AN ABSTRACT OF THE DISSERTATION OF
Tracy Shawn Peterson for the degree of Doctor of Philosophy in Microbiology presented
on February 27, 2013.
Title: Detection and Transmission of Mycobacterium marinum and Mycobacterium
chelonae in Zebrafish (Danio rerio).
Abstract approved: __________________________________________________
Michael L. Kent
Mycobacteriosis is a common disease of laboratory zebrafish (Danio rerio).
Different infection patterns occur in zebrafish depending on mycobacterial species.
Mycobacterium marinum and M. haemophilum produce virulent infections associated
with high mortality, whereas M. chelonae is more wide spread and not associated with
high mortality. Identification of mycobacterial infections to the species level provides
important information for making management decisions. Observation of acid-fast
bacilli in histological sections or tissue imprints is the most common diagnostic method
for mycobacteriosis in fish, but only allows for diagnosis to the genus level.
Mycobacterial culture, followed by molecular or biochemical identification is the
traditional approach for species identification, but recently it has been shown that DNA
of diagnostic value can be retrieved from paraffin blocks. Type of fixative, time in
fixative before processing, species of mycobacteria, and severity of infection were
investigated as parameters to determine if the hsp gene PCR assay (primer set
HS5F/hsp667R) could detect and amplify mycobacterial DNA from paraffin-embedded
zebrafish. Whole zebrafish were experimentally infected with either M. chelonae or M.
marinum, and then preserved in 10% neutral buffered formalin or Dietrich’s fixative for
3, 7, 21 and 45 days. Subsequently, fish were evaluated by H&E and Fite’s acid-fast
stains to detect mycobacteria within granulomatous lesions. The PCR assay was quite
effective, and obtained PCR product from 75% and 88% of the M. chelonae and M.
marinum infected fish, respectively. Fixative type, time in fixative, and mycobacterial
species showed no statistical relationship with the efficacy of the PCR test.
Regarding natural transmission, zebrafish are capable of contracting
mycobacterial infections by feeding on infected fish tissue, but other natural routes have
not been clearly elucidated. Free living amoebae have been shown to be vectors for
mycobacteria and their virulence is enhanced when residing in these protozoans.
Paramecium caudatum are commonly used as a first food for zebrafish, and I
investigated this ciliate’s potential to serve as a vector of Mycobacterium marinum and
M. chelonae. The ability of live P. caudatum to transmit these mycobacteria to larval,
juvenile and adult zebrafish was evaluated. Infections were defined by histologic
observation of granulomas containing acid-fast bacteria in extraintestinal locations. In
both experiments, fish fed paramecia containing mycobacteria became infected at a
higher incidence than controls. Larvae (exposed at 4 days post hatch) fed paramecia with
M. marinum exhibited an incidence of 30% (24/80) and juveniles (exposed at 21 days
post hatch) showed 31% incidence (14/45). Adult fish fed gelatin diets containing
bacteria within paramecia or mycobacteria alone for 2 wk resulted in infections when
examined 8 wk after exposure: M. marinum OSU 214; in paramecia 47% (21/45; 3.5 x
105 dose/fish/day), M. marinum CH in paramecia 47% (9/19; 3.6 x 105 dose/fish/day), M.
chelonae in paramecia 38% (5/13; 3.5 x 105 dose/fish/day).
I investigated the ability of mycobacteria to persist within paramecia, as this has
previously been demonstrated in amoebae. Gram negative bacteria ingested by paramecia
were processed within an hour. In contrast, I determined using GFP-labeled
Mycobacterium marinum that mycobacteria can persist within paramecia digestive
vacuoles. The concentration of M. marinum at 1 hour was similar to that at the time of
ingestion. Twenty-four hours post-ingestion and later there was significant decline in M.
marinum concentrations compared to time of ingestion, but M. marinum continued to
persist inside digestive vacuoles for up to one week. My results demonstrate for the first
time that Paramecium caudatum can act as a vector for mycobacteria. This provides a
useful animal model for evaluation of natural mycobacterial infections and demonstrates
the possibility of mycobacterial transmission in zebrafish facilities via contaminated
paramecia cultures.
©Copyright by Tracy Shawn Peterson
February 27, 2013
All Rights Reserved
Detection and Transmission of Mycobacterium marinum and Mycobacterium
chelonae in Zebrafish (Danio rerio)
by
Tracy Shawn Peterson
A DISSERTATION
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Doctor of Philosophy
Presented February 27, 2013
Commencement June 2013
Doctor of Philosophy dissertation of Tracy Shawn Peterson presented on
February 27, 2013.
APPROVED:
Major Professor, representing Microbiology
Chair of the Department of Microbiology
Dean of the Graduate School
I understand that my dissertation will become part of the permanent collection of Oregon
State University libraries. My signature below authorizes release of my dissertation to
any reader upon request.
Tracy Shawn Peterson, Author
ACKNOWLEDGEMENTS
Mike Kent has been a stalwart mentor over the years. He is the definition of
success in his field and in his life. I would like to thank him for providing me with
numerous opportunities to advance my career. What separates him from most mentors
that I have worked with is that he seeks to develop his students both personally and
professionally. This is a rare phenomenon in academia and I know Mike has positively
impacted the lives of many former and present students. I would like to thank the
members of my committee for their service and contributions to my work: Jerry Heidel,
Christiane Löhr, Bruce Geller and Larry Curtis. Robert Tanguay has been instrumental in
helping me to establish my career, by giving me the opportunity to be the first student
selected for the NIH/NCRR grant T32 RR023917. Thank you to all those uncommon and
brave individuals who were fit to “pack the gear” and wage war by my side in the
Coliseum of Academia. May your struggles find worthiness in this desolate place that
hardens the heart and softens the mind, and may your battles be symmetrical, thus
providing insight to those who shall follow. Lastly, I wish to extend profound gratitude to
the common and unpredictable enemy, the zebrafish, those tenacious aquatic guerrilla
fighters that they are, who in the darkest hours of my research valiantly and unselfishly
gave their lives and suffered many indignities in this protracted asymmetrical war, so that
others of their kind may live. There is no higher or nobler sacrifice.
CONTRIBUTION OF AUTHORS
Justin Sanders provided both photographic and statistical assistance as well as advice on
experimental design for the paramecia experiments and photographic editing for the brain
tumor descriptive work. Doctors Christopher Whipps and Jayde Ferguson assisted with
performance and data gathering on the PCR assays and statistical interpretations,
respectively, for the PCR work and Doctor Jayde Ferguson again assisted with statistical
interpretations on the transmission studies performed for this thesis. Doctors Stephen
Feist and Jan Spitsbergen provided microsporidian samples and salient wisdom for the
Luna stain project. Doctors Don Ennis and Nadine Mutoji donated the GFP-labeled
Mycobacterium marinum used in the paramecia studies and gave me the recipe for the
gelatin feed matrix as well as inspiration to pursue novel methods of mycobacterial
transmission, due to their groundbreaking work with mosquito larvae. Doctors Jerry
Heidel and Katy Murray provided zebrafish samples that resulted in discovery of an
entirely new brain tumor affecting zebrafish and gave critical insights into the
development of this tumor. Virginia Watral assisted with injection of zebrafish and
provided excellent technical support for the PCR and transmission studies.
TABLE OF CONTENTS
Page
Chapter 1. Introduction……………………………………………………………………1
Historical perspective of piscine mycobacteriosis………………………………...1
Mycobacterial species of importance……………………………………………...3
Pathology of piscine mycobacteriosis……………………………………………..4
Transmission of mycobacteria in fish……………………………………………..5
Treatment and control of mycobacteriosis in fish………………………………..10
Mycobacterial diagnostics in fish………………………………………………..14
Piscine mycobacteriosis and human health concerns……………………………16
Mycobacteriosis in fishes used in biomedical research………………………….17
Models of mycobacteriosis in laboratory fishes…………………………………20
Mycobacteriosis in laboratory zebrafish…………………………………………24
Summary…………………………………………………………………………29
References………………………………………………………………………..30
Chapter 2. Comparison of fixatives, fixation time, and severity of
infection on PCR amplification and detection of Mycobacterium marinum
and Mycobacterium chelonae DNA in paraffin-embedded zebrafish
(Danio rerio)……………………………………………………………………………..46
Abstract…………………………………………………………………………..47
Introduction………………………………………………………………………48
Material and Methods ……………………………………………………………50
Results ……………………………………………………………………………55
TABLE OF CONTENTS (Continued)
Page
Discussion………………………………………………………………………..56
References………………………………………………………………………..62
Chapter 3. Paramecium caudatum enhances transmission and infectivity
of Mycobacterium marinum and Mycobacterium chelonae in zebrafish
(Danio rerio)……………………………………………………………………………..68
Abstract…………………………………………………………………………..69
Introduction………………………………………………………………………70
Materials and Methods…………………………………………………………...72
Results……………………………………………………………………………79
Discussion………………………………………………………………………..84
References………………………………………………………………………..94
Chapter 4. Persistence of Mycobacterium marinum in digestive vacuoles
of Paramecium caudatum………………………………………………………………..99
Abstract…………………………………………………………………………100
Introduction……………………………………………………………………..101
Materials and Methods………………………………………………………….102
Results …………………………………………………………………………..106
Discussion………………………………………………………………………107
References………………………………………………………………………116
Chapter 5. Discussion…………………………………………………………………..119
Preface………………………………………………………………………….119
TABLE OF CONTENTS (Continued)
Page
Detecting mycobacterial pathogens with PCR from paraffin-embedded
zebrafish tissues………………………………………………………………...............119
Unlocking the mystery of mycobacterial transmission in zebrafish……………123
References………………………………………………………………………134
Chapter 6. Summary and Conclusions………………………………………………….142
Bibliography……………………………………………………………………………145
Appendix………………………………………………………………………………..170
The Luna stain, an improved selective stain for detection of microsporidian spores
in histologic sections……………………………………………………………………171
Abstract………………………………………………………………………….172
Introduction……………………………………………………………………...172
Materials and Methods…………………………………………………………..174
Results…………………………………………………………………………...175
Discussion……………………………………………………………………….177
References……………………………………………………………………….185
Malignant dysembryoplastic neuroepithelial tumour in a zebrafish (Danio rerio)…….187
Summary………………………………………………………………………...188
Note……………………………………………………………………………...188
References……………………………………………………………………….199
LIST OF FIGURES
Figure
Page
Figure 3.1 Histological presentations in zebrafish exposed to M. marinum and
M. chelonae residing within paramecia food vacuoles………….……………………….90
Figure 3.2 Prevalence of extraintestinal mycobacteriosis and presence of
mycobacteria in the intestinal lumen…………………………………………………….91
Figure 4.1 Paramecia containing GFP-labeled Mycobacterium marinum……………...114
Figure 4.2 Concentration of GFP-labeled M. marinum in paramecia…………………..115
Appendix Figure 1.1 Microsporidia in histologic stains from various fishes…………..181
Appendix Figure 1.2 Microsporidia in histologic stains from various fishes…………..182
Appendix Figure 2.1 Tumour growth pattern and infiltrative nature…………………..196
Appendix Figure 2.2 Microcystic pattern of tumour growth…………………………...197
Appendix Figure 2.3 Abnormally oriented and misshapen neurons……………………198
LIST OF TABLES
Table
Page
Table 2.1 Proportion of PCR positives and mean granuloma intensity………………….61
Table 3.1 Larval and juvenile zebrafish exposed by feeding Paramecium caudatum
containing mycobacteria…………………………………………………………………92
Table 3.2 Adult zebrafish fed gelatin food matrix containing mycobacteria or
Paramecium caudatum containing mycobacteria………………………………………..93
Appendix Table 1.1 Characteristics of microsporidian spores in histological sections
with special stains……………………………………………………………………....184
Chapter 1. Introduction
Historical Perspective of Piscine Mycobacteriosis
Of all the diseases affecting fishes, mycobacteriosis remains one of the most
studied and perplexing. My thesis attempts to provide valuable insight into the disease, as
it pertains to accurate species level diagnosis in laboratory zebrafish, and to provide an
answer as to how it is transmitted among zebrafish and by extension, other fishes.
Mycobacterial infections involving several different species of aquatic mycobacteria
occur across a broad taxonomic range of fishes worldwide, reported in at least 160
different species (Kaattari et al. 2006). The disease occurs in diverse ecological habitats
ranging from the tropics to the arctic and in brackish, salt and fresh water. Confirmed
mycobacterial infections have been reported in multiple wild fish species encompassing
different genera and occasionally intraspecific fishes, ranging from salmonids and
sturgeon to reef and tropical fishes (Diamant et al. 2000, Bercouvier and Vincent 2001,
Jacobs et al. 2009). Many of these wild fish were introduced into aquaculture and along
with other fish have been domiciled, intensively managed or harvested as a food source,
and such examples include Chinook salmon (Oncorhyncus tshawytscha), Atlantic salmon
(Salmo salar), rainbow trout (Oncorhyncus mykiss), cod (Gadus morhua L.), hybrid
striped bass (Morone saxatilis x Morone chrysops), sea bass (Dicentrachus labrax),
sturgeon (Acipenser spp.), carp (Cyprinus carpio L.), turbot (Scophthalmus maximus),
California rockfish (Sebastes mystinus), sablefish (Anoplopoma fimbria), mackerel
(Scomber scombrus L.), yellowtail (Seriola quinqueradiata), greater amberjack (Seriola
dumerili), yellowtail amberjack (Seriola lalandi), striped jack (Pseudocaranx dentex),
2
sevenband grouper (Epinephelus septemfasciatus) and tilapia (Oreochromis
mossambicus). (Nakanaga et al. 2012, Imajoh et al. 2012, Majeed and Gopinath 1983,
Moser et al. 1986, Moser and Sakanari 1986, Diamant et al. 2000, dos Santos et al. 2002,
Hastings et al. 1982, Ashburner 1977, Arakawa and Fryer 1984, Bruno et al. 1998,
Gauthier et al. 2003, Heckert et al. 2001, Hedrick et al. 1987, Noga et al. 1990, Parisot
1958, Ross 1959, Sakanari et al. 1983, Wolf and Smith 1999).
Historically, mycobacteriosis has been reported among wild, cultured and
commercial trade fishes going back almost 120 years to the turn of the century, when
Bataillon et al. (1897) discovered acid-fast bacilli associated with granulomatous
“tuberculosis” disease in a moribund carp, which was caught in a pond contaminated with
human sputum from tuberculosis patients (Wolke and Stroud 1978). This was not the
likely source of the mycobacteria, as human Mycobacterium tuberculosis and piscine
Mycobacterium marinum are phylogenetically and genotypically distinct bacteria despite
being closely related (Tobin and Ramakrishnan 2008). After this discovery, they further
characterized the bacterial isolate in a series of papers and compared it with human and
avian isolates (Bataillon et al. 1897, Bataillon and Terre 1897, Bataillon et al. 1902). The
earliest 20th century reports of mycobacteriosis in fish, not including the original
observations of “tuberculosis” or “tubercular disease” in cod (Gadus callarias) and
halibut (Hippoglossus hippoglossus) where the causal agent was not identified, were
made separately by Aronson and Baker in salt and fresh water fish, respectively (Aronson
1926, Baker 1943). In both of these discoveries, the bacteria responsible for the observed
lesions was subsequently identified and helped define what would become the
3
paradigmatic aquatic mycobacterial species, Mycobacterium marinum. There has been
several comprehensive and exhaustive review papers previously published on piscine
mycobacteriosis, establishing it as a well-characterized and understood disease in fishes
(Parisot 1958, Snieszko 1978, Wolke and Stroud 1978, Bercouvier and Vincent 2001,
Decostere et al. 2004, Gauthier and Rhodes 2009, Jacobs et al. 2009).
Mycobacterial Species of Importance
Originally, when M. marinum was recovered, cultured and isolated from a variety
of salt water fishes at the Philadelphia aquarium (sergeant major, Abudefduf mauritii;
croaker, Micropogon undulatus; sea bass, Centropristes striatus; lane snapper, Lutjanus
griseus) and the Mexican platyfish (Platypoecilus maculatus), it was misclassified by
Aronson as Mycobacterium piscium and subsequently in Baker’s report as well, as
Mycobacterium platypoecilus. In both of these initial characterizations of M. marinum,
isolated cultures were re-introduced to fish experimentally, following Koch’s postulates
and were found to cause identical “tubercular disease” in the experimental hosts. By
contrast, few mycobacterial infections of salmonid or cold water fishes have been
documented (Bruno et al. 1998, Whipps et al. 2007a) and this likely reflects both the
range of growth temperatures of mycobacteria, between 18 and 28 oC, and the relatively
low ambient water temperature inhabited by salmonids, although mycobacteria can
survive at temperatures lower than 20 oC for several months and even years (Wayne and
Kubica 1986). Mycobacterial infections have been reported from deep water marine fish
(Sebastes spp.) living at temperatures <10oC (Whipps et al. 2003).
4
The occasional culture and isolation of atypical non-tuberculous mycobacterial
species in ornamental fish, such as Mycobacterium terrae, Mycobacterium gordonae,
Mycobacterium peregrinum, Mycobacterium scrofulaceum, Mycobacterium interjectum
and others have been reported (Lansdell et al. 1993, Prearo et al. 2004, Zanoni et al.
2008). In many, but not all, of these unusual cases where mycobacterial infection was
suspected, the culture and identification of some atypical mycobacteria can be attributed
to extrinsic contamination from the water source (Goslee and Wolinski 1976) and in the
absence of correlating clinical disease with a truly pathogenic mycobacteria, are more
properly termed a pseudoinfection or pseudo-outbreak (Panwalker and Fuhse 1986,
Lalande et al. 2001).
Pathology of Piscine Mycobacteriosis
The insidious and often clinically inapparent nature of mycobacterial infections in
both cultured and aquarium fishes, as well as zebrafish (Danio rerio), makes it difficult to
diagnose in live fish, as infected fish may have no clinical signs or may have non-specific
external signs such as lepidorthosis, dermal ulceration, scale loss, exophthalmia,
hyperpigmentation (dermomelanosis), integumentary hyperemia and hyphema. Internal
examination of affected fish often gives the initial evidence of mycobacterial infection,
by visual detection of granulomas in tissues, most commonly the liver, kidney and spleen
and thus infection can be regarded as a systemic disease (Ferguson 2006).
The hallmark microscopic lesion is considered to be granulomatous inflammation
with formation of characteristic granulomata, which are comprised of concentric layers of
histiocytes (epitheloid cells) ensheathed by fibrocytes with scant to minimal numbers of
5
lymphoplasmacytic inflammatory cells surrounding individual granulomas. The centers
of these granulomata frequently contain caseonecrotic debris and in typical cases of
mycobacteriosis, acid-fast positive bacterial rods can be observed both within the necrotic
material and intracellularly in the adjacent macrophages. Granulomas participating in
control of mycobacterial infections have a morphologically definable lifespan, from
loosely organized aggregates of histiocytes early in the infective process that eventually
mature into well-organized and discrete encapsulating structures as the chronic infection
persists (HW Ferguson 2007). Multinucleated giant cells and the characteristic
lymphoplasmacytic infiltrate surrounding granulomas, which are common in mammalian
mycobacterial infections, are infrequently seen in fish (Astrofsky et al. 2000, Gauthier
and Rhodes 2009). The target organs in piscine mycobacterial infections include the
blood filter organs such as spleen, liver and kidney (Jacobs et al. 2009) although in
especially severe infections multiple organs are affected, including the choroid rete, heart,
gills, skin and gas (e.g. swim) bladder. In some cases of zebrafish Mycobacterium
haemophilum infections, the central nervous system is prominently infected in addition to
the typical organs (Whipps et al. 2007b).
Transmission of Mycobacteria in Fish
There are several known and widely-accepted modes of mycobacterial
transmission, representing non-natural experimental infections, which includes
intraperitoneal and intramuscular injection, oral gavage and bath exposure (Prouty et al.
2003, Harriff et al. 2007, Watral and Kent 2007). It is of critical importance to
understand the possible origins of mycobacteria within an aquatic laboratory system, in
6
order to implement appropriate control strategies. Potential sources of infection in the
water source as well as biofilm in water pipes and tanks and live food items, particularly
those fed to larval zebrafish, that can serve as hosts and vectors for mycobacterial
transmission are major concerns of zebrafish husbandry.
There are currently three routes that established the hypothesis for natural
transmission of mycobacteriosis, all of them oral-enteric and which have substantial
confirmatory evidence of natural transmission of mycobacteria among fishes. These
routes of transmission include ingestion of contaminated food, cannibalism of infected
fish and feeding on environmental detritus (Post 1987, Chinabut et al. 1990, Grady et al.
1992, Decostere et al. 2004). Direct proof of the transmission cycle first became known
in 1958, when Wood and Ordal observed 100% infection rates in salmon fry and
fingerlings from Pacific Northwest hatcheries that were fed contaminated, unpasteurized
ground fish meal (Wood and Ordal1958). Current information regarding natural
transmission and vectors of mycobacteria in laboratory fishes, particularly zebrafish, is
not substantive.
There are several known and utilized methods of mycobacterial transmission in
zebrafish and Japanese medaka (Oryzias latipes), representing non-natural experimental
infections, which includes intraperitoneal and intramuscular injection, oral gavage and
bath exposure (Prouty et al. 2003, Harriff et al. 2007, Watral and Kent 2007). Per os
infection by feeding on infected fish tissues is one recognized route of transmission (Ross
1970, Hedrick et al. 1987, Mutoji 2011) and could be considered as a more natural route
of infection. It has been previously shown that mycobacteria residing within granulomas
7
have increased virulence, which provides an explanation for the ease of per os infections
by feeding on infected tissue (Volkman et al. 2004). Water bath exposure of medaka
(Oryzias latipes) to mycobacteria directly from cultures is ineffective and does not result
in establishing a high prevalence of infection in laboratory transmission studies (Mutoji
2011).
Free-living aquatic protozoans, such as Acanthamoeba and Dictyostelium,
evolved as phagocytic cells that actively prey on bacteria within air-water interface
biofilms (Barker and Brown 1994) and inhabit the same niches as environmental
mycobacteria. These protozoans ingest the mycobacteria into food vacuoles, where they
are processed for energy then eliminated via excretory vacuoles, or in some instances
may persist within the protozoan host. From an evolutionary perspective, it is possible
that mycobacteria and certain Gram-negative bacteria (i.e. Legionella pneumophila) may
have adapted to survive intracellularly in amoebae with the development of eventual
endosymbiosis (Winiecka-Krusnell and Linder 2001). In support of this concept, Jadin
(1975) as well as Krishna Prasad and Gupta (1978) discovered that Mycobacterium
leprae and other mycobacteria not only survived within Acanthamoeba spp., but could
actively replicate in the organism.
Since those initial reports, many more mycobacterial species (up to 26 different
species, including Mycobacterium marinum and Mycobacterium chelonae) have been
discovered that exploit a potential endosymbiotic relationship with free-living amoebae,
making a living within Acanthamoeba spp. trophozoites and amoebic cysts or cyst walls
(Steinert et al. 1998, Adekambi et al. 2006). The ability to capitalize on this potential
8
symbiosis, with residence in amoebic cysts, ultimately would protect mycobacteria from
suboptimal environmental conditions including exposure to chlorine, chlorine
compounds, ozone and metal ions (Thomas et al. 2004, Whan et al. 2006) and
antimicrobial compounds that constitute front-line chemotherapy for mycobacterial
infections (Miltner and Bermudez 2000).
Although an endosymbiotic relationship exists between amoebae and
mycobacteria, it should be noted that many bacterial pathogenicity traits co-evolved
along with amoebae as a means to resist predation and digestion, such as bacterial
encapsulation, toxin secretion, blockage of phagolysosome fusion and intracellular
replication within the host amoeba (Cosson and Soldati 2008). As an example, it has been
demonstrated by Cirillo (1997) that Mycobacterium avium pathogenicity is increased if it
is phagocytized by Acanthamoeba castellanii and replicates within amoebic vacuoles,
specifically due to M. avium stopping phagolysosome fusion and enhanced cellular entry
and intracellular replication. Solomon et al. (2003), in concordance with this observation,
found that M. marinum effectively replicates within Dictyostelium spp. by halting
phagosomal maturation. Mechanisms of cellular invasion elicited by M. marinum may be
putatively conserved between amoebae and mammalian macrophages (Hagedorn and
Soldati 2007).
Live foods, typically aquatic protozoans, worms and crustacean larvae, are a
dietary staple of both larval and adult zebrafish because they offer balanced nutritional
profiles (Watanabe et al. 1983, Lawrence 2007, Best et al. 2010). Most commonly, these
foods consist of prey items such as paramecium, rotifers, water fleas (Daphnia spp.),
9
Artemia and chironomid larvae (bloodworms). The critical window for feeding larval
zebrafish occurs around five days post-fertilization, when the larval fish inflate their gas
bladder and can begin to move through the water column in active search for prey items.
It is also around this time that the larval zebrafish have depleted their yolk reserves and
must begin to feed in order to grow and survive. Paramecia and rotifers are considered
the ideal first feeding prey items, as they are relatively slow-moving and have the correct
size for easy ingestion by the larval fish. As the fish matures, at around 10 to 12 days
post-fertilization, larger prey items such as Artemia and water fleas can be offered. It is
known, as discussed previously, that environmental protozoans actively graze biofilms
for bacteria and that mycobacteria take up residence in other environmental protozoans
(amoebae) and by doing so, their pathogenicity and survivability increases significantly.
It is also established that the zebrafish gastrointestinal tract is the primary site of entry for
mycobacteria (Harriff et al. 2007).
Another potential route of infection is by feeding on invertebrates colonized by
mycobacteria. Tubificid worms (Tubifex tubifex) and water fleas (Daphnia spp.) in
particular have been implicated as a source of mycobacteria within established
aquaculture systems (Grange 1985, Conroy and Conroy 1999, Somsiri et al. 2005, Nenoff
and Uhlemann 2006). Mutoji (2011) showed that mosquito larvae infected with M.
marinum induced a high prevalence of infection in medaka following feeding on these
larvae. Mycobacteria residing within amoebae are also more infectious than their
counterparts from axenic cultures. Cirillo et al. (1997) showed that M. avium
pathogenicity is increased if it is phagocytized by A. castellanii and replicates within
10
amoebic vacuoles. Free-living aquatic protozoans, such as Acanthamoeba and
Dictyostelium, evolved as phagocytic cells that actively prey on bacteria within air-water
interface biofilms (Barker and Brown 1994) and inhabit the same niches as
environmental mycobacteria. Mycobacteria residing in macrophages have enhanced
virulence, and hence these amoebae act as surrogate macrophages in the environment.
Harriff et al. (2007) demonstrated that per os infections of M. marinum in zebrafish were
enhanced by passing the bacteria through amoebae.
Whereas amoebae are not deliberately feed to zebrafish, the ciliate Paramecium
caudatum is a common first food for larval zebrafish (Westerfield 2007, Harper and
Lawrence 2011) and similar to environmental amoebae, they are indiscriminant
bacterivores. Therefore, I conducted experiments to determine if P. caudatum could serve
as a vector and enhance natural transmission of M. marinum and M. chelonae in larval,
post-larval and adult zebrafish (Chapter 3).
Treatment and Control of Mycobacteriosis in Fish
Mycobacteriosis in zebrafish presents many challenges to effective management
and control of the disease once established within research colonies (Astrofsky et al.
2000, Kent et al. 2009). The mycobacterial species infecting laboratory zebrafish
including M. marinum, M. haemophilum and M. chelonae, can be commonly isolated
from the local environment especially water sources, pipes and tanks (Primm et al. 2004,
Whipps et al. 2007b, Whipps et al. 2008). Persistence and replication of mycobacteria in
the aquatic environment is aided by the bacteria’s inherent resistance to chlorine, survival
in low-nutrient environments, (starvation resistance), thermoresistance and ability to form
11
biofilms (Nyka 1974, Schulze-Robbecke and Buchholtz 1992, Schulze-Robbecke et al.
1992, Hall-Stoodley and Lappin-Scott 1998, Taylor et al. 2000, Le Dantec et al. 2002,
September et al. 2004). These capabilities enable mycobacteria to establish populations
and persist in flowing water systems, like those found in the laboratory setting, despite
relatively slow growth. Mycobacterial species capable of causing disease in fish are
common in aquaria and aquatic habitats, but often infections in fish do not occur when
held in the same system or environment that contains the bacteria (Beran et al. 2006).
Traditional treatment and control methods that were initially established for
piscine mycobacteriosis centered on individual fish within a population within the
ornamental fish industry, and at the whole population level of hatchery fish (Ross 1970).
The usual approach at the facility level has been depopulation of infected stocks of
aquarium and hatchery fish (Chinabut 1999). Antimicrobial treatment of mycobacteriosis
in fish cultured for human consumption is impractical because combating mycobacterial
infections involves an economy of scale, such as the large numbers of fish that must be
treated, bioavailability of the antimicrobial drugs in fish, feasible delivery of the drug,
relatively high costs of antibiotics, treatment regimens lasting weeks to months, and
protracted drug withdrawal times in fish destined for human consumption.
Mycobacterium chelonae, which is widely distributed in many laboratory fish
facilities, as well as other opportunistic mycobacterial species, rapidly recolonize surface
biofilms in laboratories within about six weeks following complete eradication of stocks
and disinfection of holding facilities (Whipps et al. 2012), which makes complete
eradication of this ubiquitous species in a facility impractical. Transport of fish into new
12
facilities using only surface disinfected eggs, the “eggs-only policy”, is a cornerstone of
the aquaculture industry (Kent and Kieser 2003). This method, including surface
disinfection with chlorine, is used by most zebrafish labs (Harper and Lawrence 2011).
The “eggs-only” chlorine disinfection method does not prevent transmission of
intraovarial pathogens, such as Mycobacterium spp., or other pathogens outside the egg
shed at spawning that are chlorine resistant (Ferguson et al. 2007). Thus, the best way to
avoid infections would be to use pathogen-free broodstock. However, laboratories usually
exchange fish amongst themselves with no knowledge of the prior disease history
(Lawrence et al. 2012), and still many researchers use zebrafish procured from pet fish
dealers. Moreover, maternally transmitted pathogens could impede studies involving the
development of gnotobiotic zebrafish (Pham et al. 2011).
Antibiotics could be employed under special circumstances for control of
mycobacterial infections in a laboratory setting. Minocycline, a tetracycline-class drug,
was reported in the initial review of M. marinum aquarium-related human skin infections
by Huminer et al. (1986) to be more efficacious than tetracycline. Ciprofloxacin, a
quinolone-class drug, has become the most common quinolone used to treat
mycobacterial infections (Aubry et al. 2002). From clinical isolates, rifampin and
rifabutin are known to be effective against M. marinum (Aubry et al. 2000), and amikacin
and clarithromycin are effective against M. chelonae and other rapid growers (BrownElliott & Wallace, 2002). There are a few reports of successful treatment of
mycobacteriosis in aquaculture. Kanamycin added to water was reported to successfully
treat M. marinum in guppies (Conroy & Conroy 1999) and gouramis (Santacana et al.
13
1982). Kawakami and Kusuda (1990) reported that rifampin, streptomycin, and
erythromycin were effective for reducing mortalities in cultured yellowtail with only two
doses within 24 hours. However, Hedrick et al. (1987) found M. marinum infected striped
bass were refractory to rifampin treatment. The majority of antimycobacterial drug
studies have focused on M. marinum infections and consequently there is no reliable and
standard efficacy data of antimicrobial therapy in treating rapid growers like M. chelonae
in fish.
Susceptibility of M. chelonae isolates from zebrafish could be evaluated using the
broth microdilution method (CLSI 2011). Ten drugs are currently recommended for
testing clinical isolates of rapid growers (CLSI 2011), however Brown-Elliott & Wallace
(2002) report that three of these drugs are less efficacious against M. chelonae
(ciprofloxacin, doxycycline, imipenem). Only two of the seven other drugs have been
used in fish (amikacin and trimethoprim-sulfamethoxazole) with the others completely
untested, as treatment typically focuses on food fish and none of these are currently
approved by the Food and Drug Administration for therapeutic use in food fish (cefoxitin,
clarithromycin, linezolid, moxifloxacin, tobramycin) and must be used off-label (i.e. not
in a manner for which the drug was initially intended to be used). Amikacin, an
aminoglycoside class antimicrobial drug, causes acute subtotal renal tubular necrosis
when administered at repeated therapeutic doses in fish. This can be a transient
phenomenon as most teleost fish retain regenerative neonephrogenesis, including renal
tubules, throughout life (Yasutake and Wales 1983), and the severe renal tubular injury
14
that occurs post-therapeutically with aminoglycoside usage reverses within two to three
weeks after discontinuing chemotherapy (Reimschuessel and Williams 1995).
Mycobacterial Diagnostics in Fish
Mycobacterial culture from clinical specimens suspected of being infected,
usually after an acid-fast stain provides initial evidence of infection, has long been the
gold standard for identification to the species level. Traditionally, specimens have been
inoculated from tissues onto either Lowenstein-Jensen or Middlebrook solid media
(Mediel et al. 2000). Piscine mycobacterial species have an optimal growth temperature
between 23 and 30 oC (Kaattari et al. 2006), as demonstrated by the temperature
differential observed between isolates of Mycobacterium shottsi and Mycobacterium
ulcerans, which preferentially grow at 24 oC and 28 oC, respectively (Tsukamura 1983,
Kaattari et al. 2006). M. marinum grows relatively fast compared with other
mycobacterial species within the M. tuberculosis clade, taking 7 to 10 days for
observable colonies to form on agar plates, versus two to three months for many of the
others (Kaattari et al. 2006).
In the modern diagnostic setting, traditional approaches to diagnosing
mycobacterial infections of zebrafish often are unsatisfactory, due to delays in obtaining
culture results, occasional lack of culture growth, contamination with other bacteria, and
the lack of fresh tissues from which to obtain cultures. To overcome the limitations of
mycobacterial culture and biochemical identification, polymerase chain reaction (PCR)
assays have been developed over the last 10 years, including restriction fragment length
15
polymorphism (RFLP) and Taqman real-time PCR, to expedite diagnosis and specieslevel identification (Colorni et al. 1993, Talaat et al. 1997).
Histology is the primary diagnostic method that is employed for laboratory
zebrafish, but diagnosis of mycobacterial infections by histology only allows for
identification of the bacteria to the genus level. Of the over 150 cases diagnosed by
histology at the Zebrafish International Resource Center (ZIRC), identification of the
bacteria to the species level using culture or molecular methods has been achieved on less
than 20 cases (Watral and Kent 2007, Whipps et al. 2007b, Whipps et al. 2008, Whipps et
al. 2012). Several studies have previously demonstrated that mycobacterial DNA can be
amplified from human and animal (including fish) tissues from paraffin blocks (Ghossein
et al. 1992, Miller et al. 1997, Marchetti et al. 1998, Zink and Nerlich 2004, Pourahmad
et al. 2009a). Efforts to develop more reliable PCR assays that would reduce the time
required for diagnosis as well as increase both the specificity and sensitivity of detecting
mycobacteria in formalin-fixed, paraffin-embedded (FFPE) tissues have been ongoing,
and are mostly focused on human mycobacteriosis (Pao et al. 1988, Pao et al. 1990,
Fiallo et al. 1992, Hardman et al. 1996, Rish et al. 1996, Osaki et al. 1997, Salian et al.
1998, Whittington et al. 1999, Singh et al. 2000, Baba et al. 2008) and to a lesser extent,
mycobacteria infections of animals including fish (Gyimesi et al. 1999, Puttinaowarat et
al. 2002, Pourahmad 2009 a,b).
Results have been mixed with this approach, and time in fixative before
processing into paraffin blocks appears to be an important factor for successfully
obtaining mycobacterial DNA from tissues (Tokuda et al. 1990, Greer et al. 1991).
16
Fixative formulations, especially those containing acids or alcohol in various
concentrations can also influence the ability to retrieve DNA (Eltoum et al. 2001). Most
human and veterinary laboratories use 10% neutral buffered formalin as a fixative, which
is also used for preservation of zebrafish and tissues from other fishes (Ferguson 2006,
Harper and Lawrence 2010). However, zebrafish have been traditionally preserved in
Dietrich’s fixative, a mixture of fixatives that contains both acetic acid and 95% alcohol
in addition to formaldehyde.
The ZIRC zebrafish diagnostic program is largely based on histologic evaluation
of whole fish specimens; however, there is a pressing need to identify mycobacterial
infections to the species level in order to allow for more informed management of these
infections. In Chapter 2, I investigate the development of a PCR test for retrieval of
mycobacteria DNA and evaluate the influence of time and fixative (either 10% neutral
buffered formalin or Dietrich’s) on the ability to recover DNA for PCR assays. This was
achieved by experimentally infecting zebrafish with either M. chelonae or M. marinum,
and preserving the infected fish in both fixatives for various time points up to 45 days.
Piscine Mycobacteriosis and Human Health Concerns
All the mycobacteria species isolated from zebrafish to date have been reported to
be infective for humans, and infected zebrafish may pose a health risk to researchers,
technical staff and other workers directly handling the fish or inadvertently exposed to
water and equipment contaminated with mycobacteria. Mycobacterium marinum human
infections have been documented to be directly related to the handling aquarium fishes
(Clifton and Hatwick 2011). Nationwide, zebrafish research facilities employ hundreds of
17
undergraduate and graduate students, researchers, technicians and husbandry staff, thus
the exposed human population is significant. Indeed, two technicians at a large zebrafish
laboratory that experienced an M. marinum outbreak became infected, resulting in
chronic hand lesions. Human infections were directly linked with infected fish at an
aquaculture facility using DNA fingerprinting (Ostland et al. 2008). These infections
result in chronic ulcerative lesions on the extremities, as has been classically described,
and on rare occasion results in disseminated disease in immune compromised humans
(Fabricius et al. 2009). Infections are seldom life threatening, but require prolonged
antimicrobial therapy lasting several months. Therefore, the impact of subclinical
mycobacteria infections in zebrafish should be considered a significant human health
concern for those working with the fish.
Mycobacteriosis in Fishes Used in Biomedical Research
Naturally-occurring mycobacterial infections have been reported in a wide variety
of fishes used in biomedical research, including the two most prominently used species,
zebrafish and Japanese medaka. M. marinum is most frequently identified as the causal
agent in these fish, but several other mycobacterial species have been isolated from
infected fish such as M. chelonae and M. haemophilum. This has led to the use of fish as
suitable models of mycobacteriosis. Historically, animal models to study various aspects
of Mycobacterium tuberculosis human infection were first developed more than 60 years
ago and have traditionally involved rabbits, guinea pigs, mice and less often non-human
primates, with each model system except non-human primates failing in some respects to
faithfully reproduce all features of the human disease. The rabbit and guinea pig models
18
approach functional utility using M. tuberculosis or M. bovis, because of their close
approximation of human infectivity and pathological changes including virulence,
granuloma and giant cell formation. The mouse model appears unsuitable due to the
inherent resistance of mice to M. tuberculosis or M. bovis infection and the almost
complete lack of comparable pathology to the human disease (Sakamoto 2012).
Paradoxically, despite these drawbacks, mice have remained the most popular animal
model to study human tuberculosis. All three common mammalian animal models of
human tuberculosis are less than ideal when their high costs, requirement for laboratory
personnel experienced with these animals and substantial generation of biologically
hazardous waste are considered (Turner et al. 2003).
Over 28,900 species of fish have been identified and cataloged that represent
approximately 96% of all known vertebrate species (www.fishbase.org). Among these,
there are 13,000 fresh water species (Leveque et al. 2008). Freshwater fishes comprise
the majority of fish used in biomedical research, but only a relative handful have been
employed as model organisms. Paradigmatic laboratory fishes that have been used as
comparative model organisms encompass a broad “phylogenic cascade” of teleost fishes
including salmonids, catfish, medaka, poecilids, killifish, stickleback, cichlids,
scombrids, silversides, pufferfish, fathead and sheepshead minnows, goldfish and
zebrafish. Many of these fish were initially used in for investigations into physiology,
immunology, melanin formation and function, carcinogenicity testing and aquatic
toxicology (Wolke 1984).
19
Among the teleost fishes used in biomedical research, the zebrafish has emerged as the
predominant model organism owing to the canonical and highly tractable instrumental
traits such as small size, high fecundity, robust tolerance of variable environmental
conditions and well-conserved comparative synteny with the human genome, despite 220
million years of evolutionary divergence and genomic polyploidy/re-duplication
(Streisinger et al. 1981, Spence et al. 2008, Meunier 2012). Originally described as a
laboratory fish for developmental studies by Creaser (1934), the zebrafish gained
prominence in 1970 for its role in the development of artificial parthenogenesis and
subsequent mutation mapping (Streisinger et al. 1981) and subsequently in 1996, when
large-scale ethyl nitrosourea mutagenesis screens conducted independently by NussleinVollhard in Tubingen, Germany and Fishman at Harvard generated hundreds of zebrafish
mutant phenotypes (Driever and Fishman 1996, Haffter et al. 1996, Stainier et al. 1996).
In the ensuing 43 years since the introduction of the zebrafish as a lab animal and model
organism, it has been widely employed in the genomic and postgenomic eras for
comparative studies of developmental biology, toxicology, infectious diseases,
hemodynamic disorders, aging and cancer (Dooley and Zon 2000, Amatruda et al. 2002,
North and Zon 2003, Amsterdam et al. 2004, Berghmans et al. 2005, Grabher and Look
2006, Beckman 2007, Gerhard 2007, Kari et al. 2007, Amatruda and Patton 2008, Mione
and Trede 2010, Bourque and Houvras 2011). Additionally, zebrafish offer many
comparative advantages over other animal model systems, which include similarities in
organ histology and cell-signaling pathways, imaging of live animals (i.e. optical
20
transparency), rapid genetic and drug screens at the organism level (Prendergast 2008)
and a completed genome sequence (http://www.sanger.ac.uk).
Models of Mycobacteriosis in Laboratory Fishes
The rise of laboratory fish as a model organism for studying human
mycobacteriosis can be traced back to 1963, when fish and other poikilothermic animals
were identified as being susceptible to M. marinum infection. Exploration of developing
piscine models of mycobacterial infection became desirable because these models
fulfilled key criteria that had significant advantages over the more conventional
mammalian models such as lower costs, ease of husbandry and natural susceptibility to a
mycobacterial pathogen, M. marinum (Wolke 1984, Trucksis 2000). Eventually, some 33
years later, the swordtail (Xiphophorus helleri) was notably used as a potential model for
immunohistochemical detection of Mycobacterium spp., conferring the benefit of
visualizing previously undetectable mycobacteria in host tissues other than granulomas
(Gomez et al. 1996).
Goldfish (Carassius auratus) emerged two years later to become a model for
mycobacteria research that is still in use today, emphasizing host-pathogen interaction
and pathogenesis, with initial model development by Talaat et al. (1998). This model
produced many firsts, including establishing the definitive histomorphologic pathology of
M. marinum infection in a laboratory fish, by fully characterizing the unique features of
goldfish granulomas and inflammatory response, dose-dependent differences between
acute and chronic progressive mycobacterial infection and determination of the LD50 for
M. marinum infection in any animal model (Talaat et al. 1998). The nascent goldfish-M.
21
marinum model was promoted as an ideal system in which to identify and characterize
virulence genes of various mycobacterial species, with focus on M. marinum because of
its close homology with M. tuberculosis (Trucksis 2000). In subsequent work, the
goldfish model was more fully established and pathogenicity studies of M. fortuitum and
M. smegmatis revealed that these rapid-growing mycobacteria produce productive and
persistent infections in goldfish, and documented for the first time that M. smegmatis
was, in fact, more pathogenic to fish than M. fortuitum (Talaat et al. 1998). They were
also able to make a persuasive argument against the previously developed leopard frog
model of mycobacteriosis (Ramakrishnan et al. 1997), as even a high inoculum (108 cfu
of M. smegmatis) did not produce persistent infection and mycobacteria was not
recovered from infected tissues, even after 28 days. More recently, the goldfish-M.
marinum infection model has been exploited to investigate host-pathogen interactions,
demonstrating the antimycobacterial responses of goldfish monocyte and macrophages
and showing that the gene expression of pro-inflammatory cytokines and cytokine
receptors becomes elevated as part of the immune response to infection (Grayfer et al.
2011, Hodgkinson et al. 2012).
Japanese medaka (Oryzias latipes) enjoyed a long history (over 50 years) as a
laboratory fish prior to their use as a model for mycobacteriosis. Indeed, there were
several undocumented accounts of possible natural mycobacterial infections occurring in
medaka (Abner et al. 1994, Sanders and Swaim 2001, Broussard and Ennis 2007).
Following on the success of the goldfish-M. marinum model, medaka (Oryzias latipes)
were utilized in a similar fashion by Broussard and Ennis (2007), as they sought to create
22
a fish model of tuberculosis that would be a more effective surrogate for human TB, by
allowing the disease progression to be studied in a transparent fish that permitted the use
of GFP labeled M. marinum. They developed an infection model that closely paralleled
the goldfish model, characterizing mycobacteriosis in the medaka and demonstrating both
acute and chronic infections that were dose-dependent. In addition, they were able to
track infections and monitor progression as well as colonization in real-time within a
living fish (Broussard and Ennis 2007).
During the development of the medaka model, Broussard and Ennis became the
first researchers to use zebrafish for direct comparison to the Japanese medaka, and
discovered that M. marinum infections were more severe in zebrafish. They ultimately
concluded that medaka may be more suitable for chronic mycobacteria infection studies.
In further studies using medaka, the Ennis group was able to show that chronic M.
marinum infections and hence, the chronic inflammation accompanying these infections
may potentiate carcinogenicity in medaka colonies. They also showed that a GFP (green
fluorescent protein) reporter plasmid construct in engineered Mycobacterium marinum
may reduce the native virulence of the bacteria in the fish host, through an unresolved
mechanism relating to GFP over-expression (Broussard et al. 2009, Mutoji and Ennis
2012).
Departing from the more traditional M. marinum model, Mosi et al. (2012) began
using Mycobacterium ulcerans, the causative agent of Buruli ulcer in humans, to test
pathogenicity of the bacteria in medaka. This was the first attempt to model M. ulcerans
infection in a fish species and curiously, they could not reproduce the disease in medaka
23
even with a high inoculating dose (108 CFU) and none of the infected medaka showed
any signs of disease. However, in a concurrent study they were able to demonstrate that
the M. marinum DL strain (myolactone-producing) causes acute mortality in medaka and
is toxic to goldfish macrophages (Mosi et al. 2012).
Primary work to develop the zebrafish-M. marinum model that set the stage for
other researchers in the zebrafish infectious disease community was conducted by Prouty
et al. (2003), when they created a M. marinum pathogenesis model that successfully
recapitulated several aspects of human tuberculosis, including definitive granuloma
formation and bimodal (acute versus chronic) disease based upon inoculum dose. This
initial model was further capitalized upon by Swaim et al. (2006) when they created the
rag1 zebrafish mutant to examine the adaptive immune response to mycobacterial
infection, proving that in fish as well as humans, the outcome of mycobacterial infection
was dependent on adaptive immunity. This study, in addition to demonstrating that adult
zebrafish could mount an analogous immune response to mycobacteria, provided more
evidence that zebrafish produce lesions, particularly caseous granulomas, consistent with
human mycobacteriosis (Swaim et al. 2006).
While efficient models of M. marinum infection were being worked out in adult
zebrafish, including the innate and adaptive immune response, Lesley and Ramakrishnan
(2008) began to focus on the earliest events that occur in mycobacterial pathogenesis,
using larval zebrafish. They discovered, using an in vivo model made possible by larval
zebrafish transparency, that 30 hours post-fertilization (hpf) larval zebrafish macrophages
home to and phagocytose individual and aggregated mycobacteria at sites of infection,
24
with subsequent systemic migration and continued growth of mycobacteria within
macrophages. They were then able to show that these infected macrophages aggregate
into nascent granulomas and these granulomas undergo maintenance relying on the
mycobacterial ESX-1 virulence determinant before maturation (Lesley and Ramakrishnan
2008). The zebrafish-M. marinum model is currently being employed for high-throughput
antimycobacterial drug screens (Takaki et al. 2012) and vaccine development (Cui et al.
2010).
Mycobacteriosis in Laboratory Zebrafish
The occurrence of mycobacteriosis in laboratory fishes is a serious concern for
various reasons: 1.) Acute disease in a colony setting can cause devastating losses and
destroy months or years of research, 2.) Non-protocol induced variation, particularly in
studies using zebrafish to investigate mycobacterial disease, can confound results and
lead to erroneous interpretation of data, 3.) Mycobacterial disease or subclinical
infections in laboratory fish pose a concern to human health and to staff working with the
fish. Mycobacterium chelonae is the most common species found in laboratory zebrafish,
and often presents as a subclinical infection (Watral and Kent 2007; Whipps et al. 2008;
Whipps et al. 2012). Mycobacterium haemophilum is more pathogenic and infections are
associated with high mortalities and severe infections (Whipps et al. 2007b). In contrast,
Mycobacterium marinum is not frequently seen in zebrafish, but when it occurs it has
been associated with acute to chronic infections and also high mortality. Importantly, we
(the Kent lab) have verified outbreaks of M. marinum in two separate, large-scale
(commercial) zebrafish facilities.
25
The mycobacterial species infecting laboratory zebrafish, including M. marinum,
can be commonly isolated from the local environment especially water sources, pipes and
tanks (Primm et al. 2004). Persistence and replication of mycobacteria in the aquatic
environment is aided by the bacteria’s inherent resistance to chlorine, survival in lownutrient environments, (starvation resistance), thermoresistance and ability to form
biofilms (Nyka 1974, Schulze-Robbecke and Buchholtz 1992, Schulze-Robbecke et al.
1992, Hall-Stoodley and Lappin-Scott 1998, Taylor et al. 2000, Le Dantec et al. 2002,
September et al. 2004). These capabilities enable mycobacteria to establish populations
and persist in flowing water systems, like those found in the laboratory setting, despite
relatively slow growth.
Similar to other laboratory animal models, underlying mycobacterial infections
can compromise research. In addition to causing acute diseases, chronic subclinical
diseases are of concern as they induce non-protocol induced variation in experiments
(Baker 2003, Kent et al. 2011, Kent et al. 2012, Lawrence et al. 2012). Chronic disease
caused by Mycobacterium spp. is the second most common infectious disease in
zebrafish research colonies (http://zebrafish.org/zirc/health/diseaseManual.php) after the
microsporidium Pseudoloma neurophilia. Historically, in many instances mycobacterial
infections in laboratory fish, including zebrafish, have been misinterpreted or missed
entirely. As late as 1959, fish health professionals working in Germany claimed to have
discovered a “kidney tumor virus” (KTV) affecting zebrafish (Danio rerio), which
produced abdominal, skin, retrobulbar and renal masses that preferentially involved the
kidney (Wessing 1959, Wessing and Bargen 1959). A separate study by Beckwith and
26
Malsberger (1980) paralleling the work of Wessing, using a variety of common aquarium
fish (Danio rerio, Danio albolineatus, Devario aequipinnatus and Poecilia reticulata)
obtained from a local pet store, was performed using tissue homogenate extracts of the
abdominal masses from the aquarium fish. The extracts did eventually cause similar
“tumors” in intracoelomically inoculated fish after 4 months, but did not produce any
cytopathogenic effects in cell culture, as would be expected with a viral pathogen.
Ultimately, it was revealed by histology and bacterial culture that the tumor “masses”
observed by the previous investigators were, in fact, tubercles containing acid-fast bacilli
resulting from Mycobacterium fortuitum infection. Beckwith and Malsberger postulated
that cell fractions of the extract, notably cord factor (6-6’ trehalose mycolic acid) could
have elicited tubercle (i.e. granuloma) formation in the experimentally exposed fish in the
absence of live bacteria, although it is more likely that they were using a contaminated
filtrate.
Mycobacterium spp. infections were found in 43% of zebrafish research facilities
submitting cases to the ZIRC diagnostic laboratory
(http://zebrafish.org/zirc/health/diseaseManual). Review of the ZIRC pathology data base
revealed approximately 150 cases, comprising over 1,000 fish, of mycobacteriosis
diagnosed by histology. Infections were identified to the species level in about 20 of
these cases when combining culture and molecular methods (Kent et al. 2004, Whipps et
al. 2007b, Whipps et al. 2008, Whipps et al. 2012). With the rare exception of severe
mycobacteriosis outbreaks caused by M. marinum or M. haemophilum (Whipps et al.
2007b), these diseases are chronic in nature and have caused low level mortalities.
27
Nevertheless, in some of these cases, sublethal lesions and infections showed high
prevalence in populations, even when mortalities were within acceptable limits. The
occurrence of underlying diseases causing low mortalities may be acceptable for
production aquaculture or with pet fish, but (as with any laboratory animal) should be
avoided in fish used in research.
Over the last 10 years important diseases in zebrafish have been described, modes
of transmission documented (Kent and Bishop-Stewart 2003, Harriff et al. 2007),
strategies provided to avoid these infections in zebrafish research settings (Kent et al.
2009, Sanders et al. 2012, Whipps et al. 2012), diagnostic tests developed (Whipps and
Kent 2008, Peterson et al. 2011, Sanders and Kent 2011) and there has been development
of an SPF laboratory that is now providing Pseudoloma-free fish to research community
(Kent et al. 2011). Hence, tools are now available to prevent the two most common
diseases, microsporidiosis and mycobacteriosis (except for M. chelonae).
In contrast to researchers using rodent models, many zebrafish researchers have
been reluctant to adopt strategies to avoid these infections. This is likely because they
have not been convinced that the level of impact on their research caused by using fish
with underlying diseases warrants the efforts that would be involved in avoiding such
infections. For example, the ZIRC diagnostic services conducts disease evaluations at
only about 5% of the zebrafish facilities listed in the Zebrafish Model Organism Database
(zfin.org, Lawrence et al. 2012). Some of these labs are submitting fish to other
diagnostic labs, but the vast majority are not documenting pathogens or diseases in their
fish. Perhaps this is because principle investigators working with zebrafish do not
28
appreciate the impacts that mild or subclinical disease could have on their research. It is,
therefore, important to further elucidate the influences of these infections on endpoints
beyond histology. In addition to reports on pathological changes associated with
mycobacteriosis in zebrafish, significant changes in the transcriptome of zebrafish
experimentally infected with M. marinum have been documented (Meijer et al. 2005,
Hegedus et al. 2009). Clearly, significant inflammatory changes throughout the visceral
organs, as seen with mycobacteriosis, or in the central nervous system or muscle with
Pseudoloma would likely influence a variety of experimental end points, including the
secretion of inflammatory cytokines that will greatly alter the microenvironment and
tissues where infections occur. Tumor necrosis factor alpha (TNFα), nuclear factor kappa
B (NFκB) and other inflammatory mediators are key regulators of normal development
and homeostasis of many tissues in vertebrates and invertebrates (Boersma and Meffert,
2008; Doherty 2007, Hernandez-Gutierrez et al. 2006), and their dysregulated expression
as a result of chronic infections will have far reaching consequences on experimental
endpoints. For instance, TNFα is also a potent endogenous mutagen acting by production
of reactive oxygen radicals (Yan et al. 2006) and its secretion in infected animals will
alter the growth phenotype of implanted tumors.
Chronic inflammation of the ovary, leading to the condition known as eggassociated inflammation and fibroplasia (EAIF), is a common lesion in zebrafish with
long-standing mycobacterial oophoritis (Kent et al. 2004). Important endpoints for
developmental biology research, such as fecundity, are negatively impacted by this
chronic disease.
29
Therefore, avoidance of disease in zebrafish continues to increase in importance
as the use of zebrafish models has expanded to areas of research that require experimental
animals to live in a healthy state for long periods. Pathogens and infectious diseases are
actually common in zebrafish facilities, even though they are reared in a laboratory
setting. Certainly acute diseases involving high mortality can impact research,
particularly when valuable lines or mutants are lost. Based on over a decade of data
obtained from the ZIRC diagnostic service, subclinical infections are by far the most
commonly diagnosed histopathological changes observed in zebrafish, often through
sentinel fish surveys (http://zebrafish.org/zirc/health/index.php). Although often not
associated with obvious morbidity, these infections should still be avoided as well as they
lead to several documented or potential impacts in research (Kent et al. 2012).
Summary
The objectives of the research comprising my thesis are to address and perhaps
resolve two current areas of need in the laboratory zebrafish community, those of
diagnostics, specifically the ability to identify mycobacterial pathogens from paraffinembedded tissue sections (Chapter 2) and to elucidate how mycobacterial pathogens are
transmitted among zebrafish within a colony setting (Chapters 3 and 4). One of the
remaining diagnostic hurdles to understanding how mycobacteriosis affects laboratory
zebrafish has been the inability to reliably culture and identify mycobacteria to the
species level from clinical cases, and to perform retrospective epidemiologic studies from
previous case material. The singular reason for these issues is that the majority of
zebrafish diagnostic cases are sent to the ZIRC for evaluation, and this facility does not
30
allow live or freshly dead zebrafish to be shipped there for biosecurity reasons. Thus, all
screening and identification of potential cases of mycobacteriosis rely on zebrafish that
have been placed into fixative solutions (either Dietrich’s or 10% neutral buffered
formalin) and are subsequently processed and paraffin-embedded for histology.
Similarly, another problematic area is our understanding of how mycobacteria are
transmitted among zebrafish and what vectors might serve as a vehicle for transmission,
which has important ramifications in the control and elimination of mycobacterial
pathogens from established zebrafish colonies. The mechanisms of mycobacterial
transmission in zebrafish, especially the identification of a primary vector (Paramecium
caudatum) and natural oro-enteric route of transmission, which I have significantly
contributed to in this thesis, will allow more reliable transmission studies to be developed
for future use as a potential natural model for early transmission (larval zebrafish) and in
studies using adult zebrafish.
Additionally, I have contributed two papers that make significant progress in
zebrafish diagnostics, including the discovery and refinement of the Luna stain for
detection of microsporidian parasites, specifically Pseudoloma neurophilia, in tissue
sections and the discovery of a novel brain tumor in zebrafish, which has particular
relevance because zebrafish are frequently used in neurodevelopmental studies
(Appendix).
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46
Chapter 2
Comparison of Fixatives, Fixation Time, and Severity of Infection on PCR
Amplification and Detection of Mycobacterium marinum and Mycobacterium
chelonae DNA in Paraffin-Embedded Zebrafish (Danio rerio)
Tracy S. Peterson1, Michael L. Kent1,2, Jayde A. Ferguson4, Virginia G. Watral1 and
Christopher M. Whipps3
Diseases of Aquatic Organisms, 2013. In Press
1
Department of Microbiology. 220 Nash Hall, Oregon State University, Corvallis, OR
97331-3404
2
Department of Biomedical Sciences, Oregon State University, Corvallis, OR 97331
3
Department of Environmental Science and Forestry, State University of New York
College of Environmental Science and Forestry, SUNY-ESF, 1 Forestry Drive, Syracuse,
NY 13210
4
Alaska Department of Fish and Game, Commercial Fisheries Division, Pathology
Laboratory, Anchorage, AK 99518
47
ABSTRACT
Mycobacteriosis is a common disease of laboratory zebrafish (Danio rerio).
Different infection patterns occur in zebrafish depending on mycobacterial species.
Mycobacterium marinum and M. haemophilum produce virulent infections associated
with high mortality, whereas M. chelonae is more wide spread and not associated with
high mortality. Identification of mycobacterial infections to the species level provides
important information for making management decisions. Observation of acid-fast
bacilli in histological sections or tissue imprints is the most common diagnostic method
for mycobacteriosis in fish, but only allows for diagnosis to the genus level.
Mycobacterial culture, followed by molecular or biochemical identification is the
traditional approach, but recently it has been shown that DNA of diagnostic value can be
retrieved from paraffin blocks. Type of fixative, time in fixative before processing,
species of mycobacteria, and severity of infection were investigated as parameters to
determine if the hsp gene PCR assay (primer set HS5F/hsp667R) could detect and
amplify mycobacterial DNA from paraffin-embedded zebrafish. Whole zebrafish were
experimentally infected with either M. chelonae or M. marinum, and then preserved in
10% neutral buffered formalin or Dietrich’s fixative for 3, 7, 21 and 45 days.
Subsequently, fish were evaluated by H&E and Fite’s acid-fast stains to detect
mycobacteria within granulomatous lesions. The PCR assay was quite effective, and
obtained PCR product from 75% and 88% of the M. chelonae and M. marinum infected
fish, respectively. Fixative type, time in fixative, and mycobacterial species showed no
statistical relationship with the efficacy of the PCR test.
48
INTRODUCTION
Piscine mycobacteriosis is a well-characterized disease, with several
mycobacterial species infecting multiple genera and species of fish (Decostere et al.
2004, Lewis and Chinabut 2011). Zebrafish (Danio rerio) are now a widely used
vertebrate animal model in biomedical research (Lele and Krone 1996, Dooley and Zon
2000, Grunwald and Eisen 2002, Rubinstein 2003, Alestrom et al. 2006, Cosma et al.
2006, Allen and Neely 2010) with an ever-increasing number of academic and private
laboratories using zebrafish. Since the emergence of the zebrafish as a pre-eminent
animal model, there has been a corresponding interest in diseases that may occur in this
species within a laboratory setting. The Zebrafish International Resource Center (ZIRC)
in Eugene, Oregon (http://zebrafish.org/zirc/home/guide.php) has provided a diagnostic
service to the zebrafish research community since 2000. Mycobacteriosis, based on
histological observation of acid-fast bacteria within typical tuberculoid granulomas
(Swaim et al. 2006), has been detected at 41% of submitting facilities in over 150 cases
involving greater than 500 fish (http://zebrafish.org/zirc/health/diseaseManual.php).
Several species of Mycobacterium have been reported to cause disease in zebrafish,
including M. chelonae (Astrofsky et al. 2000, Kent et al. 2004, Murray et al. 2011), M.
peregrinum (Kent et al. 2004), M. haemophilum (Whipps et al. 2007, 2012), and M.
marinum (Ostland et al. 2007). Based on observations reported in diagnostic cases
(Astrofsky et al 2000, Kent et al. 2004, Whipps et al. 2008, Murray et al. 2011, Whipps et
al. 2012) and laboratory transmission studies (Watral and Kent 2007, Whipps et al. 2007,
Ostlander al. 2008), the severity of mycobacteriosis is usually related to the
49
Mycobacterium species causing the infection and in some cases can be correlated with
husbandry stress (Ramsay et al. 2009). Mycobacterium chelonae is relatively widespread
and causes chronic infections but minimal mortalities (Whipps et al. 2008, Murray et al.
2011). There have been six outbreaks of M. haemophilum, which caused severe
infections with high but chronic mortality (Whipps et al. 2007). Mycobacterium marinum
is uncommonly isolated from cases of mycobacteriosis in zebrafish, but when infection
occurs it is associated with acute disease and high mortalities (Watral and Kent 2007,
Ostland et al. 2008). Histology is the primary diagnostic method that we use with
zebrafish, but diagnosis of mycobacterial infections by histology only allows for
identification of the bacteria to the genus level. Of the over 150 cases diagnosed by
histology by ZIRC, identification of the bacteria to the species level using culture or
molecular methods has been achieved on less than 20 cases (Watral and Kent 2007,
Whipps et al. 2007, 2008, 2012). Several studies have previously demonstrated that
mycobacterial DNA can be amplified from human and animal (including fish) tissues
from paraffin blocks (Ghossein et al. 1992, Miller et al. 1997, Marchetti et al. 1998, Zink
and Nerlich 2004, Pourahmad et al. 2009a). Efforts to develop PCR assays that would
reduce the time required for diagnosis as well as increase both the specificity and
sensitivity of detecting mycobacteria in formalin-fixed, paraffin-embedded (FFPE)
tissues have been ongoing, and are mostly focused on human mycobacteriosis (Pao et al.
1988, Pao et al. 1990, Fiallo et al. 1992, Hardman et al. 1996, Rish et al. 1996, Osaki et
al. 1997, Salian et al. 1998, Whittington et al. 1999, Singh et al. 2000, Baba et al. 2008)
and to a lesser extent, mycobacteria infections of animals including fish (Gyimesi et al.
50
1999, Puttinaowarat et al. 2002, Pourahmad 2009 a,b). Results have been mixed with this
approach, and time in fixative before processing into paraffin blocks appears to be an
important factor for successfully obtaining mycobacterial DNA from tissues (Tokuda et
al. 1990, Greer et al. 1991). Fixative formulations, especially those containing acids or
alcohol at various concentrations can also influence the ability to retrieve DNA (Eltoum
et al. 2001). Most human and veterinary laboratories use 10% neutral buffered formalin
as a fixative, which is also used for preservation of zebrafish and tissues from other fishes
(Ferguson 2006, Harper and Lawrence 2011). However, zebrafish have been
traditionally preserved in Dietrich’s fixative, a mixture of chemicals that contains acetic
acid and 95% alcohol in addition to formaldehyde. The ZIRC zebrafish diagnostic
program is largely based on histologic evaluation of whole fish specimens; however,
there is a need to identify mycobacterial infections to the species level in order to allow
for informed management of these infections. Subsequently, a PCR test was developed
for retrieval of mycobacteria DNA and evaluated the influence of time and fixative
(either 10% neutral buffered formalin or Dietrich’s) on the ability to recover DNA for
PCR assays. This was achieved by experimentally infecting zebrafish with either M.
chelonae or M. marinum, and preserving the infected fish in both fixatives for various
time points up to 45 days.
MATERIALS AND METHODS
Mycobacteria cultures used for inoculation were prepared in the following
manner. Stock cultures from the Kent laboratory reference collection of M. marinum
OSU 214 (M. marinum) and M. chelonae H1E2 (M. chelonae) were incubated on
51
Columbia CNA (colistin and nalidixic acid) 5% sheep blood and Middlebrook 7H10 agar
(Remel, Lenexa, KS) for 14 days at a temperature of 28 °C. Fresh culture material from
agar plates was loop inoculated into sterile PBS and washed three times to eliminate
carry-over of antimicrobial chemicals from the agar plates and then adjusted in sterile
PBS to an optical density of 1.0 based on the MacFarlane scale, yielding an approximate
dose of 1 x 106 bacteria/25 µl for the final injection amount.
AB strain adult zebrafish of mixed sex were obtained from the Sinnhuber Aquatic
Research Laboratory at Oregon State University as experimental animals. Fish were
housed in a biosafety-level 2 (ABSL-2) laboratory and maintained for twelve weeks after
inoculation, allowing for progression of infection, in triplicate flow-through tanks at 2527 °C with supplemental aeration and a 14/10 light/dark photoperiod schedule. For
inoculation with M. marinum and M. chelonae, fish were anesthetized with 100 mg/l-1
tricaine methanosulfonate (MS-222) then aseptically given intraperitoneal injections in
the right flank, to avoid accidentally lancing the spleen, with 25 µl of either sterile PBS
(sham-inoculated control group) or the prepared M. marinum and M. chelonae inocula.
At the end of 12 weeks, moribund fish and all surviving fish were euthanized by overdose
of MS-222, the operculae removed and fish incised along the ventrum in order to expose
viscera to the fixative solutions, then immediately placed into 15 ml of either Dietrich’s
fixative or 10% neutral buffered formalin (10% NBF). Fish were held in Dietrich’s
fixative or 10% NBF for 3, 7, 21 and 45 days (Table 2.1).
52
Histology and PCR
Decalcification was performed on the fixed zebrafish samples as follows: Fish
preserved in Dietrich’s fixative were decalcified in 5% trichloroacetic acid (TCA)
overnight (following the Zebrafish International Resource Center protocol), while fish
preserved in 10% NBF were placed in CalExII (Fisher Scientific, Fair Lawn, New Jersey)
for 48 h. After decalcification and rinsing for 30 min with de-ionized water, all fish were
transferred into 70% ethanol and held no longer than 48 h before processing. Following
fixation, zebrafish were processed for paraffin embedding by routine procedures.
Sections examined for histology were cut at 5 µm and stained with hematoxylin and
eosin as well as Fite’s acid fast stain for detection of acid-fast bacilli (Luna 1968).
PCR scrolls were made from dry tissue sections cut from blocks at 5 µm thickness after
decontamination and prepared for DNA extraction in the following manner. Prior to
sectioning the scrolls, each tissue block was wiped down with one DNA AWAYTM wipe
(Molecular BioProducts, Inc. San Diego, CA), then faced with a new microtome blade.
The block face was wiped with a fresh DNA AWAYTM wipe, the microtome blade
changed and twelve 5 µm sections cut in a scroll. Two scrolls, handled with clean
wooden toothpicks, were placed into a sterile, pre-labeled 1.5 mL microfuge tube.
Microtome blades were discarded after cutting scrolls for each tissue block, in order to
avoid cross-contamination. The entire microtome was then wiped down with xylene (to
remove excess paraffin waste) and 100% ethanol, allowed to air dry then wiped down
with a DNA AWAYTM wipe and air-dried. Gloves were changed between microtome
cleaning, blade changes and tissue block handling. After every five blocks, scrolls were
53
obtained from a block containing a negative zebrafish from the Sinnhuber Aquatic
Research Laboratory SPF colony. All tissue blocks and PCR scrolls were prepared in an
identical manner.
Paraffin-embedded tissue sections were washed twice with 1.2 ml of xylene and
then twice with the same volume of 100% ethanol, which followed the recommended
protocol for the Qiagen DNeasy Blood & Tissue Kit (Qiagen Inc., Valencia, California).
The tissue pellet was suspended in 375 µl of ATL buffer and Antifoam A (SigmaAldrich) was added to a final concentration of 1%. This was transferred to a 0.5ml screw
cap tube containing 300µl of 0.1mm Zirconia/Silica Beads (BioSpec Products
Bartlesville, OK) and placed on a Mini-BeadBeater 16 (BioSpec Products) for 3 minutes.
The bead beating step was incorporated as preliminary studies showed that this yielded
more positive results. Thirty µl of 20 mg/ml Proteinase K was added to this disrupted
tissue and digested overnight in a 45 °C water bath. The sample was centrifuged, then
200µl removed and DNA extracted following the manufacturer’s instructions. PCR was
carried out with primers HS5F (GTC ATC ACC GTC GAG GAG) and hsp667R
(Selvaraju et al. 2005), yielding a product 156 bp of hsp65 sequence. Amplifications were
performed on a C1000™ Thermal Cycler (BioRad Laboratories, Hercules, CA) with
initial denaturation at 95° C for 3 min, followed by 35 cycles of 94° C for 30s, 54° C for
45s, 68° C for 60s, and a final extension at 68° C for 7 min. Product amplification was
evaluated by observation on a 2% agarose gel. To confirm the specificity of the PCR
reactions, products from a subset of three positives each of M. chelonae and M. marinum
samples were sequenced. Amplification products were purified using the E.Z.N.A. Cycle
54
Pure Kit (Omega Bio-Tek, Norcross, GA) and direct sequencing was performed using
primer HS5F on the ABI BigDye Terminator Cycle Sequencing Ready Reaction Kit v3.1,
using the ABI3730xl Genetic Analyzer (Applied Biosystems, Foster City, CA).
Severity of Infection
Individual infected fish were scored as light or heavy infections by enumerating
granulomas containing acid-fast bacteria in two whole body sagittal sections of fish per
slide. For both M. marinum and M.chelonae, light and heavy infections were classified
as 1-6 granulomas and greater than 6 granulomas, respectively.
Statistics
All statistical analyses were conducted with the program R, version 2.7.2 (R
Development Core 315 Team). Significance was set at p ≤ 0.05 and p-values are 2-tailed.
Differences in PCR prevalence between Mycobacterium species, fixative types and
fixation duration was tested with the Fisher’s exact test. A generalized linear regression
model was used to evaluate the influence of fish sex, Mycobacterium species, fixative
type, fixation time and number of granulomas (indicating level of infection) detected by
histology had on the ability of the PCR (primer set HS5F/hsp667R) to successfully detect
Mycobacterium DNA. Logistic regression was the most appropriate technique to evaluate
these effects because the response of PCR detection is a binary variable. This model had
a logit link function and an interaction effect of fixative and fixation time was explored.
The likelihood ratio test was used to determine how well the model fit the data compared
to the null model of just the intercept. Individual variables and combinations thereof were
also tested using a backwards elimination technique.
55
RESULTS
DNA was successfully recovered from a large number of the histology-positive
samples, irrespective of total fixation time, type of fixative, or Mycobacterium spp.
(Table 2.1). There was no significant interaction effect of fixative and fixation time (p >
0.99) so this variable was removed and the resulting model that contained the variables of
interest did not fit the data as well as the null model (p = 0.37; chi-square of 7.5 with 7
df), meaning that none of these variables explained the variation in the detection of
mycobacteria DNA by PCR. Similar results were obtained by evaluating individual
variables and combinations thereof. Fixation time of 45 days showed a trend towards
fewer PCR positives, and one of the four groups at this time period (M. chelonae in 10%
neutral buffered formalin) had only one of five positive samples. Although this time in
fixative was not a statistically significant influence, it was suggestive of having an effect
(p = 0.08). None of the histology negative control fish that were included throughout the
evaluations were positive by PCR. Sequencing of three M. marinum and three M.
chelonae PCR positive samples confirmed their identity.
Severity of infection was also evaluated as an explanatory variable. Mortality
occurred in the M. marinum group, and overall these fish exhibited about three times the
number of granulomas containing acid fast bacteria than the M. chelonae-infected fish
(29.6 versus 9.4 granulomas/fish, respectively) that were positive by PCR (Table 2.1).
The M. chelonae groups yielded fewer positive fish by PCR, but there was no significant
relationship with PCR compared to either Mycobacterium species or number of
granulomas/fish. There was also no significant difference between fixative types.
56
DISCUSSION
Evaluation of DNA sequences has become a cornerstone in bacterial species
identification. This is certainly the case for Mycobacterium species, as they are often
fastidious, grow slowly and typically provide few culture-specific or biochemical traits
by conventional methods that facilitate species-level identification (Daniel 1990, Cousins
et al. 1992). It is often more useful to obtain sequences for diagnoses directly from
infected tissues, particularly for Mycobacterium species that are slow growing or require
specialized media. This approach has been used to diagnose mycobacteria infections
directly from frozen or fresh infected fish (Kaattari et al. 2005, Poort et al. 2006, Whipps
et al. 2003, Whipps et al. 2007). Often only formalin preserved tissues embedded in
paraffin are available, which has led to attempts to obtain mycobacterial DNA from
paraffin-embedded tissues following observation of either chronic lesions (i.e.,
granulomas) or the presence of acid-fast bacteria in tissues that are indicative of
mycobacterial infection (Ghossein et al. 1992, Marchetti et al. 1998, Zink and Nerlich
2004, Miller et al. 2007, Pourahmad et al. 2009a,b).
Fixatives used to preserve tissues (regardless of whether it is a coagulant or noncoagulant fixative) and the total time the tissues are held in a fixative can result in
degradation of DNA and RNA within tissues (Dubeau et al. 1986, Fiallo et al. 1992, Foss
et al. 1994). Exposure of mycobacterial DNA to formalin causes production of Schiff
bases on the free amino groups of nucleotides (Fraenkel-Conrat 1954, Dubeau et al.
1986), and subsequent cross-linking between tissue proteins and DNA (Jackson and
Chalkey 1974). Therefore, time in formalin is considered to be one of the most important
57
criteria for successful DNA retrieval and PCR amplification following tissue processing
for histology (Ben-Ezra et al. 1991, Greer et al. 1991). Once the tissues are embedded in
paraffin wax, this degradation process is slowed and subsequent amplification of short
DNA sequences is successful (Shibata et al. 1988). Fish are often preserved at a research
laboratory, aquaculture facility or in the field, and then shipped many days later to
another laboratory for histology processing. Hence, there was particular interest in the
effects of time in fixative. Positive results were consistently obtained from the majority of
fish held in fixative up to 21 d, and the same occurred at 45 d with the exception of M.
chelonae infected fish fixed in NBF (i.e. only one fish tested positive). This positive
result at the later time points was somewhat surprising. Mycobacterium spp. has a unique,
waxy cell wall comprised of lipids and fatty acids (Kolattukudy et al. 1997), which may
exclude aldehydes to some extent. Perhaps this characteristic plays a role in preserving
DNA more than in non-acid fast bacteria exposed to formalin-based fixatives.
Dietrich’s, Davidson’s and Bouin’s fixatives are commonly used with fish tissues.
These contain acid and alcohol, both which cause denaturation and coagulation of
proteins and nucleic acids within tissue specimens by dehydration and disruption of
electrostatic and hydrogen bonding (Fournie et al. 2000, Eltoum et al. 2001). Tissues
preserved in Bouin’s solution (which contains both picric and acetic acid) are particularly
problematic for DNA retrieval (Greer et al. 1991). Interestingly, there was no reduction
in positive samples with Dietrich’s compared to 10% buffered formalin.
Some samples were negative in both fixatives at some of the early time points. The
inability to retrieve DNA logically would be influenced by the amount of bacterial DNA.
58
Incorporation of a bead beating enhances retrieval of DNA from formalinpreserved samples to liberate more DNA (Tripathi and Stevenson 2012). This was
included with the PCR protocol because from previous experiences it was found to
consistently yield more positive results with PCR from paraffin-embedded tissues.
Nevertheless, severity of infection did not affect the ability to detect mycobacterial DNA.
The results were similar to previous transmission studies regarding the virulence of M.
chelonae compared to M. marinum in zebrafish (Watral and Kent 2007, Ostland et al.
2008, Whipps et al. 2008). The M. marinum isolate was highly virulent and several fish
became moribund or died over the 8 wk period, and these fish were included in 45 d
samples. In contrast, all fish injected with M. chelonae became infected, but none
exhibited morbidity or mortality. The M. marinum infected fish had three times the
number of granulomas with acid-fast bacteria, but the recovery and detection of DNA
was not different than with fish infected with M. chelonae.
Most zebrafish cases submitted to the ZIRC and our laboratory for health
screening and diagnosis consist of multiple fish, and usually they are processed and
embedded within 3 weeks. Hence, even with only about 50-70% recovery of
mycobacterial DNA, diagnosticians can confidently use PCR on routinely processed and
paraffin-embedded zebrafish to identify these infections to the species level for many of
the fish within a particular case. This is important as the severity and distribution of
these various Mycobacterium spp. found in zebrafish are quite different, and fish health
managers and clinicians with species identifications in hand would be able to make more
informed decisions. For example, facilities with M. haemophilum infected zebrafish often
59
euthanize infected populations, disinfect the aquaria, and repopulate (Kent et al. 2009,
Kent et al. 2011, Whipps et al. 2012), and this approach is similarly recommended for M.
marinum outbreaks. In contrast, M. chelonae infections are presently managed by
cleaning tanks more frequently (Murray et al. 2011) or using different wild type strains of
zebrafish (Whipps et al. 2008).
Acid-fast bacteria are frequently observed in the intestinal lumen of zebrafish,
with and without infections in the visceral organs. Therefore, a concern with
amplification of DNA using whole fish sections may be confusion of mycobacteria in
granulomas versus intraintestinal mycobacteria that may be associated with other
constitutive microbiota in the intestinal tract. A solution for the potentially confounding
problem of intraintestinal bacterial contaminants would be to use histologic sections as a
template to guide in the removal of specific cores from the tissue block that correspond to
confirmed areas of infected organs (Sfanos et al. 2008) or, even more precisely, by using
laser capture microdissection to select individual intralesional granulomas containing
mycobacteria for PCR assays (Ryan et al. 2002, Zhu et al. 2003, Selva et al. 2004). Both
methods would allow for accurate selection of infected tissue sites within the infected
zebrafish prior to PCR analysis for mycobacterial identification and PCR primer-specific
molecular identification of Mycobacterium marinum and Mycobacterium chelonae.
60
ACKNOWLEDGEMENTS
The authors wish to thank the Oregon State University Veterinary Diagnostic
Laboratory for excellent histotechnical assistance. This work was supported by
NIH/NCRR grants T32 RR023917, R24 RR017886 and NIEHS Center grant P30
ES000210.
61
Table 2.1. Proportion of PCR positives (primer set HS5F/hsp667R) and mean granuloma
intensity in paraffin-embedded zebrafish (Danio rerio). Fish were preserved in either
10% neutral buffered formalin (NBF) or Dietrich’s fixative for various times before
processing. There was no statistical significance in prevalence between mycobacteria
species, fixatives or duration of fixation (all p ≥ 0.3, Fisher’s exact test). Granulomas/fish
are reported as the mean for the total group, followed by the mean of PCR negative and
positive fish in parentheses
Group
M. chelonae
3 days NBF
7 days NBF
21 days NBF
45 days NBF
3 days Dietrich’s
7 days Dietrich’s
21 days Dietrich’s
45 days Dietrich’s
M. marinum
3 days NBF
7 days NBF
21 days NBF
45 days NBF
3 days Dietrich’s
7 days Dietrich’s
21 days Dietrich’s
45 days Dietrich’s
PCR positive/ No. tested
Granulomas (-/+)
5/5
4/5
3/5
1/5
14.4
6.2 (9.0, 5.5)
15.4 (19.5, 12.7)
5.6 (5.5, 6.0)
4/5
4/5
4/5
5/5
8.4 (14.0, 7.0)
8.0 (1.0, 9.75)
11.8 (15.0, 11.0)
8.6
5/5
5/5
5/5
5/5
14.6
35.0
38.0
20.2
4/5
4/5
4/5
3/5
36.2 (26.0, 38.75)
33.0 (25.0, 35.0)
20.8 (16.0, 22.0)
29.0 (22.5, 33.3)
62
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Chapter 3
Paramecium caudatum enhances transmission and infectivity of Mycobacterium
marinum and Mycobacterium chelonae in zebrafish (Danio rerio)
Tracy S. Peterson1, Jayde A. Ferguson5, Virginia G. Watral1, K. Nadine Mutoji3, Don G.
Ennis4, Michael L. Kent1,2
Diseases of Aquatic Organisms, 2013. Submitted
1
Department of Microbiology, Oregon State University, Corvallis, Oregon, 97331
Department of Biomedical Sciences, Oregon State University, Corvallis, Oregon, 97331
3
Department of Biology, University of Texas-San Antonio, San Antonio, Texas, 78249
4
Department of Biology, University of Louisiana, Lafayette, Louisiana, 70504
5
Alaska Department of Fish and Game, Fish Pathology Laboratory, Anchorage, Alaska,
99518
2
69
ABSTRACT
Mycobacterial infections in laboratory zebrafish (Danio rerio) are common and
widespread in research colonies. Mycobacteria within free living amoebae have been
shown to be transmission vectors for mycobacteriosis. Paramecium caudatum are
commonly used as a first food for zebrafish, and we investigated this ciliate’s potential to
serve as a vector of Mycobacterium marinum and M. chelonae. The ability of live P.
caudatum to transmit these mycobacteria to larval, juvenile and adult zebrafish was
evaluated. Infections were defined by histologic observation of granulomas containing
acid-fast bacteria in extraintestinal locations. In both experiments, fish fed paramecia
containing mycobacteria became infected at a higher incidence than controls. Larvae
(exposed at 4 days post hatch) fed paramecia with M. marinum exhibited an incidence of
30% (24/80) and juveniles (exposed at 21 days post hatch) showed 31% incidence
(14/45). Adult fish fed a gelatin food matrix containing mycobacteria within paramecia
or mycobacteria alone for 2 wk resulted in infections when examined 8 wk after exposure
as follows: M. marinum OSU 214 47% (21/45), M. marinum CH 47% (9/19), M.
chelonae 38% (5/13). In contrast, fish feed mycobacteria alone in this diet did not
become infected, except for 2 fish (5%) in the M. marinum OSU 214 low dose group.
These results demonstrate that Paramecium caudatum can act as a vector for
mycobacteria. This provides a useful animal model for evaluation of natural
mycobacterial infections and demonstrates the possibility of mycobacterial transmission
in zebrafish facilities via contaminated paramecia cultures.
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INTRODUCTION
The use of zebrafish (Danio rerio) in biomedical research has dramatically
increased in recent years (Vascotto et al. 1997, Grunwald & Eisen 2002, Amsterdam &
Hopkins 2006). Similar to other laboratory animal models, underlying infections in
zebrafish can compromise health as well as research because chronic subclinical diseases,
such as mycobacteriosis, are of concern as they induce non-protocol induced variation in
experiments (Baker 2003, Kent et al. 2011, Kent et al. 2012, Lawrence et al. 2012).
Chronic disease caused by Mycobacterium spp. is the second most common
microbial infection in zebrafish research colonies
(http://zebrafish.org/zirc/health/diseaseManual.php). Mycobacterium chelonae is the
most common species found in laboratory zebrafish, and often presents as a subclinical
infection (Watral & Kent 2007, Whipps et al. 2008, Whipps et al. 2012). Mycobacterium
haemophilum is more pathogenic and infections are associated with high mortalities and
severe infections (Whipps et al. 2007). In contrast, Mycobacterium marinum is not
frequently seen in zebrafish, but when it occurs it has been associated with acute to
chronic infections and also high mortality (Broussard and Ennis 2007, Watral and Kent
2007, Ostland et al. 2008). Importantly, we have recently verified M. marinum outbreaks
in two separate, large-scale research zebrafish facilities.
Mycobacteriosis in zebrafish presents many challenges to effective management
and control of the disease once established within research colonies (Astrofsky et al.
2000, Kent et al. 2009). The mycobacterial species infecting laboratory zebrafish,
including Mycobacterium marinum, M. haemophilum and Mycobacterium chelonae, can
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be commonly isolated from the aquaria holding fish, water supplies and effluents (Primm
et al. 2004, Whipps et al. 2007, Whipps et al. 2008). Persistence and replication of
mycobacteria in the aquatic environment is aided by the bacteria’s inherent resistance to
chlorine, survival in low-nutrient environments, (starvation resistance), thermoresistance
and ability to form biofilms (Nyka 1974, Schulze-Robbecke and Buchholtz 1992,
Schulze-Robbecke et al. 1992, Taylor et al. 2000, Hall et al. 2006).
Current information regarding natural transmission and vectors of mycobacteria
in laboratory fishes, particularly zebrafish, is not substantive. There are several known
and utilized methods of mycobacterial transmission, representing non-natural
experimental infections, which includes intraperitoneal and intramuscular injection, oral
gavage and bath exposure (Prouty et al. 2003, Harriff et al. 2007, Watral & Kent 2007).
Water bath exposure of Japanese medaka (Oryzias latipes) to very high concentrations of
mycobacteria directly from cultures is ineffective and rarely resulted in infected fish,
usually with low infection levels, in laboratory transmission studies (Mutoji 2011).
Infection by feeding on infected fish tissues is one recognized route of transmission (Ross
1970, Hedrick et al. 1987, Mutoji 2011) and could be considered as a more natural route
of infection. It has been previously shown that mycobacteria residing within granulomas
have increased virulence, which provides an explanation for the ease of oral-enteric
transmission from infected tissues (Volkman et al. 2004). Another potential route of
infection is by feeding on invertebrates colonized by mycobacteria. Mutoji (2011)
showed that M. marinum within the gastrointestinal tract of mosquito larvae fed to
Japanese medaka were significantly more infectious than cultured mycobacteria for
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infection in Japanese medaka following feeding on these larvae. Mycobacteria residing
within amoebae are also more infectious than their counterparts from axenic cultures.
Cirillo et al. (1997) showed that Mycobacterium avium pathogenicity is increased if it is
phagocytized by Acanthamoeba castellanii and replicates within amoebic vacuoles.
Free-living aquatic protozoans, such as Acanthamoeba and Dictyostelium,
evolved as phagocytic cells that actively prey on bacteria (Barker & Brown 1994) and
inhabit the same niches as environmental mycobacteria. Mycobacteria residing in
macrophages also have enhanced virulence, and hence these amoebae act as surrogate
macrophages in the environment. Harriff et al. (2007) demonstrated that per os infections
of M. marinum in zebrafish were enhanced by passing the bacteria through amoebae.
Whereas amoebae are not deliberately feed to zebrafish, the filter-feeding ciliate
Paramecium caudatum is a common first food for larval zebrafish (Lawrence 2007,
Westerfield 2007, Harper & Lawrence 2010) and similar to environmental amoebae, they
are indiscriminant bacterivores.
Therefore, we conducted the following transmission experiments to determine if
P. caudatum could serve as a vector and enhance natural transmission of M. marinum and
M. chelonae in larval, post-larval and adult zebrafish.
MATERIALS AND METHODS
General Fish and Fish Husbandry
Larval, three week-old and five month old AB strain zebrafish free of pre-existing
background diseases were obtained from the Sinnhuber Aquatic Research Laboratory at
Oregon State University. Fish were housed in a biosafety-level 2 (BSL-2) room. Larval
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fish were held in static containers, whereas 3 wk and adult fish held in flow-through 2.8
liter tanks in a modular zebrafish rack system (AquaneeringTM, San Diego, CA).
Incoming municipal water source was filtered, dechlorinated and heated. All fish were
held at 28o C with a 14/10 light/dark photoperiod.
Mycobacterium spp.
We chose two strains of Mycobacterium marinum and one strain of
Mycobacterium chelonae for this study. The M. marinum OSU 214 strain was initially
isolated from hybrid striped bass and zebrafish (Watral & Kent 2007, Ostland et al. 2008)
where it was a significant cause of mortality, whereas the M. marinum isolate from a
recent outbreak (herein referred to as “M. marinum CH”) was cultured from a laboratory
colony of albino zebrafish experiencing a chronic and ongoing mortality event. The M.
chelonae H1/E2 isolate originated from a TU strain zebrafish, as part of a laboratory
colony with ongoing chronic morbidity and low mortality levels (Whipps et al. 2008).
Mycobacteria were prepared for inocula by making a 5 ml suspension in sterile distilled
water with an estimated optical density of 1 and 3 (McFarland standard), respectively,
which approximated the low and high dose values. The specified dosages were confirmed
by serial dilution plating (Tables 1 and 2). The total dose/fish was calculated based upon
serial dilution plating on CNA (colistin/nalidixic acid and blood) and Middlebrook 7H10
agar plates (RemelTM, Lenexa, KS 66215 USA) with a 100 µl aliquot of the gelatin diet
(minus the gelatin) including the 5 ml mycobacteria inoculum, with or without
paramecia. We include CNA medium as our previous studies have shown that this
medium is particularly useful for culture of M. marinum OSU 214 (Ostland et al. 2008).
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Paramecium culture and infections
Paramecia were cultured using standard methods
(http://zfin.org/zf_info/zfbook/chapt3/3.3.html), which included maintaining a
sustainable infusoria for the paramecia by adding sterile (autoclaved) wheat berries plus
yeast extract to sterile aquarium water. To “infect” paramecia with mycobacteria, cultures
were incubated for 8 h with either M. marinum or M. chelonae (optical density of 1-3 on
the McFarland scale). They were then were collected in 50 ml screw-cap conical tubes
(Becton Dickinson, Franklin Lakes, NJ 07417 USA) and spun on a microcentrifuge
(Eppendorf, Hauppauge, NY 11788 USA) at 4,400 g for 5 minutes to concentrate the
paramecia. The semi-concentrated paramecia were re-suspended in 5 ml of sterile
aquarium water. This step was repeated twice for a total of three rinses, to reduce the
amount of exogenous mycobacteria in planktonic suspension. The final rinse constituted
the paramecia suspension to be added to a gelatin suspension. Paramecia were visually
evaluated using a stereo microscope to confirm viability prior to adding them to a gelatin
food matrix. Mycobacterial doses in the paramecia were determined by counting of
individual bacteria within food vacuoles. To enumerate the total number of mycobacteria
per paramecium, 30 replicates of 10 µl (larval zebrafish) and 25 µl (juvenile and adult
zebrafish) aliquots of the mycobacteria- loaded paramecia were placed onto clean glass
slides, air-dried for 30 minutes, fixed in 100% methanol for 5 minutes, air-dried and then
acid-fast stained by Kinyoun’s cold acid-fast staining method (Kent Lab protocol). On
average, there were 8 paramecia/10 µl for the larval zebrafish exposure groups and 18
paramecia/25 µl for each of the juvenile and adult zebrafish exposure groups (M.
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marinum OSU 214, M. chelonae and M. marinum CH), with each individual paramecium
containing 70 to 125 mycobacteria in food vacuoles. In the adult zebrafish exposure
groups, there were approximately 720 paramecia/ml, or between 69,000 and 71,000
mycobacteria/ml in the prepared inoculum containing paramecia. Paramecia not exposed
to mycobacteria were negative by both bacterial cultures on Middlebrook 7H10 agar
plates and acid-fast staining, which ensured that there was no background contamination
of mycobacteria in stock paramecia cultures.
Gelatin delivery vehicle
The base gelatin feed formulation was comprised of 16.25 g of unflavored gelatin
powder (HyTop GelatinTM, Arlington Heights, IL 60005 USA), 12.5 g of Otohimie “A”
Marine Fish Larval and Weaning FeedTM, (Marubeni Nisshin Feed Co., Tokyo, Japan),
6.25 ml of cod liver oil (Twinlab, American Fork, UT 84003 USA) and 120 ml of sterile
aquarium water. The mixture was gently heated to 60oC while agitated until the gelatin
went into solution, removed from the heat, allowed to cool to 40oC then 5 ml of either the
mycobacteria-loaded paramecia cultures or mycobacteria in sterile aquarium water were
added. The total stock feed volume was doubled and approximately sixteen 4 g tablets
were prepared in chilled metal molds and allowed to completely solidify, after which the
tablets were refrigerated. Prepared diets in the gelatin matrix were stored by refrigeration
at 5oC for two weeks. The diet was minced to pieces approximately 2 mm in diameter
prior to feeding.
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Survival of mycobacteria at refrigerated temperatures
We evaluated the stability of the mycobacteria under refrigerated temperatures as
the gelatin diet was prepared and held at 5oC for the 2 wk feeding regime. The feed
mixture minus gelatin was inoculated with 3 x 104 M. marinum OSU 214 and M.
chelonae, and then refrigerated at 5oC to simulate refrigerated storage conditions of the
gelatin diet. The feed mixture was cultured after 1 and 3 weeks, respectively, to
determine if mycobacteria in the feed continued to grow, decline or remained static.
Larval fish exposure
For the larval zebrafish feeding trial, 208 AB strain zebrafish were divided into
control, M. marinum OSU 214 and M. chelonae groups. Four days post-fertilization
(dpf), larvae were transferred to sterile 12 well cell culture plates (COSTARTM, Corning
Incorporated, Corning, NY 14831 USA flat bottom cell culture cluster with lid). One
larval fish was added to each well, which was filled with 3 ml of sterile aquarium water.
Every 8 hours, 2.5 ml of well water was replaced with fresh sterile aquarium water.
Larval zebrafish were fed 10 µl of free swimming paramecia with mycobacteria
suspension twice daily until 12 dpf, when they were switched over to “Yellow”
micronized feed (a blend of 50 g each of Zeigler AP100TM 100, 150 to 250 micron and 25
g Golden PearlTM 200 to 300 micron, Aquatic Eco Systems and Artemia International)
from 12 dpf to 24 dpf, then fish were euthanized and evaluated. Mortalities and larvae
surviving to the conclusion of the feeding trial were placed into 15 ml conical tubes
containing 14 ml of 10% neutral buffered formalin.
77
3 wk-old fish exposure
Three week-old zebrafish (juveniles) were exposed to either M. marinum OSU
214 or M. chelonae in flow through tanks. Fish were fed 25 µl paramecia containing
mycobacteria twice daily for 7 d, and the experiment ended at 56 d post exposure (pe)
(Table 3.1). Exposed and control fish were held in 3 replicate flow-through 2.8 L tanks
for each treatment, with 25 fish/tank.
Adult fish exposure
For the gelatin food matrix inoculated with either M. marinum OSU 214 strain, M.
marinum CH strain or M. chelonae H1/E2, five month-old AB strain, mixed sex zebrafish
were divided into 4 replicate 2.8 L tanks/exposure regime. Exposure groups were
comprised of fish fed both high and low dose mycobacteria in the gelatin food matrix,
mycobacteria and paramecia in gelatin food matrix, and gelatin food matrix only for
control groups. Each tank in the study was fed 1 g of the gelatin food matrix daily for 14
d. Fish were then switched to a standard zebrafish commercial diet mixture for another 6
wk until the end of the experimental exposure time. In the M. chelonae treatment groups,
fish were examined at two time points, 8 and 16 wk pe (Table 3.2).
Histology
Moribund and surviving fish were euthanized with an overdose of tricaine
methanesulfonate (FinquelTM, Argent Chemical Laboratories, Redmond, WA 98052
USA) and processed as follows. All zebrafish in the transmission experiments, with the
exception of the larval zebrafish, were processed for histology in an identical manner.
Briefly, immediately after death, the coelomic cavity of each fish was opened and half of
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each operculum removed to facilitate fixation and mitigate post-mortem autolysis. The
fish were then placed in 10X volume/fish of 10% neutral buffered formalin in 50 ml
conical tubes and allowed to fix overnight on a rocker plate. After initial fixation, the fish
were put into 10X volume/fish of CalExIITM (Fisher Scientific, Fair Lawn, NJ 07410
USA) for 48 h of decalcification, then rinsed in distilled water for 30 minutes prior to
being sagittally sectioned, placed into standard tissue cassettes and processed for routine
histology. Larval zebrafish were processed identically except for the decalcification and
rinsing step, and then placed into gelatin molds prior to processing and embedding.
Hematoxylin and eosin stain and Ziehl-Neelsen or Kinyoun’s aid-fast stains were
performed for all fish.
Statistics
All statistical analyses were conducted with the program R, version 2.7.2 (R
Development Core 315 Team). Significance was set at p ≤ 0.05 and p-values are 2-tailed.
For each experiment, differences in infection prevalence between tanks of each treatment
group were tested by the Pearson’s Chi-square test for count data in order to determine
whether data from different tanks could be pooled for group comparisons. The Monte
Carlo simulation method using 20,000 replications was used to estimate p-values for this
test. Fisher’s exact tests were used to compare infection prevalence between treatment
groups in each experiment. Because M. chelonae is less virulent than M. marinum and
can occur in a chronic state, we were also interested in comparing overall infection
prevalence of this species between experimental groups, regardless of the time point that
was sampled. We used Fisher’s exact tests for this comparison after data from different
79
time points had been assessed for permissible pooling by comparing infection
prevalences with Fisher’s exact tests.
RESULTS
Larval fish
This experiment entailed feeding fry zebrafish with paramecia that had been
incubated for 8 h in a culture of M. marinum OSU 214 or M. chelonae. Several larval fish
that were fed paramecia containing mycobacteria showed the presence of mycobacterial
aggregates within the intestinal lumen, and a few developed infections of the swim
bladder when examined between 3 and 14 days pe (Table 3.1; Fig. 3.1b,c). In the M.
marinum group, acid-fast bacteria were observed in the intestinal lumens of 30% (24/80)
of infected fish. Within the M. chelonae group, acid-fast bacteria were observed in the
intestines of 39% (44/112) of infected fish, while 2% of infections occurred in the swim
bladder. In the fish with swim bladder infections, small aggregates of mycobacteria (M.
chelonae) were also observed within the pneumatic duct (Fig. 3.1b). Mycobacteria
infections observed in the larval zebrafish groups, regardless of whether infected by M.
marinum or M. chelonae, were comprised of mycobacteria aggregates in close
association with epithelial surfaces, with no intracellular mycobacteria and minimal to no
inflammatory cell infiltrates or granuloma formation (Fig.3.1c). All control larval
zebrafish were negative (0/16) for mycobacteria infection.
Juveniles
In three week-old zebrafish fed infected paramecium, about one third of the fish
exposed to M. marinum became infected, while fish that were exposed to M. chelonae
80
showed only 3% prevalence of infections. Infections for both mycobacterial species were
confirmed by identifying granulomas containing acid-fast bacilli, which were primarily
localized to kidney, spleen (Fig.3.1e,f) and liver. Rarely, there was involvement of the
choroid rete, brain, and gonad (ovary, and less often testis). These infrequent infections
were confined to fish in the M. marinum group.
Adult fish
This experiment entailed feeding a gelatin food matrix containing M. chelonae
and two strains of M. marinum to adult zebrafish, either by placing bacteria directly into
the diet from cultures or by feeding mycobacteria to paramecia, which were subsequently
added to gelatin food matrix. Surviving fish were euthanized and evaluated by histology
at 8 wk pe for the M. marinum groups, and at 8 and 16 wk for the M. chelonae groups.
Across all groups, several fish were inadvertently lost before the conclusion of the study
due to the lack of an in-tank screen baffle, which resulted in failure of outflow
containment. Hence, between 32 and 100% of the fish in each group were available for
examination compared to initial number of the fish in each group (Table 3.2). However,
no tank within any of the groups was completely devoid of fish at the termination of the
experiment. Many fish developed extraintestinal mycobacteriosis when exposed toany of
the three mycobacteria strains. In contrast, of the 305 fish examined from either high or
low dose groups, 302 (99.4%) were free of extraintestinal infection. However, many of
the fish in all exposures, regardless of their exposure regime had acid-fast bacteria
(presumably Mycobacterium spp.) in the lumen of the intestine (Fig. 3.2).
81
M. marinum OSU 214
At 7 wk pe in this experiment in all groups there was a mortality event. This
occurred across all exposure regimens including unexposed controls, and was attributed
to gas supersaturation. These fish were included in the data analysis as this occurred
within 1 wk of the final data collection time. Ten of these fish were from the M.
marinum OSU 214 paramecia group, and 3 of these fish had severe systemic
mycobacteriosis. All the other mortalities from the other groups (n = 23) did not exhibit
mycobacteriosis. Fish fed mycobacteria with the paramecium vector had an approximate
50% incidence of infection, with the infections occurring in all replicate tanks (Table 3.2;
Fig. 3.2a). In contrast, only 2 fish in one of the other groups (low dose mycobacteria
without paramecia) demonstrated extraintestinal infections. Fish from all groups,
including those not fed mycobacteria, showed acid-fast bacteria in the intestinal lumen
(Fig. 3.2a).
Regarding statistical comparison, no difference was observed amongst tanks
within each exposure regime (Chi-squared test with Monte Carlo simulation). Therefore,
data from each tank within each exposure regime were pooled. Prevalence of infection in
the paramecia group was significantly greater than all other groups (p < 0.001; Fisher’s
exact test), whereas there was no significant difference between the low dose group
compared to the high dose, and between control groups and the low dose infected group
(< 0.001; Fisher’s exact test).
82
M. marinum CH
A similar pattern of infection occurred with the second strain of M. marinum.
Only fish exposed to M. marinum CH via the paramecia were infected (Table 3.2; Fig.
3.2b), whereas other groups had mycobacteria only in the intestinal lumen of some fish.
The infected paramecia treatment group had an overall infection prevalence of 47.4%,
which was statistically different than all other groups. (p < 0.001; Fisher’s exact test).
Infection prevalence amongst tanks within each treatment regime was not statistically
different among tanks (p = 0.532; Chi-squared test with Monte Carlo simulation), and
hence pooling data across tanks was permissible.
M. chelonae
As M. chelonae infections are more chronic and less virulent than M. marinum, it
was decided to examine fish as subsamples collected at 8 and 16 wk pe (Table 3.2; Fig.
3.2c,d). There were no extraintestinal infections in the low or high dose groups at either 8
or 16 wk pe. The infected paramecia treatment group had an infection prevalence of
38.5% at 8 wk (pe), which was statistically different from uninfected controls (p = 0.04;
Fisher’s exact test).
At 16 wk pe 18.8% of the infected paramecia treatment group was infected, which
did not differ from the unexposed controls (p = 0.22; Fisher’s exact test). Infection
prevalence in infected paramecia treated fish did not statistically differ among tanks at 8
wk (p = 0.710; Chi-squared test with Monte Carlo simulation) or 16 weeks (p = 0.092;
Chi-squared test with Monte Carlo simulation), so pooling data across tanks was
permissible. There was also interest in the overall M. chelonae infection in infected
83
paramecia treated fish, regardless of the time point sampled. There was no statistical
difference between prevalence of paramecia treated fish sampled at 8 and 16 wk pe (p =
0.420; Fisher’s exact test), so data were pooled and the resulting overall infection
prevalence of 29.6%, which was significantly different than uninfected controls and fish
exposed via the gelatin food matrix without paramecia (p = 0.005; Fisher’s exact test).
Adult fish histology
M. marinum and M. chelonae infected fish exhibited the typical multifocally
extensive pattern of mycobacteriosis, similar to the histological presentations seen in
infected juvenile fish (Fig. 3.2), with prominent granuloma formation involving the
kidney, spleen, liver, ovary, coelomic adipose tissue and less often the endomeninges,
choroid rete, testis and the inner ear. Granulomas were variable in their chronicity, with
the majority being well-organized and containing necrotic centers filled with cellular
detritus. In all granulomas, variable numbers of acid-fast bacilli were observed, some
within necrotic centers in addition to smaller intracellular aggregates within
macrophages. No colonization of the swim bladder was seen in any of the infected fish,
regardless of mycobacterial species. In a large percentage of fish exposed to M. marinum,
M. chelonae or unexposed controls, variable numbers of mycobacterial aggregates were
transiently present in the intestinal lumen, with no obvious epithelial association or
colonization of the intestine (Fig. 3.1d).
84
Survival of mycobacteria at 5o C
Feed mixture containing mycobacteria, minus the gelatin, were held for up to 2
wk in a refrigerator at 5o C, and thus it was important to determine survival and growth of
the mycobacteria under these conditions. In all cases, whether cultured at 1 or 3 wk after
preparing the diet, CFU plate counts for both M. marinum OSU 214 and M. chelonae
were initially 3 x 104 mycobacteria/5 ml, remained the same at 1 wk and decreased
slightly to 2.8 x 104 at 3 wk.
DISCUSSION
Previous studies have shown that mycobacteria within amoebae, specifically the
environmental amoebae Acanthamoeba castellani, Acanthamoeba polyphaga and
Dictyostelium discoideum, can successfully survive, replicate, and may be potentiated for
infectivity and spread of infection in the host (Cirillo et al. 1997, Adekambi et al. 2006,
Hagedorn et al. 2009). Mycobacteria from granulomas are more infective than those
from media cultures (Li et al. 2002). The enhanced infectivity of mycobacteria residing in
amoeba vectors has been attributed to them acting somewhat as surrogate macrophages,
with up-regulation of virulence genes when mycobacteria are within the intracellular
environment (Danelishvili et al. 2004, Tenant & Bermudez 2006). Solomon et al. (2003),
in concordance with this observation, found that Mycobacterium marinum effectively
replicates within Dictyostelium by halting phagosomal maturation, and that mechanisms
of cellular invasion elicited by M. marinum may be putatively conserved between
amoebae and mammalian macrophages (Hagedorn & Soldati 2007).
85
This study demonstrated for the first time that a ciliate protozoan, Paramecium
caudatum, like amoebae, can be a transmission vector for mycobacteria as mycobacterial
infections occurred by feeding paramecia containing either M. marinum or M. chelonae.
Moreover, transmission was dramatically increased with the paramecia vector compared
to fish fed bacteria alone. It is likely the prevalent infections with the ciliate were due to a
similar phenomenon of increased infectivity as reported with macrophages and amoebae,
as fish fed mycobacteria in the paramecia actually received much lower amounts of
bacteria compared to those fed the high doses of bacteria alone (Table 3.2).
Mycobacterial concentrations in paramecia were determined by directly counting
bacteria within paramecia using acid-fast stains. It is likely that the total counts were
actually slightly less for the paramecia group as non-viable mycobacteria could not be
eliminated in the counts. Mycobacteria could not be counted after being placed in
gelatin; perhaps due to inhibitory effects of the collagenous matrix, but 40o C does not
appreciably reduce the viability of mycobacteria (Merkal & Whipple 1980, Gao et al.
2002). We also showed that mycobacteria counts were essentially unchanged at 5 oC. The
total amount of mycobacteria given to adult zebrafish within the paramecia vector over
the 2 wk period using the three mycobacterial stains was about 6 x 106 bacteria/fish.
Harriff et al. (2007) achieved 25% M. marinum infection (based on granuloma formation)
in zebrafish gavage fed with amoebae containing 1.3 x 105 mycobacteria/fish in a single
dose. This was a lower total dose compared to this study, but similar to the daily dose.
These authors also achieved 67% and 35% prevalence of M. marinum infection by
exposing zebrafish fish by gavage feeding at 3 x 107 and 8 x 105, respectively. These
86
findings are quite different than those from this study using two strains of M. marinum
fed without paramecia, as no fish became infected when exposed to a total dose of either
6.4 x 108 or 8.5 x 106 . One plausible explanation is that the fish in this study were
exposed under more natural feeding conditions and over a 2 wk period, rather than a
single bolus dose forcibly introduced to the intestine. Also, zebrafish are relatively small
and agastric, so it may be possible that the intestinal epithelium was traumatically
breached with the plastic gavage tube during exposure in earlier study. Another
comparable transmission study was conducted by Mutoji (2011), where mosquito larvae
containing M. marinum were fed to Japanese medaka. Fish receiving 4 or 7 feedings
resulted in about 90% prevalence of infection, with a calculated dose of about 4 to 7 x 104
mycobacteria/fish, respectively.
Only two fish that were not fed paramecia developed extraintestinal infections.
These occurred in the low dose M. marinum OSU 214, but the high dose group feed the
same bacteria were not infected. It is possible that these two fish had pre-existing
background infections, as we have documented minimal background mycobacteriosis in
zebrafish at the laboratory providing the fish for the present study (Kent et al. 2004, Kent
et al. 2011).
Mycobacterium species capable of causing disease in fish may occur in aquaria in
the absence of disease (Beran et al. 2006). Indeed, all groups of the fish in this study,
including controls that were not deliberately exposed to mycobacteria, exhibited the
presence (perhaps colonization) of large amounts of acid-fast bacteria in the intestinal
lumen. This demonstrates that the presence of mycobacteria, even if ingested, does not
87
always result in extraintestinal infections or may represent non-virulent intestinal
microflora. The latter is plausible as we frequently observed acid-fast bacteria in the
intestinal lumen of control fish. As discussed above, one factor allowing for these
infections may be changes in gene expression as may occur whether the mycobacteria are
in macrophages, amoebae or invertebrate hosts.
Natural mycobacterial infections occasionally present as aerocystitis (swim
bladder infection) in zebrafish (Whipps et al. 2008). We observed this presentation in
larval fish, but not adults. Physostomous teleost fish, such as the zebrafish, have open
communication (pneumatic duct) between the distal oropharynx and swim bladder and
several mycobacterial species are known to colonize the swim bladder and cause
aerocystitis (Matthews 2004, Whipps et al. 2008, Kent et al. 2011). In larval zebrafish, it
may be anatomically favorable for non-motile mycobacteria to colonize the swim
bladder, as the total distance traveled from the oropharynx to pneumatic duct and swim
bladder is considerably shorter than in juvenile and adult zebrafish, and this may provide
one explanation for the predilection of mycobacteria to localize in the larval swim
bladder. Another possible explanation for the affinity of mycobacteria to the zebrafish
swim bladder may be a correlation between the higher cholesterol content of swim
bladder surfactant lipid, when compared to mammalian lung surfactant, and the
requirement of mycobacterial species to import and catabolize cholesterol to maintain
persistent infection in the host (Orgeig et al. 2003, Pandey & Sassetti 2008). Two of the
present authors (M.K. and T.P.) have provided histologic interpretations for numerous
diagnostic cases submitted to the Zebrafish International Resource Center diagnostic
88
program, Eugene, Oregon. As reported by Whipps et al. (2008), there have been several
confirmed cases of mycobacterial aerocystitis in zebrafish with mycobacteriosis from
many laboratories both in the United States and internationally. Perhaps these represent
fish with very chronic infections that were established as larvae or juvenile fish.
The teleost swim bladder shares many conserved anatomical features with the
mammalian lung, from comparative ontogeny (Perry et al. 2004) to conserved gene
expression and transcriptomics in molecular developmental pathways (Winata et al. 2009,
Zheng et al. 2011), and thus the model that we developed with larval zebrafish may be a
useful surrogate for investigating certain aspects of human pulmonary tuberculosis.
Since amoebae and ciliates occurring naturally in the aquatic environment can be
vectors for mycobacteria, these organisms are likely an important route of transmission
and subsequent infection in zebrafish. Moreover, live foods such as aquatic protozoans,
worms and crustacean larvae are deliberately included as a dietary staple of larval,
juvenile and adult broodstock zebrafish (Watanabe et al. 1983, Lawrence 2007, Best et al.
2010). Commonly, these live feeds consist of natural prey items such as paramecia,
rotifers, water fleas (Daphnia spp.), Artemia spp. and chironomid insect larvae. Water
fleas in particular have been implicated as a source of mycobacteria within established
aquaculture systems (Grange 1985, Conroy & Conroy 1999, Somsiri et al. 2005).
Considering that paramecia are a common first feed for larval zebrafish, the
results shown in this study demonstrate one specific potential source of infection.
Therefore, facility staff involved in propagating zooplankton, such as paramecia, as a
food source for larval zebrafish should exercise caution to avoid exposing these cultures
89
to potential sources of mycobacteria. Future mycobacteria transmission studies with live
feeds commonly used in laboratory zebrafish larviculture, including rotifers and Artemia
spp., are warranted to determine whether these microinvertebrates can also act as vectors
for mycobacterial infection within zebrafish colonies. Given that a wide variety of ciliate
species occur in the aquatic environment and undoubtedly encounter mycobacteria,
further studies are warranted to compare and contrast the inter-relationship of paramecia
with mycobacteria to the amoebae models.
ACKNOWLEDGEMENTS
The authors thank the Oregon State University Veterinary Diagnostic Laboratory,
especially Kay Fisher, Misty Corbus and Renee Norred for their excellent histotechnical
assistance with slide preparation as well as Colleen Paquette and Justin Sanders for
additional technical assistance. This work was supported by NIH/NCRR grants T32
RR023917, R24 RR017886 and NIEHS Center grant P30 ES000210.
90
Figure 3.1. Danio rerio. Histological presentations in zebrafish exposed to
Mycobacterium marinum and Mycobacterium chelonae residing within paramecia food
vacuoles. (a) Paramecium caudatum cultured with M. marinum OSU 214 for 8 h, with
numerous acid-fast bacteria in food vacuoles. Kinyoun’s acid-fast. Scale bar = 10 µm. (b)
Mycobacteria (arrow) in pneumatic duct of larval fish exposed to M. chelonae. ZiehlNeelsen acid-fast. Scale bar = 10 µm. (c) M. chelonae aggregate in larval fish swim
bladder (arrows). Ziehl-Neelsen acid-fast. Scale bar = 25 µm. (d) Mycobacteria (arrow)
in the intestinal lumen of an adult fish. Kinyoun’s acid-fast. Scale bar = 25 µm. (e) Liver
(L) and spleen (S) granulomas in a juvenile fish fed paramecia containing M. marinum
OSU 214. Arrow = granuloma demonstrated in Fig. 1f. Hematoxylin and eosin. Scale
bar = 100 µm. (f) High magnification of spleen granuloma with numerous mycobacteria.
Kinyoun’s acid-fast. Scale bar = 25 µm.
91
Figure 3.2. Prevalence (%) of extraintestinal mycobacteriosis (white bars) and presence
of mycobacteria in the intestinal lumen (gray bars). Data points indicate means from
each of four tanks for each mycobacteria strain. A. Mycobacterium marinum OSU 214 at
8wk pe. B. Mycobacterium marinum CH at 8 wk pe. C. Mycobacterium chelonae at 8
wk. D. M. chelonae at 16 wk pe.
92
Table 3.1. Danio rerio. Larval and juvenile zebrafish exposed by feeding Paramecium
caudatum containing mycobacteria for 7 d. Larval and juvenile fish were examined at 24
d and 8 wk post initial feeding, respectively.
1
Total fish infected are numbers expressed (without parenthesis) and relative percentages
infected (within parenthesis)
Mycobacteria
Larval M. marinum (OSU 214)
Larval M. chelonae (H1/E2)
Larval Control
Juvenile M. marinum (OSU
214)
Juvenile M. chelonae (H1/E2)
Juvenile Control
Dose/Fish/Day
4 x 105
3.8 x 105
0
3.2 x 105
No. Fish
80
112
16
45
Total Fish Infected1 (%)
24(30)
44(39)
0(0)
14(31)
3.3 x 105
0
70
35
2(3)
0(0)
93
Table 3.2. Danio rerio. Adult zebrafish fed gelatin food matrix containing mycobacteria or Paramecium caudatum containing
mycobacteria
1
No. Fish Examined includes mortalities within M. marinum (OSU 214) experimental and control groups occurring 1 week
prior to conclusion of experiment
2
Total fish infected are numbers expressed (without parenthesis) and relative percentages infected (within parenthesis)
3
Total fish positive for intestinal acid-fast bacilli are numbers expressed (without parenthesis) and relative percentages infected
(within parenthesis)
Mycobacteria
Dose/Fish/Day
Days
Fed
Duration
(Post-Exposure)
No. Fish
Examined1
Total Fish
Infected (%)
Intestinal AcidFast Bacilli (%)
M. marinum (OSU
214)
Paramecia
High Dose
Low Dose
Control
3.4 x 105
6.1 x 105
3.6 x 104
0
14
14
14
14
8 wk
8 wk
8 wk
8 wk
45
56
42
60
21(47)2
0(0)
2(5)
0(0)
12(34)3
20(41)
18(47)
31(51)
M. marinum (CH)
Paramecia
High Dose
Low Dose
Control
3.6 x 105
4.6 x 107
3.8 x 106
0
14
14
14
14
8 wk
8 wk
8 wk
8 wk
19
22
21
20
9(47)
0(0)
0(0)
0(0)
8(42)
14(64)
10(48)
18(90)
M. chelonae (H1/E2)
Paramecia
High Dose
Low Dose
Control
3.4 x 105
8.3 x 107
3.5 x 106
0
14
14
14
14
8 wk
8 wk
8 wk
8 wk
13
14
16
11
5(38)
0(0)
0(0)
0(0)
12(92)
11(78)
11(69)
5(45)
Paramecia
High Dose
Low Dose
Control
3.4 x 105
8.3 x 107
3.5 x 106
0
14
14
14
14
16 wk
16 wk
16 wk
16 wk
14
16
15
12
3(21)
0(0)
0(0)
0(0)
11(79)
12(75)
6(40)
9(75)
94
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zebrafish swimbladder: The requirements of Hedgehog signaling in specification and
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Zheng W, Wang Z, Collins JE, Andrews RM, Stemple D, Gong Z (2011) Comparative
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Chapter 4
Persistence of Mycobacterium marinum in Paramecium caudatum Digestive Vacuoles
Tracy S. Peterson1, Justin L. Sanders1, Michael L. Kent1,2
Journal of Eukaryotic Microbiology, 2013. Submitted
1
Department of Microbiology, 220 Nash Hall, Oregon State University, Corvallis,
Oregon, 97331
2
Department of Biomedical Sciences, Oregon State University, Corvallis, Oregon, 97331
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ABSTRACT
The freshwater ciliate Paramecium caudatum is not only a popular model for
protist research, but serves as a food source for fry fish reared in culture. Historically,
Paramecium spp. has been an important first food source for laboratory-reared larval
zebrafish (Danio rerio) and many other aquarium fishes in the commercial pet trade.
Mycobacteriosis, caused by Mycobacterium marinum and other Mycobacterium species,
is a common disease of aquarium fishes, including zebrafish. Amoebae have been shown
previously to serve as vectors for Mycobacterium spp. and potentiate these infections. We
recently demonstrated a similar phenomenon with Paramecium caudatum serving as
vectors for orally transmitted M. marinum and M. chelonae in zebrafish. Environmental
bacteria serving as a food source for paramecia are engulfed by membrane-bound
digestive vacuoles (phagolysosome) and are further processed during cyclosis,
culminating in degradation and elimination of bacterial products. A key question
remaining to be answered involving this transmission cycle is whether mycobacteria
would persist within digestive vacuoles of the paramecia vector longer than the typical
digestion cycle reported with Gram-negative bacteria. Our results show that
Mycobacterium marinum persists in paramecia digestive vacuoles for many days after
ingestion, while Escherichia coli is eliminated from vacuoles within an hour. We
conducted two trials using GFP-labeled M. marinum, and evaluated the concentrations
and persistence for several days. In both trials, mycobacteria significantly decline by 24
hours post-ingestion and persisted in paramecia up to one week post-exposure.
101
INTRODUCTION
Bacteria are the natural food source for ciliates present in the environment
(Hargitt and Fray 1917). Feeding habits of Paramecium spp., a ciliate protozoan
organism, demonstrate that they are filter-feeding bacterivores of planktonic bacteria in
aquatic systems (Hausmann 2002). They select bacteria based upon criteria such as size,
shape, age, fatty acid composition, and whether or not the bacteria form clumps or long
chains, which may be detrimental to engulfment and digestion (Wichterman 1953, Barna
and Weis 1973). Food processing in paramecia is accomplished initially by the formation
of digestive vacuoles, which takes approximately one minute after the recycling of
existing vacuole membranes to be formed and then released into the cytoplasm (Fok et al.
1988). Flow of the digestive vacuole is compartmentalized, with immediate postingestion acid phosphatase activity, merging of the acidosome and phagosome, rapid
acidification of the intravacuolar space that lowers the pH from 7 to <3, secondary
lysosome fusion with the digestive vacuole, followed by vacuolar re-alkalization (Fok
and Allen 1993). The entire sequence of ingestion, processing and egestion of food
material (i.e., clearance rate) in paramecia, including inert bacteria-sized latex beads,
occurs over approximately 20 minutes, with an average lifespan of a digestive vacuole
between 20 minutes to 1 hour (Fok and Allen 1993, Verni and Gualtieri 1997).
Zebrafish (Danio rerio) are important model species in biomedical research, and
hence they are reared at many institutions (Vascotto et al. 1997, Grunwald & Eisen 2002,
Amsterdam & Hopkins 2006). Mycobacteriosis is one of the most common diseases
afflicting zebrafish in research facilities (Whipps et al. 2012). Amoebae have been
102
shown to act as vectors and potentiate infections by mycobacteria, including infections in
zebrafish (Harriff et al. 2007). Whereas amoebae are not deliberately feed to zebrafish,
the filter-feeding ciliate Paramecium caudatum is a common first food for larval
zebrafish (Lawrence 2007, Harper & Lawrence 2010) and similar to environmental
amoebae, they are indiscriminant bacterivores.
We have previously demonstrated successful ingestion and digestive vacuole
sequestration of Mycobacterium marinum and Mycobacterium chelonae, two established
zebrafish bacterial pathogens, by Paramecium caudatum, and we also confirmed that
Paramecium caudatum can serve as an effective transmission vector of zebrafish
mycobacteriosis (Peterson et al. 2013). In contrast to more readily digested bacteria,
mycobacteria in amoebae survive for a more extended period and inhibit phagolysosome
formation (Salah et al. 2009). Here we investigated the duration and intracellular location
of Mycobacterium marinum within paramecia.
MATERIALS AND METHODS
Paramecia Culture
Paramecia (Paramecium caudatum) were cultured for the series of experiments
following the established Zebrafish International Resource Center (ZIRC), University of
Oregon, Eugene, Oregon protocol. Briefly, a 3 ml subset of stock paramecia culture
obtained from ZIRC was inoculated into paramecia culture media (ZIRC paramecia
procedure, rev. 2011) in 250 mm sterile plastic petri dishes containing 150 ml of sterile
aquarium water, yeast and wheat berries. This results in indiscriminate bacterial growth,
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which the paramecia feed upon. The paramecia cultures were incubated at 25oC for two
weeks or until culture density was at least 10 paramecia/5 µl.
We obtained mature paramecia for the exposure trials as follows. Culture media
containing both immature and mature forms of paramecia was filtered through a 20 µm
nylon mesh filter to separate the age classes, as only large, mature paramecia were
desired for the series of experiments. The retained paramecia were then rinsed three times
with a total of 60 ml culture media filtrate (containing no paramecia or other
microinvertebrates) in order to separate immature paramecia from the remainder. The
retained paramecia were filtered into a 35 x 10 mm plastic sterile petri dish (Falcon®
Easy Grip Petri Dish, Becton Dickinson, Franklin Lakes, NJ USA 07417-1886) and
saved for future use.
Mycobacteria Exposure and Persistence Evaluations
Mycobacterium marinum OSU 214 GFP (M. marinum GFP) and Escherichia coli
strain DH5α/DsRed (E. coli DsRed) were prepared in 5 ml sterile aquarium water at an
optical density of approximately 1.0 McFarland scale, with the CFU plate counts on
Middlebrook 7H10 and LB + gentamicin agar 1.17 x 107 for M. marinum OSU 214 GFP
and 9.2 x 106 for E. coli DH5α DsRed, respectively. All mycobacteria were filtered
through a 20 µm nylon screen to remove large aggregates prior to inoculating paramecia
cultures. The OSU 214 isolate was selected as it was originally isolated from zebrafish
(Ostland et al. 2008) and was used in our experiments evaluating paramecia as a vector
for mycobacteria (Peterson et al. 2013).
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Paramecia in a 5 ml suspension were inoculated with 1 ml of M. marinum OSU 214 GFP
strain and exposed for 20 minutes as previously described. After 20 min., the paramecia
suspension was filtered through a 20 µm nylon screen to retain mature paramecia,
copiously rinsed with 50 ml of clean paramecia media filtrate to remove exogenous
mycobacteria, with the final rinse into a sterile 35 x 10 mm petri dish (Falcon® Easy
Grip Petri Dish, Becton Dickinson, Franklin Lakes, NJ). The culture was then held at
25o C.
Microscopy
Mycobacteria within the paramecia were evaluated in two separate trials, at
various time points up to 7 d (Figs 1- 8) as follows. Ten paramecia at each time point
were preserved in 10% neutral buffered formalin in a 0.75 ml microcentrifuge tube. After
incubation, paramecia were transferred to a 0.75 ml microcentrifuge tube in 250 µl
media, then an equal volume of 10% neutral buffered formalin was added to tube, and
allowed to fix the paramecia for 15 minutes. Then, a 10 µl of solution with paramecia
was placed between a coverslip bridge on a glass slide and observed using a Leica DMR
epifluorescence microscope (Leica Microsystems, GmbH) with the 40X objective using
FITC at an excitation BP 480 nm and emission BP 527 nm for GFP and rhodamine at an
excitation BP 546 nm and emission BP 600 nm for RFP fluorescence. Digital
photomicrographs were obtained with a SPOT RT3 Type 2.0 Mp Color RT2540 Slider
camera, serial # 275965 (Diagnostic Instruments, Sterling Heights, MI).
The amount of fluorescence in paramecia was used to quantitate and compare the
relative concentrations of GFP mycobacteria in digestive vacuoles by analysis of images
105
from ten paramecia at each time point. Both the density of fluorescence and area
occupied by fluorescence (white in gray scale images) was determined using ImagePro
Plus software (Version 5.1, MediaCybernetics). Areas of fluorescence for each
paramecium were selected using the range of intensity values which encompassed those
areas of the image that contained fluorescing bacteria, and were reported as the number
of pixels in each selected area (Fig. 2, 6).
Paramecia Cohabitation
We evaluated lateral transmission of mycobacteria between paramecia. Paramecia
that were not exposed to the mycobacteria were cohabitated with “infected” paramecia to
determine whether the large numbers of mycobacteria within vacuoles was due to actual
persistence versus egestion and re-ingestion of the bacteria. A negative paramecia
suspension was identically prepared as the infected paramecia. One infected paramecium
was placed along with a negative paramecium into each well of an 8 well chamber slide
(Lab-Tek® Chamber Slide™ System, Nalge Nunc International, Naperville, IL),
incubated in a humidity chamber at 25oC, and examined at 24 and 48 h. For this
experiment, paramecia on the slides were air-dried on the chamber slides, aqueous
mounting medium (VectaMount™ AQ Aqueous Mounting Medium, Vector
Laboratories, Burlingame, CA USA 94010) was added and the slide coverslipped. Slides
were then examined for fluorescing bacteria in paramecia as described above.
Statistical Analysis
Significant difference in the area of fluorescence between paramecia at the time
points was tested using a one-way analysis of variance (ANOVA) on the log-transformed
106
area measurements. Tukey’s honestly significant difference post hoc test was used to
find time points with a mean area of fluorescence which differed significantly with that of
time zero. All statistical analyses were performed using the software package R version
2.7.2 (R Development Core 315 Team).
RESULTS
Mycobacteria were quickly ingested, and digestive vacuoles were replete with
fluorescing bacteria (Fig. 4.1, 1 and 2), and the concentration of bacteria at 1 h was
similar to time zero (Fig. 4.1, 6). The second trial included evaluations at 4, 8 and 12 h
post-exposure and showed little decline, without statistical difference, compared to time
zero. At 24 h and later there was significant decline in bacterial concentrations in both
trials compared to time zero (Fig. 4.2, 7 and 8), and mycobacteria persisted within
digestive vacuoles for up to 1 wk (Fig. 4.1, 5). These calculations were derived from the
number of pixels within vacuoles that were scored as positive (i.e., white). We also
evaluated concretions by amount of fluorescence based on total brightness with vacuoles.
Results were very similar, and hence these data are not presented
Cohabitation of infected with uninfected paramecia showed very limited lateral
transmission. At 24 h, none of the 10 initially uninfected paramecia exhibited bacteria
exhibited fluorescing bacteria, whereas at 48 h few bacteria were observed in the
digestive vacuoles of two initially uninfected paramecia.
In contrast to mycobacteria, Dsred labeled E. coli was observed at time zero but
not 1 hour in one trial (n = 10). In a second trial, E. coli were observed within paramecia
digestive vacuoles at time zero and one hour, with none observed 24 h or thereafter. No
107
fluorescence was observed in control paramecia which had not been exposed to the
labeled E. coli or M. marinum.
DISCUSSION
Aquatic ciliates, such as Paramecium caudatum, have evolved as natural
predators of environmental bacteria (Winiecka-Krusnell and Linder 2001). Paramecia
track prey in aquatic environments by chemosensory mechanisms that cause changes in
membrane fluidity and initiate both K+ and Ca2+ action potentials, using chemical cues in
the local environment originating from dissolved or suspended bacterial components
(amino acids, metabolic by-products) as an attractive homing mechanism to find and
ingest the bacteria (Tanabe et al. 1980, Doughty and Dryl 1981, Snyder 1991).
Paramecia have been shown to be naturally infected with algae (Chlorella spp.) and
bacteria (Holospora obtusa, Caedibacter taeniospiralis) that eventually take up residence
inside digestive vacuoles or macronucleus and become symbionts (Gortz 1982,
Karakashian and Rudzinska 1981, Gortz 2001). We are unaware of reports of
mycobacteria in paramecia or other free-living ciliates.
The initial description of infective bacterial-paramecia mutual commensalism is
best illustrated by the role of Caedibacter taeniospiralis, a symbiont that provides the
host paramecium with resistance to elaborated toxins that kill adjacent uninfected
paramecia (Preer et al. 1974). These symbiotic relationships between paramecia, algae
and environmental bacteria may have originated as antagonistic pathogen-host
interactions that evolved to provide both paramecia and the nascent symbionts
mechanisms for adapting to novel environments, including entry into mammalian cells
108
and antibiotic resistance (Harb et al. 2000, Winiecka-Krusnell and Linder 2001).
Whereas not previously reported in ciliates, other studies have shown that
mycobacteria within amoebae, specifically the environmental amoebae Acanthamoeba
castellani, Acanthamoeba polyphaga and Dictyostelium discoideum, can successfully
survive, replicate, and may be potentiated for infectivity and spread of infection in
vertebrate hosts (Cirillo et al. 1997, Adekambi et al. 2006, Harriff et al. 2007, Hagedorn
et al. 2009, Salah et al. 2009).
The enhanced infectivity of mycobacteria residing in amoeba vectors has been
attributed to them acting somewhat as surrogate macrophages, with up-regulation of
virulence genes when mycobacteria are within the intracellular environment (Danelishvili
et al. 2004, Tenant & Bermudez 2006). Solomon et al. (2003), in concordance with this
observation, found that Mycobacterium marinum effectively replicates within
Dictyostelium by halting phagosomal maturation, and mechanisms of cellular invasion
elicited by M. marinum may be putatively conserved between amoebae and mammalian
macrophages (Hagedorn & Soldati 2007).
Phagolysosomal inhibition has been previously demonstrated in amoebae and mouse
macrophages infected with Mycobacterium avium and Mycobacterium tuberculosis
(Armstrong and Hart 1971, Thomas and McDonnell 2007). The mechanism responsible
for mycobacterial persistence within paramecia digestive vacuoles may be due to
abrogation of phagolysosomal fusion and subsequent maturation arrest of the
mycobacteria-containing vacuole due to combined elaboration of the ESX-1 type VII
secretion system (ESAT-6) and the mycobacterial mimicry of phosphatidylinositol,
109
stopping downstream actions of phosphoinositide 3-kinase (Philips 2008), ultimately
causing arrest of the vacuole in the early endosomal stage (MacGurn and Cox 2007).
In particular, ESAT-6 secretion by M. marinum allows membrane pore formation
to occur and facilitates vacuolar escape, with intracytoplasmic establishment of M.
marinum and subsequent cell-to cell spread (Smith et al. 2008). In our study, there was a
decrease of M. marinum in digestive vacuoles at 24 h post-ingestion, which then
remained relatively static over 1 week. Although numbers of M. marinum decreased
within digestive vacuoles over 1 week, many continued to persist inside vacuoles.
A reasonable explanation for this decline is that a subpopulation of infecting
mycobacteria were killed and processed during the initial phase of digestive vacuole
formation, with persisting mycobacteria upregulating key survival and pathogenicity
genes in a manner similar to Mycobacterium avium within amoebae and macrophages
(Danelishvili et al. 2004, Tenant and Bermudez 2006). Danelishvili et al. (2004)
discovered that common housekeeping genes involved in metabolism and protein
translation were upregulated, as were genes governing pathogenicity of mycobacteria
including nirB, lprC and dehydrogenases. This gene upregulation does not happen
immediately upon ingestion; rather, it occurs over days, from 24 to 48 hours after the
mycobacteria reside in digestive vacuoles and different genes are turned on nonsequentially. A similar pattern to M. avium was noticed with M. marinum inside digestive
vacuoles, with large numbers initially present and gradual loss over 24 h to 1 week,
probably reflecting the upregulation of survival and pathogenicity genes of persisting
mycobacteria, and perhaps as similar phenomenon with mycobacteria in other protozoa.
110
Another possibility is that mycobacteria are egested from digestive vacuoles back
into the environment. Based upon our observations that uninfected paramecia acquire
some mycobacteria within 48 hours when cohabitated with infected paramecia, vacuolar
egestion and reacquisition of environmental mycobacteria may also play a minor role in
the continued occurrence after initial exposure. In paramecia where mycobacteria
persisted for 1 week, we noticed that within 24 hours post-infection some mycobacteria
had escaped from the digestive vacuoles, relocated to the cytoplasm and had colonized
paramecia.
Phagosome escape and the corresponding cytoplasmic translocation have been
observed for other bacterial species that naturally infect paramecia and become
symbionts (Gortz 2001). It has been reported with pathogenic bacteria, notably
Legionella pneumophila residing in macrophages and amoebae (Molmeret et al. 2005)
and Mycobacterium marinum (Hagedorn and Soldati 2007, Cosson and Soldati 2008).
Mycobacterium marinum naturally infects environmental amoebae (Salah et al. 2009),
and now we have shown that it will also infect paramecia in a natural setting as well as
serving as a transmission vector for zebrafish mycobacteriosis (Peterson et al. 2013).
The gradual loss of paramecia over 1 wk could also be associated with simple
attrition. It is possible that infection with M. marinum eventually causes lysis of
paramecia, as reported by Krishna Prasad and Gupta in 1978. This could be caused by
intravacuolar accumulation of mycobacteria, replication and/or elaboration of a
mycobacterial toxin such as mycolactone. Cohabitation of M. marinum infected
paramecia with naïve cohorts resulted in low levels of cross-infection after 48 h and
111
demonstrates that mycobacterial transmission can occur between intact individual
paramecia.
Persistence of mycobacteria in paramecia digestive vacuoles and subsequent
spread to other cohort paramecia very likely results in enhanced mycobacterial
pathogenicity, similar to the discovery by Cirillo et al. (1997). They found that
Mycobacterium avium persisting within an amoeba, Acanthamoeba castellani, more
efficiently entered and replicated in both macrophages and epithelial cells. Paramecia
may become infected or spread mycobacterial infections by binary fission or conjugation
during reproduction. This may be similar to naturally occurring Holospora obtusa
macronucleus infection of paramecia, where the bacteria infect paramecia then are
released back into the environment to infect other paramecia (Gortz and Brigge 1998).
One concern in our study was whether intravacuolar GFP-labeled M. marinum
were viable. GFP fluorescence is regulated at the gene expression and hence, protein
level and thus bacteria lose fluorescence shortly after death. Based upon the previous
work of Lowder et al. (2000), where they demonstrated that dead GFP-tagged
Pseudomonas fluorescens lost their ability to fluoresce, we performed a similar
experiment to assess viability and subsequent fluorescence in GFP-tagged M. marinum.
Hydrochloric acid treatment of the M. marinum isolate at pH of 2.0 and 3.0, respectively,
for 10 minutes was performed in order to effectively mimic paramecia intravacuolar pH
post-phagolysosomal fusion, and bacteria completely lost fluorescence.
Our study is the first investigation on the interactions of mycobacteria in a ciliate.
We attempted to evaluate persistence in the paramecia by using bacterial culture, but
112
even with the use of selective media for mycobacteria and decontamination with cetylpyridinium chloride (CPC), overgrowth with Gram negative bacteria prevented accurate
counts of M. marinum. Therefore, future studies using axenic paramecia cultures (Fok
and Allen 1979), axenic cultures combined with selective antibiotic-resistant
mycobacteria, or a more defined culture media (i.e. media containing a single bacterial
species like Klebsiella sp.) are needed to allow for more precise evaluation of this
interaction. Determination of the actual mechanism of survival, e.g., prevention of
phagosome/lysosome fusion as seen with amoebae (Solomon et al. 2003), is also a next
step. As we showed in a previous study that both M. marinum and M. chelonae in
paramecia are more infective to zebrafish than mycobacteria from cultures (Peterson et
al. 2013), we plan to determine if similar mycobacterial genes are differentially regulated
within paramecia as seen in amoebae (Tenant and Bermudez 2006).
It is likely that the persistence of M. marinum within paramecia digestive vacuoles would
occur similarly with interactions of other mycobacterial species and other environmental
ciliates. For example, the environmentally complex bovine and ovine rumen contains
numerous ciliates, which by mass are the second-most populous rumen microbe
(Williams 1986). Rumen ciliates such as Epidinium, Entodinium, Diplodinium and others
establish symbiotic relationships with many rumen bacteria, and they may be predisposed
to harbor and potentially enhance virulence of Mycobacterium paratuberculosis, the
causative agent of Johne’s disease of cattle and sheep. Hence, investigation of this
potential phenomenon is also warranted.
113
ACKNOWLEDGEMENTS
The authors would like to thank Allison Creason in the Department of Botany and Plant
Pathology at Oregon State University for generously donating E. coli strain DH5α/DsRed
and the Zebrafish International Resource Center (ZIRC) for supplying paramecia
(Paramecium caudatum) for this experimental work. GFP-labeled M. marinum was
provided by Dr. Don Ennis in the Department of Biology at the University of LouisianaLafayette. ZIRC is supported by NIH/NCRR grant P40 RR012546. This work is
supported by NIH/NCRR grants T32 RR023917, R24 RR017886 and NIEHS Center
grant P30 ES000210.
114
Figure 4.1. Paramecia containing GFP-labeled Mycobacterium marinum. Figures 1, 3, 4,
6 were photographed with low-level brightfield and ultraviolet light (UV) to demonstrate
location of mycobacteria. Other figures are images photographed with UV only, used to
calculate number of pixels containing fluorescence using the ImagePro Plus program.
Insets in Figures 1 and 6 are high magnification of digestive vacuoles. Bars = 25 µm. Fig
1, 2 = time zero. Fig. 3 = 24 h post-exposure (pe). Fig., 4 and 5 = 72 h pe. In Fig. 5, 15 of
the 52 areas of specific fluorescence are labeled with their numerical value. Fig. 6 = 1 wk
pe.
115
Figure 4.2. Concentration of GFP-labeled Mycobacterium marinum in paramecia based
on area of fluorescence in digestive vacuoles from two trials. * = statistically different
compared to time zero (p < 0.01).
116
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Chapter 5. Discussion
Preface
The two objectives of my thesis research were to evaluate the ability to retrieve
DNA from zebrafish infected with clinically relevant mycobacterial species, and to
evaluate paramecia as a transmission vector for pathogenic mycobacteria in laboratory
zebrafish colonies.
Detecting mycobacterial pathogens with PCR from paraffin-embedded zebrafish
tissues
Evaluation of DNA sequences has become a cornerstone in bacterial species
identification. This is certainly the case for Mycobacterium species, as they are often
fastidious, grow slowly and typically provide few culture-specific or biochemical traits
by conventional methods that facilitate species-level identification (Daniel 1990, Cousins
et al. 1992). It is often more useful to obtain sequences for diagnoses directly from
infected tissues, particularly for Mycobacterium species that are slow growing or require
specialized media. This approach has been used to diagnose mycobacteria infections
directly from frozen or fresh infected fish (Kaattari et al. 2005, Poort et al. 2006, Whipps
et al. 2003, Whipps et al. 2007). Often only formalin preserved tissues embedded in
paraffin are available, which has led to attempts to obtain mycobacterial DNA from
paraffin-embedded tissues following observation of either chronic lesions (i.e.,
granulomas) or the presence of acid-fast bacteria in tissues that are indicative of
mycobacterial infection (Ghossein et al. 1992, Marchetti et al. 1998, Zink and Nerlich
2004, Miller et al. 1997, Pourahmad et al. 2009a,b).
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Fixatives used to preserve tissues (regardless of whether it is a coagulant or noncoagulant fixative) and the total time the tissues are held in a fixative can result in
degradation of DNA and RNA within tissues (Dubeau et al. 1986, Fiallo et al. 1992, Foss
et al. 1994). Exposure of mycobacterial DNA to formalin causes production of Schiff
bases on the free amino groups of nucleotides (Fraenkel-Conrat 1954, Dubeau et al.
1986), and subsequent cross-linking between tissue proteins and DNA (Jackson and
Chalkey 1974). Therefore, time in formalin is considered to be one of the most important
criteria for successful DNA retrieval and PCR amplification following tissue processing
for histology (Ben-Ezra et al. 1991, Greer et al. 1991). Once the tissues are embedded in
paraffin wax, this degradation process is slowed and subsequent amplification of short
DNA sequences is successful (Shibata et al. 1988). Fish are often preserved at a research
laboratory, aquaculture facility or in the field, and then shipped many days later to
another laboratory for histology processing.
Hence, I was particularly interested in the effects of time in fixative. Positive
results were consistently obtained with fish held in fixative up to 21 days, and the same
occurred at 45 days with the exception of one group. This positive result at the later time
points was somewhat surprising. Mycobacterium spp. has a unique, waxy cell wall
comprised of lipids and fatty acids (Kolattukudy et al. 1997), which may exclude
aldehydes to some extent. Perhaps this characteristic plays a role in preserving DNA
more than in non-acid fast bacteria exposed to formalin-based fixatives.
Dietrich’s, Davidson’s and Bouin’s fixatives are commonly used with fish tissues.
These contain acid and alcohol, both which cause denaturation and coagulation of
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proteins and nucleic acids within tissue specimens by dehydration and disruption of
electrostatic and hydrogen bonding (Fournie et al. 2000, Eltoum et al. 2001). Tissues
preserved in Bouin’s solution (which contains both picric and acetic acid) are particularly
problematic for DNA retrieval (Greer et al. 1991). Interestingly, there was no reduction
in positive samples with Dietrich’s compared to 10% buffered formalin.
Some samples were negative in both fixatives at some of the early time points.
The inability to recover DNA could logically be influenced by the total amount of
bacterial DNA present in the sample(s). Incorporation of a bead beating enhances
retrieval of DNA from formalin-preserved samples to liberate more DNA (Tripathi and
Stevenson 2012). Bead beating was used with the PCR protocol because from previous
experiences it was found to consistently yield more positive results with PCR from
paraffin-embedded tissues. Nevertheless, there was no significance in the ability to
detect mycobacterial DNA based on severity of infection. Mycobacterium marinum
causes more severe infections than M. chelonae in zebrafish (Watral and Kent 2007,
Ostland et al.2008, Whipps et al. 2008). In my study, the M. marinum strain was highly
virulent and several fish became moribund or died over the 8 week period, and these fish
were included in 45 day samples. In contrast, all fish injected with M. chelonae became
infected, but none exhibited morbidity or mortality. The M. marinum-infected fish had
three times the number of granulomas with acid-fast bacteria, but the ability of retrieving
DNA was not different than with fish infected with M. chelonae. Moreover, between
mycobacterial species I found no correlation of successful retrieval of DNA with the
severity of infection. Most zebrafish cases submitted to the ZIRC and our laboratory for
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health screening and diagnosis consist of multiple fish, and usually they are processed
and embedded within 3 weeks. Hence, even with only about 50-70% ability to obtain
mycobacterial DNA, diagnosticians can confidently use PCR on routinely processed and
paraffin-embedded zebrafish to identify these infections to the species level for many of
the fish within a particular case. The ability to identify mycobacteria from zebrafish to
the species level is important as the severity and distribution of various Mycobacterium
spp. are quite different. Therefore, fish health managers and clinicians with species
identifications in hand will be able to make more informed management decisions. For
example, facilities with M. haemophilum infected zebrafish often euthanize infected
populations, disinfect the aquaria, etc., and repopulate (Kent et al. 2009, Kent et al. 2011,
Whipps et al. 2012), and this approach is similarly recommended for M. marinum
outbreaks. In contrast, M. chelonae infections are presently managed by cleaning tanks
more frequently (Murray et al. 2011) or using different wild type strains of zebrafish
(Whipps et al. 2008). Due in part to my research, now at least two veterinary diagnostic
laboratories in United States are routinely receiving requests to identify mycobacteria in
zebrafish in paraffin blocks.
Identification of mycobacterial infections from formalin fixed paraffin-embedded
(FFPE) zebrafish can be further improved. My research (Chapter 2) used paraffin scrolls
for PCR amplification. Acid-fast bacteria are frequently observed in the intestinal lumen
of zebrafish, with and without infections in the visceral organs. Therefore, a concern
with amplification of DNA using whole fish sections may be confusion of mycobacteria
in granulomas versus intraintestinal mycobacteria that may be associated with other
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constitutive microbiota in the intestinal tract. A solution for the potentially confounding
problem of intraintestinal bacterial contaminants would be to use histologic sections as a
template to guide in the removal of specific cores from the tissue block that correspond to
confirmed areas of infected organs (Sfanos et al. 2008) or, even more precisely, by using
laser capture microdissection to select individual intralesional granulomas containing
mycobacteria for PCR assays (Ryan et al. 2002, Zhu et al. 2003, Selva et al. 2004). Both
methods would allow for accurate selection of infected tissue sites within the infected
zebrafish prior to PCR analysis for mycobacterial identification and PCR primer-specific
molecular identification of Mycobacterium spp.
Unlocking the mystery of mycobacterial transmission in zebrafish
The second area of my thesis research investigated natural transmission of
mycobacteriosis in fish using the zebrafish model (Chapters 3 and 4). Mycobacteria in
fish can be readily transmitted by feeding on infected tissues, but placing the fish into
aqueous suspensions of mycobacteria (even high concentrations) is not effective.
Previous studies have shown that mycobacteria within amoebae, specifically the
environmental amoebae Acanthamoeba castellani, Acanthamoeba polyphaga and
Dictyostelium discoideum can successfully survive, replicate, may be potentiated for
infectivity and spread of infection in the host (Cirillo et al. 1997, Adekambi et al. 2006,
Hagedorn et al. 2009). Mycobacteria from granulomas are more infective than those
from media cultures (Li et al. 2002). The enhanced infectivity of mycobacteria residing in
amoeba vectors has been attributed to them acting somewhat as surrogate macrophages,
124
with up-regulation of virulence genes when mycobacteria are within the intracellular
environment (Danelishvili et al. 2004, Tenant & Bermudez 2006). Solomon et al. (2003),
in concordance with this observation, found that Mycobacterium marinum effectively
replicates within Dictyostelium by halting phagosomal maturation, and that mechanisms
of cellular invasion elicited by M. marinum may be putatively conserved between
amoebae and mammalian macrophages (Hagedorn & Soldati 2007).
My research demonstrated for the first time that a ciliate protozoan, Paramecium
caudatum, like amoebae, can be a transmission vector for mycobacteria as mycobacterial
infections occurred by feeding paramecia containing either M. marinum or M. chelonae.
Moreover, transmission was dramatically increased with the paramecia vector compared
to fish fed mycobacteria alone. It is likely the prevalent infections with the ciliate was due
to a similar phenomenon of increased infectivity as reported with macrophages and
amoebae, as fish fed mycobacteria in the paramecia actually received much lower
amounts of bacteria compared to those fed the high doses of bacteria alone.
Mycobacterium species capable of causing disease in fish may occur in aquaria in
the absence of overt disease (Beran et al. 2006). In my study, all groups of fish, including
controls that were not deliberately exposed to mycobacteria, exhibited the presence
(perhaps colonization) of variable amounts of acid-fast bacteria in the intestinal lumen.
This demonstrates that the presence of mycobacteria, even if ingested, does not always
result in extraintestinal infections or may represent non-virulent intestinal microflora. As
discussed above, one factor allowing for these infections may be changes in gene
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expression as could occur whether the mycobacteria are in macrophages, amoebae or
invertebrate hosts such as paramecia.
Natural mycobacterial infections occasionally present as aerocystitis (swim
bladder infection) in zebrafish (Whipps et al. 2008). This presentation was observed in
larval fish, but not adults. Physostomous teleost fish, such as the zebrafish, have open
communication (pneumatic duct) between the distal oropharynx and swim bladder and
several mycobacterial species are known to colonize the swim bladder and cause
aerocystitis (Matthews 2004, Whipps et al. 2008, Kent et al. 2011). In larval zebrafish, it
may be anatomically favorable for non-motile mycobacteria to colonize the swim
bladder, as the total distance traveled from the oropharynx to pneumatic duct and swim
bladder is considerably shorter than in juvenile and adult zebrafish, and this may provide
one explanation for the predilection of mycobacteria to localize in the larval swim
bladder. Another possible explanation for the affinity of mycobacteria to the zebrafish
swim bladder may be a correlation between the higher cholesterol content of swim
bladder surfactant lipid, when compared to mammalian lung surfactant, and the
requirement of mycobacterial species to import and catabolize cholesterol to maintain
persistent infection in the host (Orgeig et al. 2003, Pandey & Sassetti 2008). I have
provided histologic interpretations for numerous diagnostic cases submitted to the
Zebrafish International Resource Center diagnostic program, Eugene, Oregon.
As reported by Whipps et al. (2008), I have observed several confirmed cases of
mycobacterial aerocystitis in zebrafish with in many laboratories. Perhaps these represent
126
fish with very chronic infections that were established as larvae or juvenile fish. The
teleost swim bladder shares many conserved anatomical features with the mammalian
lung, from comparative ontogeny (Perry et al. 2004) to conserved gene expression and
transcriptomics in molecular developmental pathways (Winata et al. 2009, Zheng et al.
2011), and thus the model that I developed with larval zebrafish may be a useful
surrogate for investigating certain aspects of human pulmonary tuberculosis.
Given that a wide variety of ciliate species occur in the aquatic environment and
undoubtedly encounter mycobacteria, further studies are warranted to compare and
contrast the inter-relationship of paramecia with mycobacteria to the amoebae models.
Aquatic ciliates, such as Paramecium caudatum, have evolved as natural predators of
environmental bacteria (Winiecka-Krusnell and Linder 2001). Paramecia track prey in
aquatic environments by chemosensory mechanisms that cause changes in membrane
fluidity and initiate both K+ and Ca2+ action potentials, using chemical cues in the local
environment originating from dissolved or suspended bacterial components (amino acids,
metabolic by-products) as an attractive homing mechanism to find and ingest the bacteria
(Tanabe et al. 1980, Doughty and Dryl 1981, Snyder 1991). Paramecia have been shown
to be naturally infected with algae (Chlorella spp.) and bacteria (Holospora obtusa,
Caedibacter taeniospiralis) that eventually take up residence inside digestive vacuoles or
macronucleus and become symbionts (Gortz 1982, Karakashian and Rudzinska 1981,
Gortz 2001), but reports of mycobacteria in paramecia or other free-living ciliates are
unknown. The initial description of infective bacterial-paramecia mutual commensalism
is best illustrated by the role of Caedibacter taeniospiralis, symbiont that provides the
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host paramecium with resistance to elaborated toxins that kill adjacent uninfected
paramecia (Preer et al. 1974). These symbiotic relationships between paramecia, algae
and environmental bacteria may have originated as antagonistic pathogen-host
interactions that evolved to provide both paramecia and the nascent symbionts
mechanisms for adapting to novel environments, including entry into mammalian cells
and antibiotic resistance (Harb et al. 2000, Winiecka-Krusnell and Linder 2001).
Whereas as not previously reported in ciliates, other studies have shown that
mycobacteria within amoebae, specifically the environmental amoebae Acanthamoeba
castellani, Acanthamoeba polyphaga and Dictyostelium discoideum, can successfully
survive, replicate, may be potentiated for infectivity and spread of infection in vertebrate
hosts (Cirillo et al. 1997, Adekambi et al. 2006, Hagedorn et al. 2009). Moreover,
mycobacteria in amoebae are more infectious to vertebrate hosts (Cirillo et al. 1997,
Harriff et al. 2007, Salah et al. 2009). The enhanced infectivity of mycobacteria residing
in amoeba vectors has been attributed to them acting somewhat as surrogate
macrophages, with up-regulation of virulence genes when mycobacteria are within the
intracellular environment (Danelishvili et al. 2004, Tenant & Bermudez 2006). Solomon
et al. (2003), in concordance with this observation, found that Mycobacterium marinum
effectively replicates within Dictyostelium by halting phagosomal maturation, and
mechanisms of cellular invasion elicited by M. marinum may be putatively conserved
between amoebae and mammalian macrophages (Hagedorn & Soldati 2007).
128
Phagolysosomal inhibition has been previously demonstrated in amoebae and
mouse macrophages infected with Mycobacterium avium and Mycobacterium
tuberculosis (Armstrong and Hart 1971, Thomas and McDonnell 2007). The mechanism
responsible for mycobacterial persistence within paramecia digestive vacuoles may be
due to abrogation of phagolysosomal fusion and subsequent maturation arrest of the
mycobacteria-containing vacuole due to combined elaboration of the ESX-1 type VII
secretion system (ESAT-6) and the mycobacterial mimicry of phosphatidylinositol,
stopping downstream actions of phosphoinositide 3-kinase (Philips 2008), ultimately
causing arrest of the vacuole in the early endosomal stage (MacGurn and Cox 2007). In
particular, ESAT-6 secretion by M. marinum allows membrane pore formation to occur
and facilitates vacuolar escape, with intracytoplasmic establishment of M. marinum and
subsequent cell-to cell spread (Smith et al. 2008).
I observed a decrease of M. marinum in digestive vacuoles at 24 h pe, which
remained relatively static over one week. Although numbers of M. marinum decreased
within digestive vacuoles over 1 wk, many continued to persist inside vacuoles. A
reasonable explanation for this decline is that a subpopulation of infecting mycobacteria
were killed and processed during the initial phase of digestive vacuole formation, with
persisting mycobacteria upregulating key survival and pathogenicity genes in a manner
similar to Mycobacterium avium within amoebae and macrophages (Danelishvili et al.
2004, Tenant and Bermudez 2006). Danelishvili et al. (2004) discovered that common
housekeeping genes involved in metabolism and protein translation were upregulated, as
were genes governing pathogenicity of mycobacteria including nirB, lprC and
129
dehydrogenases. This gene upregulation does not happen immediately upon ingestion;
rather, it occurs over days, from 24 to 48 hours after the mycobacteria reside in digestive
vacuoles and different genes are turned on non-sequentially. A similar pattern to M.
avium was observed with M. marinum inside digestive vacuoles of paramecia, with large
numbers initially present and gradual loss over 24 hours to 1 week, probably reflecting
the upregulation of survival and pathogenicity genes of persisting mycobacteria, and
perhaps as similar phenomenon with mycobacteria occurs in other protozoa.
Another possibility is that mycobacteria are egested from digestive vacuoles back
into the environment. Based upon our observations that uninfected paramecia acquire
some mycobacteria within 48 hours when cohabitated with infected paramecia, vacuolar
egestion and reacquisition of environmental mycobacteria may also play a role in the
continued occurrence after initial exposure. In paramecia where mycobacteria persisted
for one week, I noticed that within 24 hours post-infection some mycobacteria had
escaped from the digestive vacuoles, relocated to the cytoplasm and had colonized
paramecia.
Phagosome escape and the corresponding cytoplasmic translocation has been
observed for other bacterial species that naturally infect paramecia and become
symbionts (Gortz 2001) and has been reported for pathogenic bacteria, notably
Legionella pneumophila residing in macrophages and amoebae (Molmeret et al. 2005).
Mycobacterium marinum naturally infects environmental amoebae (Hagedorn and Soldati
2007; Cosson and Soldati 2008; Salah et al. 2009), and I have shown that it will also
130
infects paramecia in a natural setting, as well as serving as a transmission vector for
zebrafish mycobacteriosis (Chapter 3).
The gradual loss of paramecia over one week could also be associated with simple
attrition. It is possible that infection with M. marinum eventually causes lysis of
paramecia, as reported by Krishna Prasad and Gupta (1978). This could be caused by
intravacuolar accumulation of mycobacteria, replication and/or elaboration of a
mycobacterial toxin such as mycolactone. Cohabitation of M. marinum infected
paramecia with naïve cohorts resulted in low levels of cross-infection after 48 h and
demonstrates that mycobacterial transmission can occur between intact individual
paramecia. The persistence in paramecia digestive vacuoles and spread to other cohort
paramecia very likely results in enhanced mycobacterial pathogenicity, similar to the
discovery by Cirillo et al. (1997) that Mycobacterium avium persisting within an amoeba,
Acanthamoeba castellani more efficiently entered and replicated in both macrophages
and epithelial cells. Paramecia may become infected or spread mycobacterial infections
by binary fission or conjugation during reproduction. This may be similar to naturally
occurring Holospora obtusa macronucleus infection of paramecia, where the bacteria
infect paramecia then are released back into the environment to infect other paramecia
(Gortz and Brigge 1998).
One concern in my study was whether intravacuolar GFP-labeled M. marinum
GFP were alive. The fluorescence is regulated at the gene expression and hence, protein
level and thus bacteria lose fluorescence shortly after death. Based upon the previous
131
work of Lowder et al. in (2000), where they demonstrated that dead GFP-tagged
Pseudomonas fluorescens lost their ability to fluoresce, a similar experiment was
performed to assess viability and subsequent fluorescence in GFP-tagged M. marinum.
Hydrochloric acid treatment of our M. marinum isolate at pH of 2.0 and 3.0, respectively,
for 10 minutes was performed in order to effectively mimic paramecia intravacuolar pH
post-phagolysosomal fusion, and bacteria completely lost fluorescence.
It is paramount to those working in zebrafish health management and husbandry
to realize that paramecia, and perhaps other live feeds, can transmit mycobacteria to
larval and adult fish alike. During their first five to seven days post-hatching, zebrafish
larvae are fed an assortment of live microinvertebrates, including Paramecium caudatum,
as their primary food source. Paramecium spp. is a well-studied bacterivorous ciliate
protozoan that inhabits a wide range of freshwater habitats, serving a dual role as a
biological filter feeder and as a food item for other raptorial ciliate filter feeders,
copepods and larval fishes. Although prepared diets are the major components of the
juvenile and adult zebrafish diet (McClure et al. 2006; Harper and Lawrence 2011),
aquatic ciliates play a key role in the diet of larval zebrafish. Zebrafish larvae display a
“pause-travel” ambush predation strategy similar to that of the white crappie larvae
(Pomoxis annularis) (Browman and O’Brien 1992), and the majority of fish larvae are
opportunistic visual predators (Stenson 1985, Lazzaro 1987). Visual tracking of prey by
larval fish apparently supersedes both electro- and mechanosensory prey detection and
capture (Hairston et al. 1982). Lair et al. (1994) discovered that the larval guppy (Lebistes
reticularis) consumes up to 1,450 paramecia per hour, depending on paramecia
132
concentration, with an eventual plateau and diminishing of ingestion of paramecia over
time.
My study is the first investigation on the interactions of mycobacteria in a ciliate
(Chapter 4). I attempted to evaluate persistence in the paramecia by using bacterial
culture, but even with the use of selective media for mycobacteria and pretreatment with
cetylpyridinium chloride (CPC), overgrowth with Gram negative bacteria prevented
accurate counts of M. marinum. Therefore, future studies using axenic paramecia
cultures (Fok and Allen 1979), axenic cultures combined with selective antibioticresistant mycobacteria or a more defined culture media (i.e. media containing a single
bacterial species like Klebsiella sp.) are needed to allow for more precise evaluation of
this interaction. Determination of the actual mechanism of survival, e.g., prevention of
phagosome/lysosome fusion as seen with amoebae (Solomon et al. 2003), is also a next
step. As I showed in Chapter 3 that both M. marinum and M. chelonae in paramecia are
more infective to zebrafish that mycobacteria from cultures, determination whether
mycobacterial genes are up or down regulated within paramecia as seen in amoebae
(Tenant and Bermudez 2006) is warranted.
Amoebae and ciliates free in the environment can be vectors for mycobacteria, feeding on
these organisms, which naturally occur in aquaria, are likely an important natural route of
transmission and subsequent infection in zebrafish. Moreover, live foods such as aquatic
protozoans, worms and crustacean larvae are deliberately included as a dietary staple of
both larval and juvenile zebrafish (Watanabe et al. 1983, Lawrence 2007, Best et al.
133
2010). Most commonly, these foods consist of prey items such as paramecia, rotifers,
water fleas (Daphnia spp.), Artemia spp. and chironomid insect larvae. Water fleas in
particular have been implicated as a source of mycobacteria within established
aquaculture systems (Grange 1985, Conroy & Conroy 1999, Somsiri et al. 2005).
Considering that paramecia are a common first feed for larval zebrafish, the
results shown in this study demonstrate one specific potential source of infection.
Therefore, facility staff involved in propagating zooplankton, such as paramecia, as a
food source for larval zebrafish should exercise caution to avoid exposing these cultures
to potential sources of mycobacteria. Future mycobacteria transmission studies with live
feeds commonly used in laboratory zebrafish larviculture, including rotifers and Artemia
spp., are warranted to determine whether these microinvertebrates can also act as vectors
for mycobacterial infection within zebrafish colonies.
My research has implications beyond zebrafish and paramecia. It is likely that the
persistence of M. marinum within paramecia digestive vacuoles would occur similarly
with interactions of other mycobacterial species and other environmental ciliates. For
example, the environmentally complex bovine rumen contains numerous ciliated
protozoa, which by mass are the second-most populous rumen microbe (Williams 1986).
Rumen ciliates such as Epidinium, Entodinium, Diplodinium and others establish
symbiotic relationships with many rumen bacteria and they may be predisposed to harbor
and potentially enhance virulence of Mycobacterium paratuberculosis, the causative
134
agent of Johne’s disease in cattle and sheep. Investigation of this potential phenomenon is
also warranted.
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Chapter 6. Summary and Conclusions
The aims of this thesis were to develop a better and more reliable method to diagnose
mycobacteria commonly found in zebrafish to the species level and to define how
transmission of mycobacteria occurs in laboratory colonies of zebrafish. Because
mycobacteriosis is so endemic and problematic in not only zebrafish colonies but in all
commercially reared fishes, these two aims were logical and worthwhile pursuits.
Mycobacterium marinum and M. haemophilum are frequently associated with high
mortality in the colony setting, whereas the ubiquitous M. chelonae is more wide spread
and causes minimal mortality but high morbidity. This ultimately leads to potentially
distracting and confounding results in the data accrued by researchers using zebrafish.
Although I say “potentially”, there is no doubt that this is already happening in the
research community. The next obvious launch point for further investigation would be to
more fully assess how subclinical mycobacterial infections compromise the quality of
research. This is precisely why the identification of mycobacterial infections to the
species level are absolutely necessary, in order to provide important information for
making colony-level health management decisions. The current method of observing
acid-fast bacilli, which may not be mycobacteria, in histological sections or tissue
imprints is the gold standard, but this only allows for a putative diagnosis and provides
very limited epidemiologic data of value. Mycobacterial DNA of diagnostic value can be
retrieved from paraffin blocks. Type of fixative, time in fixative before processing,
species of mycobacteria, and severity of infection had no bearing on the ability to
successfully detect mycobacteria in these samples. The PCR assay was quite effective,
143
and obtained PCR product from 75% and 88% of the M. chelonae and M. marinum
infected fish, respectively. Fixative type, time in fixative, and mycobacterial species
showed no statistical relationship with the efficacy of the PCR test.
Regarding natural transmission, zebrafish like all other fishes, are capable of
contracting mycobacterial infections by feeding on infected fish tissue but other natural
routes have not been firmly established. Mycobacteria within environmental amoebae act
as transmission vectors for mycobacteria and their virulence is enhanced, with
mycobacteria becoming primed for intracellular residence and cell to cell movement.
Paramecium caudatum are commonly used as the first food for larval zebrafish. The
ability of P. caudatum to transmit these mycobacteria to larval, juvenile and adult
zebrafish has been confirmed through my experimental work. Infections were defined by
granulomas containing acid-fast bacteria in extraintestinal locations, notably the target
filter organs (liver, spleen and kidney) normally associated with mycobacterial infections.
Zebrafish fed paramecia containing mycobacteria became infected at a higher incidence
than controls. Larvae (exposed at 4 days post hatch) fed paramecia with M. marinum
exhibited an incidence of 30% (24/80) and juveniles (exposed at 21 days post hatch)
showed 31% incidence (14/45). Adult fish fed gelatin diets containing bacteria within
paramecia or mycobacteria alone for 2 wk resulted in infections when examined 8 wk
after exposure: M. marinum OSU 214; in paramecia 47% (21/45; 3.5 x 105
dose/fish/day), M. marinum CH in paramecia 47% (9/19; 3.6 x 105 dose/fish/day), M.
chelonae in paramecia 38% (5/13; 3.5 x 105 dose/fish/day).
144
The ability of mycobacteria to persist within paramecia, which seems a requisite for
transmission, has been demonstrated in amoebae. To effectively and definitively show
that mycobacteria indeed behave differently than other environmental bacteria, the Gram
negative bacteria ingested by paramecia were processed within an hour. In contrast, GFPlabeled Mycobacterium marinum, which has attenuated virulence compared to wild-type
M. marinum (i.e. the OSU 214 strain), unequivocally persist over several days within
paramecia digestive vacuoles. The concentration of M. marinum at 1 hour was similar to
that at the time of ingestion. Twenty-four hours post-ingestion and later there was
significant decline in M. marinum concentrations compared to time of ingestion, but M.
marinum continued to persist inside digestive vacuoles for up to one week. Together with
my previous transmission studies, these results clearly demonstrate for the first time, with
no doubt, that Paramecium caudatum acts as a vector for mycobacteria. This provides a
useful animal model for evaluation of natural mycobacterial infections, a stepping stone
for broader studies in other fish species where paramecia is used as a larval fish food, and
demonstrates the real possibility of mycobacterial transmission in zebrafish facilities via
contaminated paramecia cultures.
145
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APPENDICES
171
Appendix A
The Luna stain, an improved selective stain for detection of microsporidian spores
in histologic sections
Tracy S. Peterson1, Jan M. Spitsbergen1, Stephen W. Feist3 and Michael L. Kent1, 2
Diseases of Aquatic Organisms, 2011, 95:175-180
1
Department of Microbiology, 220 Nash Hall, Oregon State University, Corvallis, OR
97331
2
Department of Biomedical Sciences, Oregon State University, Corvallis, OR 97331
3
Centre for Environment Fisheries and Aquaculture Science (Cefas), Weymouth
Laboratory, Barrack Road, The Nothe Weymouth, Dorset DT4 8UB. UK
172
ABSTRACT
Microsporidia in histologic sections are most often diagnosed by observing spores
in host tissues. Spores are easy to identify when they occur in large aggregates or
xenomas when sections are stained with hematoxylin and eosin (H&E). However,
individual spores are not frequently detected in host tissues with conventional H&E
staining, particularly if spores are scattered within the tissues, areas of inflammation or
small spores in nuclei (i.e., Nucleospora salmonis). Hence, a variety of selective stains
that enhance visualization of spores are recommended. We discovered that the Luna
stain, used to highlight eosinophils, red blood cells and chitin in arthropods and other
invertebrates, also stains spores of Pseudoloma neurophilia. We compared this stain to
the Gram, Fite’s acid fast, Giemsa, and H&E stains on eight other aquatic microsporidian
organisms that were readily available in our two laboratories: Loma salmonae, Glugea
anomala, Pseudoloma neurophilia, Pleistophora hyphessobryconis, Pleistophora
vermiformis, Glugea sp., Steinhausia mytilovum and an unidentified microsporidian from
E. sinensis, UK. Based on tinctorial properties and background staining, the Luna stain
performed better for detection of 7 of the 8 microsporidia. Gram stain was superior for
the two microsporidia from invertebrates, Steinhausia mytilovum and the unidentified
microsporidian from E. sinensis.
INTRODUCTION
Microsporidian parasites are a reductionist group of eukaryotic obligate
intracellular parasites related to the fungi that infect numerous different invertebrates and
vertebrates, and infections are notably more prevalent in both arthropods and teleost
173
fishes (Keeling and Fast 2002). Morphological diagnosis of these infections is usually
based on visualization of the spore stage.
Aside from this approach, in almost all
diagnostic cases the detection of microsporidia relies upon the host organism’s infectious
burden or obvious xenoma formation (Kotler et al. 1994). Spores are usually detected in
wet tissue preparations, histological slides or tissue/fecal smears. One of the problems
encountered in routine H&E sections is that spores often are not stained and thus can be
obscured against a similarly staining background. For this reason, a litany of selective
stains have been employed to facilitate and enhance detection of spores in tissue
sections, which includes the Brown-Brenn and Brown-Hopps modified Gram, Gram
chromotrope, Giemsa, Masson’s trichrome, Periodic-acid Schiff’s, Acridine orange,
Gomori’s methenamine silver, Warthin-Starry, Ziehl-Nielsen and Fite’s acid-fast stains
(Weber et al. 1999, Lamps et al. 1997, Garcia 2002 ). Many of the selective stains used
to detect microsporidial organisms often give variable and inconsistent staining (Lamps
et al. 1997, Garcia 2002). Fluorescent stains, such as the Calcofluor White and
FungiFluorTM , are also useful for detecting spores in smears or sections (Garcia 2002,
Guzman et al. 2001, Kent and Bishop-Stewart JK 2003). Major difficulties encountered
in the interpretation of fluorescent stains include distinguishing microsporidial spores
from background, artifactual staining and determining the precise location of spores in
tissue sections. PCR tests are also available for several microsporidia including those
infecting fishes (Brown and Kent 2002, Whipps and Kent 2006). These tests have been
adapted for in situ hybridization on tissue sections (Sanchez et al. 1999). Each of these
selective stains has specific utility for detection of spores, including ease-of-use and
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interpretation, time and economy, degree of contrast with tissue elements and specificity,
but also drawbacks for routine use, for example high background “noise”, variable
staining, requirements for a fluorescent microscope or non-confirmatory, time-consuming
and expensive reagents.
The Luna stain was originally developed to detect cytoplasmic granules within
eosinophils, Negri bodies, erythrocytes and phagocytes (Luna 1968, Tomasi et al. 2008).
It also has been used to demonstrate elastin (Kligman 1981). We discovered that spores
of Pseudoloma neurophilia from zebrafish stain positive (brick red) with the Luna stain
and have minimal background interference. We then evaluated several other
microsporidia with this stain, comparing it with Fite’s acid fast, Giemsa and Gram stains
which we have routinely used for the detection of microsporidia in our laboratories.
MATERIALS AND METHODS
Histologic samples representing the 8 different microsporidia were obtained from
the case files of the Zebrafish International Resource Center (Eugene, OR), Oregon State
University Department of Microbiology (Corvallis, OR), University of California-Davis
School of Veterinary Medicine (Davis, CA) and the Centre for Environment, Fisheries
and Aquaculture Science (Weymouth Laboratory, UK). The histologic samples were
comprised of six different teleost fishes, Mitten crab (Eriocheir sinensis) and a mussel
(Mytilus sp.) that had been previously diagnosed with microsporidian infections. 10%
neutral buffered formalin and Dietrich’s fixative were used for preservation of fish tissues
and sea water Davidson’s fixative for mussel and mitten crab. Five serial sections, cut at
175
4 µm and deparaffinized, were made from each of the selected tissue blocks. These tissue
sections were then stained with H&E and a special stain panel of Luna, Gram, Fite’s
acid-fast and Giemsa stains. Luna and Gram stains were performed using Luna’s method
for erythrocytes and eosinophil granules (Luna 1968 pages 111-112) and the AccustainTM
Gram stain for tissue kit (HT90T, Sigma-Aldrich). Fite’s acid-fast and Giemsa stains
were employed using Fite’s method for acid fast organisms (Luna 1968 pages 217-218)
and May-Grunwald Giemsa method (Luna 1968 pages 121-122). Slides were evaluated
for routine histologic features, presence of microsporidian organisms, detection of
microsporidian spores by special stains, fidelity of staining for each of the special stains,
amount of background stain and artifacts.
RESULTS
Luna stain
The majority (approximately > 90%) of microsporidial spores in 7 of 8 cases,
regardless of genus or species, stained brick red and were easily detected in tissue
sections. The spores displayed exceptionally high contrast from the background stain
with minimal to no interference from the few positively-staining tissue elements
(Appendix Table 1.1, Appendix Figs.1.1 and 1.2). Spores were particularly evident in
cases of diffuse multiorgan infections. With Nucleospora salmonis, a few small
intranuclear spores were detected which stained deep brick red in contrast to blue-black
stained nuclei (Appendix Fig. 1.2i). Spores of Steinhausia mytilovum from mussel
oocytes did not stain, while the unidentified microsporidian from mitten crab enterocytes
176
showed variable staining. With fish, other tissues that also stained red included
erythrocytes, bone, lens of the eye, and eosinophilic granular cells of salmon.
Gram stain
All microsporidial spores tested with the Gram stain showed approximately 50 to
75 % positive staining (defined as staining Gram positive), with many individual spores
having a high degree of variable staining and in some cases, almost no positive staining.
The background for this stain is yellow and contrasts well with the deep blue to purple
color of spores. For most microsporidia, spores were easily distinguished from the
surrounding tissue elements. However, the Gram stain was much less sensitive for
Nucleospora salmonis as the nuclear chromatin also stained deep blue to purple
(Appendix Fig. 1.2h). Steinhausia mytilovum spores were variable with this stain, but
some aggregates were distinctly Gram positive. Spores of the unidentified
microsporidian from E. sinensis were intensely positive (deep blue). Gram stains of testes
from zebrafish revealed variably strong dark blue to purple staining of intracystic
spermatids.
Fite’s acid-fast stain
There was extensive variability among microsporidial spore staining, from no
spores staining positive to approximately 25-50% of spores in any given tissue section
staining positive. In many of the tissue sections the spores were incompletely stained with
minimal background contrast or did not stain positive at all.
177
Giemsa stain
Spores consistently stained blue to deep blue with this stain. However, there was
essentially no contrast with surrounding tissues as they stained the same color.
DISCUSSION
Microsporidian infections in histologic sections are most often diagnosed by
observation of spores in host tissues. Spores are easy to identify when they occur in large
aggregates with sections stained with hematoxylin and eosin. However, spores are not
usually stained with either dye, and thus individual spores scattered in areas of
inflammation or small spores in nuclei (i.e., N. salmonis) are difficult to detect. Hence, a
variety of selective histologic stains that highlight spores are recommended.
Our study comparing the performance of four special stains on 8 different
microsporidia (6 from fish and 2 from aquatic invertebrates) demonstrated that the Luna
stain was optimal or at least comparable to the Gram stain for most specimens. Positive
background staining with the Luna stain was confined to erythrocytes, bone, the lens of
the eye and the chitin capsule of brine shrimp nauplii (within the gastrointestinal tract of
zebrafish), all of which tended to be variably stained. None of these specific tissues in the
hosts were infected by microsporidia. A novel finding with respect to teleost tissues was
the intense positive staining of eosinophilic granular cells, noted in the tissue sections of
salmonid gills. One key question is what microsporidian cellular components have an
affinity for the stain. The endospore wall of microsporidia contain chitin (Keeling and
Fast 2002, Vavra and Larsson 1999) and the active dye (1% Biebrich scarlet) for the
178
Luna stain specifically complexes with chitosaccharides (Holler et al. 1975).
Historically, Biebrich scarlet dye has been used to identify chitinous structures of
invertebrates (Joffe and Hepburn 1973) and this was confirmed in our study by the
positive staining of brine shrimp exoskeleton seen in the intestine of zebrafish. Chitin is a
long chain polymer of N-acetylglucosamine. A reasonable hypothesis is that the charged
sulphur trioxide of the dye binds with the nitrogen of the amine in chitin, as this is
accessible. Each dye molecule contains two charged sulphur trioxides so there is the
possibility that the two sulphur trioxides bind to the two adjacent amine functions on the
polymer chain, making the binding tighter. Therefore, we conclude that the Luna stain
stains chitin in microsporidian spores. The Gram stain was also very effective for staining
spores in most of the samples, with the dramatic differentiation between blue staining
spores and a brilliant yellow background. Indeed, we used this stain recently to identify
spores of Pleistophora hyphessobryconis (Sanders et al. 2010), and variations of the
Gram stain have been used by many pathologists to highlight a wide variety of
microsporidian spores in tissue sections (Rupstra et al. 1988, Moura et al. 1996). In fact,
this stain was superior to the Luna stain for the two invertebrate species. The Gram stain
forms an insoluble moiety with peptidoglycan rather than chitin. Spores that are Gram
positive but Luna negative may contain little chitin or are less fully developed. However,
the majority of infected enterocytes contained mature spores of the unidentified
microsporidian from E. sinensis.
One major drawback of the Gram stain was for detection of intranuclear spores of
Nucleospora salmonis as nuclear material also stained dark blue to deep purple. The
179
Warthin-Starry silver stain has been used to detect microsporidian spores and Kent et al.
(1995) previously used this stain for detecting spores of N. salmonis. Spores were
positive, but there was considerable nuclear background staining that frequently
confounded detection and interpretation of the intranuclear spores. Furthermore, we are
investigating vertical transmission of Pseudoloma neurophilia. The Gram stain is
effective for demonstrating these spores in zebrafish ovaries and eggs. Similar to the
Warthin-Starry stain, intense deep basophilic staining of nuclear material and developing
sperm makes detection and interpretation of spores difficult within testes.
Acid fast stains have been used for highlighting microsporidian spores and we
previously used the Ziehl-Nielsen stain to demonstrate spores of P. neurophilia in tissue
sections (Ramsay et al. 2010). In our experience with both the Ziehl-Nielsen and Fite’s
stains, we found that results were quite variable between laboratories and between runs,
largely based on the amount of decolorization as spores do not retain carbol fuschin stain
as well as mycobacteria. As positive staining of spores may be reversed by overdecolorizing, Ramsay et al. (2010) recommended reducing the time of this step.
However, this results in more background staining. In other words, the correct balance
between excessive background staining and over-decolorization of spores is required to
obtain optimal contrast between spores and surrounding tissues.
The Giemsa stain is commonly used to highlight protozoan and myxozoan
parasites and has been recommended for microsporidia (Rupstra et al. 1988). We found
that spores from all microsporidia species in our study did stain positive but could not be
180
readily distinguished from surrounding tissues. Therefore, the Giemsa stain would still
be better than H&E, which usually does not highlight spores well.
In conclusion, the Luna stain is superior to the Gram, acid fast, and Giemsa stains
for staining microsporidian spores with the least amount of background interference.
Moreover, it is a simple stain that involves only two principle dyes and is readily
available in most histology laboratories.
ACKNOWLEDGEMENTS
The authors wish to thank Virginia Watral, Jayde Ferguson, John Bignell, Justin
Sanders, Dr. Grant Stentiford, Dr. Katy Murray and Dr. Ronald Hedrick for contributing
case material and technical assistance for this paper. We would also like to thank Kristin
Berkenkamp and the Oregon State University Veterinary Diagnostic Laboratory for
excellent histotechnical assistance. We thank Dr. Stephen Irving for discussion on
chemistry of the Luna stain. This work was supported by NIH NCRR grants
5R24RR017386-02 and 1 T32 RR023917 and UK Department for Environment, Food
and Rural Affairs contract FB001 (SWF).
181
Appendix Figure 1.1. Microsporidia in histologic stains from various fishes. Bars = 25
µm. a. Pseudoloma neurophilia individual spores (arrow) and aggregate with
presporogonic stages (p) associated with chronic inflammation in the nerve and skeletal
muscle of Danio rerio. Luna. b – e. P. neurophilia xenoma in spinal cord of zebrafish.
b, Luna; c, Gram; d, Fite’s acid fast; e, Giemsa. f-i, Loma salmonae xenoma in
Oncorhynchus kistuch gill. f, Luna; g, Gram; h, Fite’s acid fast; i, Giemsa. j-m. Glugea
stephani spores, within background of chronic inflammation, in ovarian follicle of
Pleuronectes platessa. j, Luna; k, Gram; l, Fite’s acid fast; m, Giemsa.
182
Appendix Figure 1.2. a – m.
183
Appendix Figure 1.2. a – e. Pleistophora hyphessobryconis in skeletal muscle (a) and
skin (b – e) of Danio rerio. Bar = 25 µm. a, b. Luna; c, Gram; d, Fite’s acid fast; e,
Giemsa. e –g, unidentified microsporidian within enterocytes of Eriocheir sinensis. Bar
= 25 µm. e, Luna, Arrow = red staining aggregate of intra-enterocytic spores. f, Gram,
with numerous Gram-positive spore aggregates within enterocytes; g, Fite’s acid fast. h, i,
Nucleospora salmonis in nuclei of Chinook salmon leukocytes. h, Gram, with spore
(arrow) in deep blue staining nucleus. i, Luna. Note red staining spore in nucleus (arrow).
p = presporogonic stage within nucleus. j - k, Steinhausia mytilovum spores (arrows) in
oocyte nuclei from Mytilus edulis. Note blue staining of spores with Gram (j), whereas
spores do not stain red with Luna (k). Bar = 20 µm. l, m. Testis stained with Luna (l) or
Gram (m). Note with both stains spermatids stain dark blue to black. Bar = 100 µm.
184
Appendix Table 1.1. Characteristics of microsporidian spores in histological sections with special stains. + = most or all
spores were stained and distinct from background, +/- = variable, some spores stained, 0 = less than 10% of spores colored
with special stain or spores not clearly distinguished from background.
Microsporidia
Fixative
Host
Luna
Gram
Fite's Acid-Fast
Giemsa
Glugea anomala
Dietrich’s
+
+/-
+/-
+
Loma salmonae
+
+/-
+/-
+
Nucleospora salmonis
10% neutral buffered
formalin (10% NBF)
Davidson’s
+/-
0
0
0
Pseudoloma neurophilia
Dietrich’s/ 10% NBF
Gasterosterus
aculeatus
Oncorhynchus
kisutch
Oncorhynchus
tshawytscha
Danio rerio
+
+/-
+/-
+/-
Pleistophora
hyphessobryconis
Pleistophora vermiformis
Dietrich’s
Danio rerio
+
+/-
+/-
+/-
10% NBF
Cottus gobio
+
+
0
+
Glugea stephani
10% NBF
+
+
0
+/-
Steinhausia mytilovum
Sea water Davidson’s
Pleuronectes
platessa
Mytilus edulis
0
+/-
0
0
Unidentified
microsporidian
Sea water Davidson’s
Eriocheir sinensis
+/-
+
0
0
185
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187
Appendix B
Malignant Dysembryoplastic Neuroepithelial Tumour in a Zebrafish (Danio rerio)
T. S. Peterson*, J. R. Heidel‡, K. N. Murray§, J. L. Sanders*, W. I. Anderson¶
and M. L. Kent*, †
Journal of Comparative Pathology, 2013, 148:220-224
Department of Microbiology, †Department of Biomedical Sciences and ‡Veterinary
Diagnostic Laboratory, College of Veterinary Medicine, Oregon State University,
Corvallis, Oregon 97331
§
Zebrafish International Resource Center, University of Oregon, Eugene, Oregon 97403
and ¶28 Hobbs Road, Plattsburgh, NY 12901, USA
*
188
SUMMARY
Neuroectodermal tumours in man, including medulloblastoma,
medulloepithelioma, neuroblastoma, esthesioneuroblastoma, primitive neuroectodermal
tumour and dysembryoplastic neuroepithelial tumour, typically occur in children and
young adults. These tumour types are occasionally observed in juvenile and adult
zebrafish (Danio rerio), either as induced tumours in carcinogen-exposed zebrafish or as
an incidental finding in zebrafish > 2 years of age. An adult zebrafish submitted for
routine histological examination was sent for a second opinion consultation after an
uncharacteristic brain mass was identified. Microscopically, the expansile and infiltrative
extracortical mass arising from the cerebellum had a diffuse microcystic pattern with
solid hypercellular regions occupying 80% of the extrameningeal space and effacing the
endomeninx and significantly displacing the metencephalon. The mass was composed of
dense sheets of oligodendrocyte-like cells, random neurons and pseudocysts containing
‘floating neurons’ within a scant mucinous matrix. Neoplastic cells demonstrated
positive perinuclear and intracytoplasmic expression of S-100. Malignant
dysembryoplastic neuroepithelial tumour was diagnosed based upon the histologic
features of the brain mass, which were indistinguishable from the human tumour. To our
knowledge, this is the first report of a dysembryoplastic neuroepithelial tumour in a
zebrafish.
NOTE
Dysembryoplastic neuroepithelial tumour (DNET) is a benign brain tumour
arising from dysplastic brain development of the subpial granular layer (Daumas-Duport
189
et al. 1988, Cabiol et al. 1999, Adesina and Rauch 2010). The tumour affects children
and young adults, ranging in age from 9 to 17 years old at the time of diagnosis (DaumasDuport 1993, Raymond et al. 1994). Dysembryoplastic neuroepithelial tumour of man
has three recognized histological patterns: multinodular, nodular and diffuse, with most
of the tumour masses localized to the frontal and temporal lobes and occasional
occurrence in the parietal and occipital lobes (Hirose 1999, Honavar et al. 1999).
Originally described by Daumas-Duport et al. (1988), DNET is composed of invariant
specific glioneuronal elements (oligodendrocyte-like cells [OLCs], astrocytes and
neurons) with the OLCs forming the bulk of the tumour mass. Hallmark
histoarchitectural features of DNET are microcystic or alveolar patterns (Leung et al.
1994, Hirose 1999), with intracystic ‘floating neurons’ in a mucinous matrix and
hypercellular background of OLCs. Cortical dysplasia, mitotic figures, nuclear atypia,
calcification and necrosis tend to be variable among the histological types of DNET and
in many cases are found inconsistently (Daumas-Duport et al. 1999, Hirose 1999,
Honavar et al. 1999).
Naturally-occurring neuroectodermal tumours of the central nervous system
(CNS) in teleost fishes are uncommonly reported (Fournie and Overstreet 1985, Kagan et
al. 2010), probably due to a low prevalence among natural fish populations and few
opportunities to acquire specimens. In wild-caught and laboratory fishes, such as
medaka and zebrafish, both spontaneous and induced CNS neuroectodermal tumours
occur (Fournie and Overstreet 1985, Reimschuessel et al. 1989, White 2004, Feitsma et
al. 2008; Ju et al. 2009, Kagan et al. 2010). Neuroectodermal tumours of fish arise during
190
development from pluripotent progenitor cells within subependymal proliferation zones
of the telencephalon, tectum and cerebellum, similar to the mammalian subventricular
zone (Becker and Becker 2008). The main difference between neuroectodermal CNS
tumours in teleost fishes and those in mammals is that adult fish retain an embryonic
plasticity and regenerative capability of neuroepithelial cells that favour development of
such CNS tumours (Grandel et al. 2006, Tshering et al. 2008).
An adult wild-type male zebrafish from a sentinel population was reported to have focal
cutaneous ulceration on the head, ventrolateral swelling of the body and scoliosis. It was
submitted to the Zebrafish International Resource Center, Eugene, Oregon for routine
diagnostic screening. The zebrafish was determined to be an older adult based upon both
thymic involution and vertebral scoliotic changes in the absence of Pseudoloma
neurophilia spinal cord and nerve root infection; although a single isolated Pseudoloma
neurophilia pseudocyst containing mature spores was observed within a cranial nerve
subjacent to the tumour mass. Vertebral scoliosis is a common indicator of senescence in
zebrafish and other teleost fishes due to sarcopenic changes in the dorsal skeletal
musculature (Gerhard et al. 2002). The specimen was fixed whole in 10% neutral
buffered formalin (total time in fixative unknown), decalcified in CalExIITM solution (10–
15% formic acid plus formalin) for 48 h, trimmed parasagittally, processed routinely and
embedded in paraffin wax. Tissue sections were stained with haematoxylin and eosin
(HE) and a special stains panel including periodic acid–Schiff (PAS), mucicarmine and
luxol fast blue (LFB).
191
Microscopically, the brain mass revealed a poorly demarcated, expansile and
infiltrative extracortical neoplastic mass occupying the endomeningeal space and
emanating from a solitary dysplastic cortical focus within the cerebellar external granular
cell layer containing dysmorphic neurons and putative ‘balloon’ cells, for which specific
diagnostic criteria have been described (Palmini et al. 2004). The brain mass extended
into the retrobulbar space adjacent to the choroid rete, around osseous bullae of the inner
ears and dorsally along the brain stem and spinal cord. Invasive sheets of tumour cells
penetrated through the exoccipital and basioccipital bones of the neurocranium,
extending into and dissecting individual skeletal muscle fibre bundles. At low
magnification the mass had a diffuse microcystic appearance with a densely cellular
background similar in appearance to the brain neuropil (Appendix Fig. 2.1). Within the
intracystic spaces there were peripheralized and infrequent central ‘floating neurons’ of
variable sizes surrounded by a sparsely mucinous matrix (Appendix Fig. 2.2). Solid areas
of the mass were composed of hypercellular regions of OLCs with round hyperchromatic
nuclei, inconspicuous nucleoli and minimal to inapparent cytoplasm, often with a
perinuclear halo or ‘fried egg’ appearance reminiscent of those seen in an
oligodendroglioma. Many of the pseudocysts appeared to be lined by variable numbers
of these OLCs. Dysmorphic neurons, rare binucleated neurons and putative balloon cells
were observed throughout the mass. OLCs occasionally displayed moderate nuclear
pleomorphism and karyomegaly. Mitotic figures, perineuronal satellitosis, perivascular
aggregates of OLCs, glomeruloid vascular proliferation and necrosis were not seen in any
areas of the tumour. The LFB stain highlighted intratumoural neurons and neurons in the
192
focal area of cerebellar cortical dysplasia. Misshapen and abnormally oriented neurons
(interpreted as dysmorphic neurons), as well as putative balloon cells, were identified in
the area of cortical dysplasia (Appendix Fig. 2.3). Mucicarmine staining revealed lightly
positive staining of the intracystic mucinous material when it was present. The PAS stain
demonstrated sparse intracystic PAS-positive mucofibrillar material. S-100
immunohistochemical labeling of the tumour cells was inconsistent, with punctate
perinuclear and diffuse intracytoplasmic labeling of individual OLCs. Glial cells serving
as an internal positive control also had inconsistent labeling.
Additional findings included a focal non-ulcerated integumentary defect devoid of
scales on the craniodorsal calvarium (most likely the ‘parietal eye’) and a spermatocytic
seminoma. A diagnosis of malignant DNET was made because the histomorphology and
cellular components of the zebrafish brain tumour resembled the diffuse form of human
DNET and closely paralleled the established human diagnostic criteria, including those
for the rare malignant version of DNET (Hammond et al. 2000). Features of malignancy
in the zebrafish DNET included OLC cellular atypia and an infiltrative growth pattern.
Malignant transformation of DNET has only been reported twice in the human medical
literature (Hammond et al. 2000, Rushing et al. 2003).
DNET and other glioneuronal tumours arise from pluripotent neuroglia (i.e.
oligodendrocyte-like or oligodendroglial cells; Pal et al. 2002) that are retained in
zebrafish and other teleost fishes throughout development (Ekstrom et al. 2001).
Neuroglia originates from specific proliferation and migratory zones in several brain
areas including ependymal and subependymal regions of telencephalic and tectal
193
ventricles. Neuroglia that differentiate into oligodendroglial cells in the zebrafish brain
initially appear and mature in the ventral hindbrain (metencephalon) by 4 days post
fertilization (dpf) and then extend rostrally to the midbrain (mesencephalon) at 10 dpf
(Wullimann and Knipp 2000, Brosamle and Halpern 2002, Yoshida and Macklin 2005).
Within cerebellar subdivisions, neuroglia in the valvula and corpus cerebelli continuously
migrates to respective granular layers, differentiating into either neurons or
oligodendrocytes (Zupanc et al. 2005). Given the anatomical location (the cerebellar
external granular layer) the DNET in this zebrafish may have derived from this
population of embryonic neuroglia during early development and developed slowly over
time. S-100 protein was selected as the immunohistochemical marker to identify
intratumoural OLCs based on the work of Hirose et al. (1994), which showed most
intratumoural OLCs to be S-100 positive. In human DNETs, OLC S-100 reactivity was
reported in the nucleus and cytoplasm, but neurons were weakly reactive or negative
(Hirose et al. 1994, Gyure et al. 2000). In the zebrafish brain tumour, patchy S-100
expression was observed in the perinuclear and intracytoplasmic regions of OLCs.
Labeling variation of the S-100 zebrafish OLCs may be attributable to using unvalidated
mammalian S-100 antibody (1 in 400 dilution; rabbit anti-bovine S-100;
DakoCytomation, Dako, Carpinteria, California, USA), as mammalian antibodies
produce inconsistent results when applied to fish tissues (Kagan et al. 2010), or because
tumour cells were not differentiated enough to allow expression of S-100 protein. The
lack of strong staining by either mucicarmine or PAS is most likely due to loss of
mucinous material during tissue processing.
194
Neuroectodermal tumours in fish are caused by heritable genetic defects and
exposure of early life stage medaka and zebrafish to environmental carcinogens. Defects
in mismatch repair (MMR) of DNA replication errors induced neurofibroma and
olfactory neuroblastoma in 6-month to 2-year-old zebrafish mutants with loss-of-function
in the major MMR genes (Feitsma et al. 2008). Postembryonic increased expression of
fibroblast growth factor 8 (fgf8) in zebrafish insertional mutants has led to the
development of neuroblastoma by preventing terminal differentiation of neural precursor
cells (Amsterdam et al. 2009). Co-expression of both the hedgehog and AKT pathways
has led to the development of glioblastoma and astrocytoma in transgenic zebrafish <3
months of age (Ju et al. 2009). Environmental carcinogens such as the direct-acting
nitrosamide N-methyl-N’-nitro-N-nitrosoguanidine (MNNG) has been reported to cause
esthesioneuroblastoma in medaka (Spitsbergen et al. 2000b). Polycyclic aromatic
hydrocarbon 7, 12-dimethyl-benz[a]anthracene (DMBA) has been shown to cause
neuroblastoma in zebrafish (Spitsbergen et al. 2000a).
Based on histomorphology, staining characteristics and S-100 OLC
immunohistochemistry (IHC), the brain tumour described herein is, to our knowledge, the
first reported case of DNET in a zebrafish. Although its aetiology is uncertain, heritable
genetic defects or environmental carcinogens have been shown to cause neuroectodermal
tumours in zebrafish.
195
ACKNOWLEDGEMENTS
The authors would like to thank K. Fischer, M. Corbus and R. Norred for
excellent histotechnical assistance. This work was supported by National Institutes of
Health NCRR grants 5R24RR017386-02, 1 T32 RR023917 and P40 RR12546.
196
Appendix Figure 2.1. Tumour growth pattern and infiltrative nature is revealed by sheets
of neoplastic cells (asterisks) extending through the neurocranium into the retrobulbar
space, around the inner ear and dissecting between skeletal muscle fibre bundles. HE.
Bar, 1 mm.
197
Appendix Figure 2.2. Microcystic pattern of the tumour growth containing intracystic
‘floating neurons’ embedded in sparse mucinous material (arrows) with a hypercellular
background composed of OLCs. HE. Bar, 25 µm.
198
Appendix Figure 2.3. Abnormally oriented and misshapen dysmorphic neurons with
eccentric nuclei and putative balloon cells (arrows) within a focus of cerebellar cortical
dysplasia adjacent to the tumour. LFB. Bar, 25 µm.
199
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