastal zone, and 
basic science on the ecology and evolution of symbionts in sharks and rays is a fruitful 
area for study. Herein, we examined and characterized the bacterial community present in 
the blood vascular system of wild-caught lesser electric rays captured from beach habitat 
in the northern GoM off Alabama. The principal aim of this study was to document the 
taxonomic and phylogenetic diversity of bacteria that infected their blood, thereby 
contributing to a broader understanding of bacterial symbionts and/or potential pathogens 
of elasmobranchs. 
Material and methods 
Sample collection. The studied rays were hand-netted off Fort Morgan, Alabama 
(30?13?45?N, 87?54?7?W), maintained alive in enclosed plastic transport containers filled 
with water from the collection site and fitted with water pumps and aerators powered by a 
car battery, and transported alive to Auburn University (within 5 h after collection). Ten 
lesser electric rays (24-47 cm in total length; 7 females and 3 males) were analyzed in the 
study. Immediately before necropsy, each ray was euthanized with an overdoes of 
tricaine methanesulfonate (MS-222), and immediately thereafter the area of skin 
 
 143 
 
circumscribed by the gill slits, mouth, and pectoral girdle was dried with a clean paper 
towel, disinfected with 70% ethanol, and cut away to expose the pericardial chamber. The 
exposed surfaces of the heart, including ventricle and conus arteriosus, were disinfected 
with 70% ethanol before a blood sample was taken by inserting a sterile syringe into the 
lumen of the heart. Each blood sample from each ray was immediately spread onto blood 
agar (BA) (MOLTOX, Boone, NC) and Marine Agar (MA) (Difco, Sparks, MD) using 
aseptic methods. Agar plates were incubated at 28?C for 48 h under aerobic conditions. A 
representative of each colony type on the primary isolation plate was re-streaked on MA 
to obtain pure cultures for identification. A total of 86 single isolates were preserved as 
glycerol stocks (Marine broth supplemented with 20% glycerol) at -80?C until subsequent 
analysis. Individual blood samples were labeled NB-01 through NB-09. The isolates were 
designated as FMR (Fort Morgan Ray) followed by the colony number.  
Bacterial identification. Bacterial isolates were identified by partially 
sequencing the 16S rRNA gene. DNA template was prepared using a rapid boiling 
method as follows. Five colonies from a pure isolate were selected from a 24-h culture on 
MA, and re-suspended in a centrifuge tube with 100 ?l sterile distilled H2O. Proteinase K 
was added to the cell suspension to a final concentration of 30 unit/?l. After 20 min 
digestion at 55 ?C, the lysate was heated to 100 ?C for 15 min, and spun down at 15,000 
g for 5 min. The supernatant was transferred to a new tube and used as template DNA. 
The nearly-complete16S rRNA gene of each isolate was amplified using the following 
primers: 63V (forward) 5'-CAGGCCTAACACATGCAAGTC-3', and 1387R (reverse) 
 
 144 
 
5'- GGGCGGWGTGTACAAGGC-3'. PCR conditions and reagents have been described 
elsewhere (8). 
Sequence analysis. Sequence trace files were edited with BioEdit version 7.1.9 
(9) to remove noise and untrusted ends. Sequences (n=3) having < 500 bp or > 3 
ambiguous positions were excluded from the analysis. The resulting 83 sequences were 
assigned to taxonomic units by i) the Ribosomal Database Project (RDP) Na?ve Bayesian 
Classifier (10) with a confidence threshold of 80%, ii) GreenGenes web classification 
tool (11), iii) BLAST (12) with a cut-off point for species ascription at 97% sequence 
similarity or higher (13). 
Phylogenetic analysis. Partial 16Sr RNA gene sequences were aligned using 
Clustal X2 (14). Multiple sequence alignment (MSA) was conducted by trimming the 
sequences to cover the entire alignment and subsequent realignment. The trimmed MSA 
spanned the hypervariable V2, V3 and V4 regions corresponding to the Escherichia coli 
16S rRNA gene base pair positions (15). Sequences of the type strains identified as 
nearest to the ray isolates by RDP and BLAST were incorporated into the phylogenetic 
trees as reference. Phylogenetic analysis was conducted in MEGA 5.0 software (16). 
Trees were constructed using the neighbor-joining method (17) with the Jukes-Cantor 
correction (18). The partial 16S rRNA gene sequences of bacterial isolates recovered 
from rays were submitted to the GenBank nucleotide sequence database (accession 
numbers KC439161 to KC439252). 
Results 
 
 145 
 
Isolate identification. All blood samples but one (NB-09) were culture positive, 
although the number of colony types (from approximately 3 to 16) varied among 
specimens (Figure 1). A total of 83 pure isolates were recovered from blood samples. 
Isolates were recovered on both marine agar (45 colonies) and blood agar (38 colonies) 
culture media. Three isolates yielded poor 16S rRNA gene sequence quality and were 
removed from the study. The remaining 83 sequences were ascribed to specific taxa using 
three databases. Overall, results from RDP, GreenGene, and NCBI were in agreement 
and isolates were identified unambiguously to genus. Isolates were classified into 14 
genera, 11 families, 6 orders, 4 classes, and 3 phyla. The majority of the isolates (91.5%) 
were ascribed to the phylum Proteobacteria, followed by the Bacteroidetes (6.0%) and the 
Actinobacteria (2.4%). In a few cases, there was a disagreement between the results 
obtained from different databases. For example, GreenGenes could not place 4 
Flavobacteriaceae isolates below family; whereas, RDP ascribed them to Arenibacter sp. 
Five isolates were ascribed to Vibrio by GreenGenes but RDP ascribed them to 
Vibrionaceae. We resolved the divergence by assigning the sequence to the lowest 
taxonomic level. 
Among the Proteobacteria, 73 isolates were Gammaproteobacteria while only 3 
isolates were identified as Alphaproteobacteria. Within the Gammaproteobacteria, Vibrio 
was the predominant genus with 45 isolates. Other isolates representing genera of 
Gammaproteobacteria comprised Pseudoalteromonas (n=13), Shewanella (n=5), 
Ferrimonas (n=2), Amphritea (n=3), Photobacterium (n=2), Thalassomonas (n=2) and 
Pseudomonas (n=1). All Alphaproteobacteria were assigned to Nautella (n=3). Figure 1 
 
 146 
 
shows the distribution of the predominant genera in each individual fish. Vibrio was the 
only genus recovered from all blood culture-positive fish. In fact, it was the most 
common genus in all fish except in NB-07 from which only 4 isolates were recovered and 
all of them belonged to different genera. Pseudoalteromonas and Shewanella were 
recovered from 5 and 4 rays, respectively. MA not only yielded more isolates but also 
provided a higher diversity of genera than BA (Figure 2), but a few genera (Gram 
positive bacteria) were recovered on BA only.  
Phylogenetic analysis.  The majority of Vibrio isolates (35 out of 45) comprised 
3 clades (Figure 3). Clade I included V. harveyi (n=14) plus the types species; Clade II = 
11 ray isolates of V. campbelli/V. sagamiensis (the partial 16S rRNA gene sequence used 
did not allow for differentiation between these two species); Clade III = 10 isolates not 
ascribed to any reference or type strain sequence but with V. nigripulchritudo as the 
closest relative.  
Analysis of the non-Vibrio Gammaproteobacteria isolates resulted in 9 principal 
groups (Figure 4). A Pseudoalteromonas clade had 13 ray isolates ascribed to P. 
phenolica (n=7), P. prydzensis (n=1), and P. spongiae (n=4). Two of the ray isolates 
could not be ascribed to named species. In two instances, a few ray isolates shared 
identical sequences but did not cluster with any reference strain. Five isolates within the 
Shewanella clade had a nonculterable bacterium as its closest neighbor and could not be 
assigned to any named species of Shewanella. Similarly, within the Amphritea clade, 3 
isolates clustered separately from all known species of Amphritea.  
 
 147 
 
The only two Actinobacteria recovered clustered along with Micrococcus 
luteus/M. yunnanensis and Microbacterium hominis (Figure 5). Two isolates of 
Bacteroidetes clustered with Arenibacter nanhaiticus, and another was the sister taxon to 
Zhouia amylolytica. The remaining two isolates were most similar to Tenibacillum sp. but 
could not be ascribed to a named species. 
Discussion 
We document that the blood of wild-caught lesser electric rays has bacteria. A 
perusal of the primary literature on the physiology, diseases, and health of elasmobranchs 
confirms that the blood of seemingly healthy wild-caught and captive elasmobranchs is 
neither sterile (i.e., free of living organisms) nor aseptic (i.e., free of microorganisms) 
(Table 1). While our results certainly contradict the paradigm that fish blood is sterile 
(19) and that isolation of bacteria from blood or other tissues results from contamination 
during necropsy (20), they agree with other published data suggesting presence of 
bacteria in elasmobranch tissues is not uncommon (21). 
Similar to the tissues of other vertebrates, elasmobranch blood harbors, in 
addition to bacteria, a diverse assemblage of parasites, including flagellates, amoebas, 
apicomplexans, microsporidians, and ciliates (see Goertz (22) and references therein). 
Elasmobranch blood comprises food for an array of metazoan parasites; including at least 
some platyhelminths (23, 24), isopods, copepods, and leeches (25). In addition, blood is 
the principal site of infection for a diversity of metazoan parasites that invade/colonize, 
mature, copulate, release progeny, and die in the blood vascular system of their shark and 
ray hosts (26-28). Still other metazoan parasites that infect internal tissues but occur as 
 
 148 
 
adults extra-intestinally may even foray into or transit within/through the blood vascular 
system in search of somatic muscle and epithelia where adult parasites deposit their eggs, 
e.g., species of Huffmanela (29, 30). In each aforementioned example, the blood is either 
invaded or exploited by a parasite, thereby plausibly exposing the blood vascular system 
to the milieu of bacteria ubiquitous in the aquatic environment. 
Bacteria, while markedly less studied in elasmobranchs than the aforementioned 
symbionts, have been detected in various elasmobranch tissues, including blood, liver, 
muscle, and epithelium (1, 31, 32) (Table 1). Similar to previous researchers who have 
documented bacteria in elasmobranchs (3), that healthy sharks can harbor bacteria in 
blood is common knowledge also among the aquatic animal health professionals who 
monitor the well being of captive elasmobranchs in the aquarium industry (32). In fact, a 
significant contribution to our understanding of blood physiology and disease has 
emerged from the aquarium industry?s monitoring of the health of sharks and rays as 
exhibit animals. From those observations numerous antimicrobial chemical therapies for 
captive elasmobranchs have been developed (33). What proportion of the bacteria known 
to associate with elasmobranchs are obligate pathogens is indeterminate; however, at 
least several are alleged primary pathogens, e.g., Vibrio harveyi (as V. carchariae) (4, 34-
36), Aeromonas salmonicida (37), and ?Flavobacterium sp.? (32). Taken together, 
however, one should not assume an elasmobranch is diseased if its blood is infected with 
parasites or bacteria nor should one assume a link between presence/absence of parasites 
and that of bacteria in the blood of elasmobranchs. We speculate that the blood vascular 
system is vulnerable to bacterial infections (i) indirectly (= via bacteria adhered to or 
 
 149 
 
residing within invading parasites) and (ii) directly (= via colonizing blood at the site of a 
hemorrhagic skin or gill lesion associated with intense infection by an ectoparasite 
population). Besides symbioses, the blood of elasmobranchs can be similarly exposed if 
the host is bitten by another predator or even the male mating partner (38). Doubtless, 
ample plausible scenarios exist for how the blood of elasmobranchs can be exposed to 
bacteria; however, we still lack a firm understanding of the taxonomic and phylogenetic 
diversity of bacteria that live in the blood of sharks and rays. 
Most isolates recovered from lesser electric rays belonged to the phylum 
Proteobacteria (91.5%), which is a result that is in agreement with previous reports (3, 
21). As expected, several species of Vibrio were isolated, including V. harveyi (17% of 
all isolates) and V. campbellii (13%). These Vibrio spp. have been previously reported as 
part of the normal flora in sharks (3). Conversely, we failed to recover any isolate 
ascribed to V. alginolyticus, a species common in sharks (4). Similarly, the common 
marine bacterium Photobacterium damselae, which has been isolated from internal 
organs of healthy fish (1, 39), was not isolated during our study. These discrepancies 
could be a factor of host-specificity, culture medium used for isolation, culture 
conditions, and habitat characteristics comprising the geographic locality where the rays 
were captured (19). Species of Pseudoalteromonas and Shewanella were common in the 
blood of the rays studied herein, and isolates representing these genera frequently have 
been reported from fish; however, only in a few instances have they been isolated from 
viscera. Some of the lesser-known bacteria isolated in the present study include 
Amphritea atlantica, Arenibacter nanhaiticus, and Zhouia amylolytica. These species or 
 
 150 
 
their closest phylogenetic species were first discovered in marine sediments (40-43). 
Species of Vibrio, Pseudoalteromonas, and Shewanella have also been recovered from 
marine sediments (44-46). Dean and Motta (47) theorized that the suction feeding 
behavior of the lesser electric ray facilitates the ingestion of sediment, and we think it is 
plausible that many bacteria would also be ingested during this feeding activity; however, 
we lack adequate behavior observations and microbial data to accept or reject this notion. 
The interstitial/benthic habitat of the lesser electric ray could drive the taxonomic 
composition of the microbiota. Regardless, how bacteria enter the blood is unknown and 
also seemingly exceedingly difficult to test in an open, natural system. A comparison of 
the present results with those from a pelagic ray that is phylogenetically-related to 
Narcine may be informative along these lines. 
The culture techniques used in this study likely underestimated the bacteria 
diversity of the tested samples since it is likely that only 1-10% of all bacteria can be 
cultured under laboratory conditions (48). We chose a culture-based strategy because 
culture methods are still the ?gold standard? in fish disease diagnostics laboratories (49) 
and because the low cost associated with this approach makes it seemingly more 
accessible to a broader spectrum of researchers. The type of culture medium, even 
general media such as MA and BA, inadvertently can select for specific bacterial groups, 
and we observed this in the present study: species of Pseudoalteromonas, Amphritea, 
Nautella, Arenibacter, Tenacibaculum, and Zhouia were recovered on MA only; whereas, 
species of Ferrimonas, Microbacterium, Micrococcus, Photobacterium, and 
Pseudomonas were recovered on BA only.  
 
 151 
 
In summary, the present study reported a high prevalence of bacteria in the blood 
of wild, apparently healthy lesser electric rays. Vibrio spp. were found in all but one 
individual and included opportunistic fish pathogens (V. harveyi) and potentially 
unnamed species (Figure 1, clade 3). Non-Vibrio Proteobacteria were also common and 
contained putative unnamed species within the Shewanella and Amphritea clades. These 
putative new species require further corroboration by full-length sequence of their 16S 
rRNA gene and additional taxonomic markers. Taken together, these insights on the 
elasmobranch microbiota are relevant to the fundamental ecology and evolutionary 
biology of aquatic symbioses. They are also vital to husbandry and veterinary staffers 
who are employed by the aquarium industry and tasked with keeping exhibited sharks 
and rays healthy; oftentimes following protocols that use blood picture as a means of 
assessing overall health status of the exhibited elasmobranch. 
Acknowledgments 
This work was funded by the National Oceanic and Atmospheric Administration 
(NOAA-NA08NMF4720545), Marine Environmental Science Consortium, GoM 
Research Institute, and National Science Foundation (NSF DEB Nos 
 1112729, 1051106, and 1048523) and is a contribution of the Southeastern 
Cooperative Fish Parasite and Disease Project (Auburn University). Zhen Tao is the 
recipient of graduate research fellowship funded by the Chinese Scholarship Council and 
Ocean University of China. 
 
  
 
 152 
 
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Table AP-1. Taxonomic diversity of bacterial isolates from reportedly asymptomatic 
elasmobranchs.* 
Host species Bacterial species Tissue 
Carcharhinus acronotus  Staphylococcus epidermidis blooda 
(blacknose shark)   
Carcharhinus limbatus Photobacterium damsela  blooda, c 
(blacktip shark) Vibrio alginolyticus blooda, c 
 Vibrio parahaemolyticus blooda, b 
 Photobacterium sp. kidneyc 
 Vibrio furnissii  kidneyb 
 Vibrio harveyi kidneyc 
 Vibrio sp. kidneyc 
 Vibrio sp. liverb 
 Vibrio harveyi liverc 
 Vibrio alginolyticus pancreasc 
 Photobacterium damsela spleenc 
Carcharhinus melanopterus Aeromonas hydrophila  blooda 
 (blackfin reef shark) Aeromonas caviae blooda 
 Alcaligenes spp. blooda 
 Chryseomonas luteola blooda 
 Citrobacter freundii blooda 
 Citrobacter youngae blooda 
 Morganella morganii blooda 
 Pasteurella pneumotropica blooda 
 Photobacterium damsela blooda 
 Pseudomonas aeruginosa blooda 
 Pseudomonas alcaligenes blooda 
 Pseudomonas fluorescens blooda 
 Pseudomonas stutzeri blooda 
 Shewanella putrefaciens blooda 
 Sphingomonas paucimobilis blooda 
 Staphylococcus epidermidis blooda 
 Vibrio alginolyticus blooda 
Carcharhinus plumbes Vibrio vulnificus blooda 
(sandbar shark) Staphylococcus epidermidis blooda 
Cephaloscyllium ventriosum  Staphylococcus epidermidis blooda 
(swell shark) Stenotrophomonas maltophilia blooda 
Chiloscyllium plagiosum  Vibrio alginolyticus blooda 
(whitespotted bamboo shark) Vibrio vulnificus blooda 
Galeocerdo cuvier Vibrio alginolyticus bloodc 
(tiger shark) Vibrio sp. bloodc 
 Vibrio alginolyticus liverc 
Ginglymostoma cirratum  Photobacterium damsela bloodc 
(nurse shark) Vibrio harveyi liverb 
 Vibrio harveyi pancreasb 
 Vibrio harveyi spleenb 
Himantura granulata Proteus vulgaris blooda 
(Mangrove whiptail ray) Vibrio vulgaris blooda 
 Morganella morganii blooda 
 Photobacterium damsela blooda 
 Vibrio alginolyticus blooda 
 
 159 
 
Table AP-1(Continued) 
Host species Bacterial species Tissue 
Mustelus canis    Vibrio metschnikovii   trunk kidneyd 
(spiny dogfish) Photobacterium damsela   trunk kidneyd 
Mustelus canis    Pseudomonas sp.   trunk kidneyd 
(spiny dogfish)   
Negaprion brevirostris Vibrio alginolyticus bloodb, c 
(lemon shark) Vibrio sp. bloodb, c 
 Photobacterium damsela bloodb 
 Vibrio alginolyticus eyeb 
 Vibrio sp. eyeb 
 Photobacterium damsela gall bladderb 
 Vibrio harveyi gill slitb 
 Vibrio furnissii gill slitb 
 Vibrio sp. gill slitb 
 Vibrio alginolyticus musclec 
 Vibrio harveyi musclec 
 Aeromonas salmonicida musclec 
 Photobacterium damsela kidneyb 
 Photobacterium damsela spleenc 
Orectolobus japonicus  Photobacterium damsela blooda 
(bearded shark)   
Potamotrygon sp.  Plesiomonas shigelloides blooda 
(freshwater stingrays) Pseudomonas fluorescens   blooda 
 Pseudomonas putida blooda 
 Vibrio fluvialis blooda 
 Staphylococcus epidermidis blooda 
Pristis zijsron  Pasteurella pneumotropica blooda 
(Narrowsnout sawfish)   
Rhizoprionodon terraenovae  Vibrio sp. liverb 
(sharpnose shark)   
Squalus acanthias  Alteromonas sp.   trunk kidneyd 
(smooth dogfish) Vibrio alginolyticus   trunk idneyd 
 Shewanella putrefaciens   trunk kidneyd 
 Photobacterium damsela   trunk kidneyd 
 Vibrio fluvialis   trunk kidneyd 
 Pseudomonas putida   trunk kidneyd 
Triaenodon obesus  Moraxella spp. blooda 
(whitetip reef shark) Streptococcus group D blooda 
 Pasteurella pneumotropica blooda 
 Photobacterium damsela blooda 
 Shewanella putrefaciens blooda 
 Staphylococcus epidermidis blooda 
 Proteus vulgaris blooda 
Triakis semifasciata Vibrio alginolyticus blooda 
(leopard shark)    
*Based on the published literature and from search results of keywords bacteria + 
elasmobranch? retrieved from bibliographic databases provided by ISI Web of 
 
 160 
 
Knowledge (http://apps.webofknowledge.com, Thompson Reuters 2013) and Science 
Direct (http://www.sciencedirect.com/, Elsevier B.V. 2013) 
Table AP-1 literature references:   
a. Mylniczenko ND, Harris B, Wilborn RE. 2007. Blood culture results from healthy 
captive and free-ranging elasmobranchs. Journal of Aquatic Animal Health 19:159-167. 
b. Grimes DJ, Brayton PR, Colwell RR, Gruber S. 1985. Vibrios as autochthonous flora 
of neritic sharks. Systematic and Applied Microbiology 6:221-226. 
c. Grimes DJ, Jacobs D, Swartz DG, Brayton PR, Colwell RR. 1993. Numerical 
taxonomy of Gram-negative, oxidase-positive rods from Carcharhinid sharks. 
International Journal of Systematic Bacteriology 1993:88-98. 
d. Borucinska JD, Frascas S. 2002. Naturally occurring lesions and micro-organisms in   
two species of free living sharks: the spiny dog fish, Squalus acanthias L., and the  
 smooth dogifhs, Mustelus canis (Mitchill), from the north-western Atlantic. Journal of 
 Fish Diseases 25:287-298. 
 
  
 
 161 
 
 
Figure AP-1. Bacteria recovered from blood of lesser electric rays (Narcine bancroftii, 
NB). 
  
 
 162 
 
 
Figure  AP-2. Distribution of isolates from blood of lesser electric rays (Narcine 
bancroftii) cultured in marine agar (MA) and blood agar (BA). 
  
 
 163 
 
 
 
 164 
 
Figure  AP-3. Phylogeny (partial 16S rRNA gene sequences) of bacterial isolates from 
blood of Narcine bancroftii and ascribed to species of Vibrio. Isolate number is followed 
by GenBank accession number. Sequences from type strains or the closest match were 
used for comparison. The tree topology was obtained by the neighbor-joining methods 
(Jukes-Cantor correction). The three main clades are highlighted. Numbers at nodes 
indicate boostrap values (1000 replicates). Bar = 0.5% sequence divergence. 
  
 
 165 
 
 
 
 166 
 
Figure AP-4. Phylogeny (partial 16S rRNA gene sequences) of bacterial isolates from 
blood of Narcine bancroftii and assigned as non-Vibrio Proteobacteria species. Isolate 
number is followed by GenBank accession number. Sequences from type strains, or the 
closest match, were used for comparison. The tree topology was obtained by the 
neighbor-joining methods (Jukes-Cantor correction). Each genus-clade is highlighted. 
Numbers at nodes indicate bootstrap values (1000 replicates). Bar = 5% sequence 
divergence.  
 
 167 
 
 
Figure AP-5. Phylogeny (partial 16S rRNA gene sequences) of bacterial isolates from 
blood of Narcine bancroftii and assigned as non-Proteobacteria. Isolate number is 
followed by GenBank accession number. Sequences from type strains or the closest 
match were used for comparison. The tree topology was obtained by the neighbor-joining 
methods (Jukes-Cantor correction). Each genus-clade is highlighted. Numbers at nodes 
indicate bootstrap values (1000 replicates). Scale bar represents 5% sequence divergence.