Mycobacteriosis in Zebrafish Colonies
Christopher M. Whipps, Christine Lieggi, and Robert Wagner
Abstract
Mycobacteriosis, a chronic bacterial infection, has been associated with severe losses in some zebrafish facilities and
low-level mortalities and unknown impacts in others. The
occurrence of at least six different described species (Mycobacterium abscessus, M. chelonae, M. fortuitum, M. haemophilum, M. marinum, M. peregrinum) from zebrafish complicates
diagnosis and control because each species is unique. As a
generalization, mycobacteria are often considered opportunists, but M. haemophilum and M. marinum appear to be more
virulent. Background genetics of zebrafish and environmental conditions influence the susceptibility of fish and
progression of disease, emphasizing the importance of regular monitoring and good husbandry practices. A combined
approach to diagnostics is ultimately the most informative,
with histology as a first-level screen, polymerase chain reaction for rapid detection and species identification, and culture for strain differentiation. Occurrence of identical strains
of Mycobacterium in both fish and biofilms in zebrafish
systems suggests transmission can occur when fish feed on
infected tissues or tank detritus containing mycobacteria.
Within a facility, good husbandry practices and sentinel programs are essential for minimizing the impacts of mycobacteria. In addition, quarantine and screening of animals coming
into a facility is important for eliminating the introduction of
the more severe pathogens. Elimination of mycobacteria
from an aquatic system is likely not feasible because these
species readily establish biofilms on surfaces even in extremely low nutrient conditions. Risks associated with each
commonly encountered species need to be identified and informed management plans developed. Basic research on the
growth characteristics, disinfection, and pathogenesis of
zebrafish mycobacteria is critical moving forward.
Christopher M. Whipps, PhD, is an assistant professor in the Department
of Environmental and Forest Biology, State University of New York College
of Environmental Science and Forestry in Syracuse, New York. Christine
Lieggi, DVM, DACLAM, is the associate director and head of veterinary
services at the Research Animal Resource Center, Memorial SloanKettering Cancer Center and Weill Cornell Medical College in New York,
New York. Robert Wagner, VMD, Dipl. ABVP-ECM, is the chief of surgical
veterinary services and associate professor in the Division of Laboratory
Animal Medicine, University of Pittsburgh, Pennsylvania.
Address correspondence and reprint requests to Dr. Christopher M.
Whipps, State University of New York College of Environmental Science
and Forestry, Environmental and Forest Biology, 1 Forestry Drive, Syracuse,
NY 13210 or email [email protected].
Volume 53, Number 2
2012
Key Words: biofilms; disinfection; mycobacteriosis; Mycobacterium chelonae; Mycobacterium haemophilum; Mycobacterium marinum; surveillance; zebrafish
Introduction
T
he use of zebrafish (Danio rerio) as a model in biomedical research has expanded greatly in the last 15
years. Studies range from human disease research
(Lieschke and Currie 2007) to infectious disease including
mycobacteriosis (Lesley and Ramakrishnan 2008; van der
Sar et al. 2004), cell function (Ellett and Lieschke 2010),
toxicology (Stanley et al. 2009), and behavior (Norton and
Bally-Cuif 2010). These are only a few examples and, as
research expands, so does the need for development of a
comprehensive approach to managing zebrafish health.
With a focus on infectious disease as a primary component
of zebrafish health, the most commonly encountered diseases
are microsporidiosis, caused by Pseudoloma neurophilia
(Kent and Bishop-Stewart 2003), and mycobacteriosis,
caused by several species of the genus Mycobacterium (Kent
et al. 2004). Such infectious diseases have the potential to
impact research and should be eliminated or at least controlled
to the point where the impact is minimal (Baker 2003).
Mycobacterium-related nonprotocol variation is poorly
quantified in zebrafish (Kent et al. 2012, in this issue), but
the common occurrence of these infections warrants investigation. Whereas control of the obligate parasite P. neurophilia
appears feasible through strict biosecurity policies (Kent
et al. 2011; Sanders et al. 2012, in this issue), Mycobacterium species present numerous additional challenges for
control.
Mycobacteria are facultative pathogens that can survive
outside the host in surface biofilms (Falkinham 2009;
Falkinham et al. 2001). Thus, screening of animals and eggs
alone may be ineffective for eliminating mycobacteria that
may colonize a facility by water, food, or personnel (clothing,
hands, equipment), and may grow and spread in the absence
of a host animal. This is exemplified in the aforementioned efforts of Kent and colleagues (2011) to establish a
P. neurophilia–free colony of fish; the obligate parasite was
eliminated but mycobacteriosis was still reported in a small
proportion of fish. There is no single etiological agent of mycobacteriosis; instead, several species and strains have been
identified (Kent et al. 2004; Whipps et al. 2008). As such, manifestation of disease varies by species and strain, complicating
95
diagnosis and management. In addition to bacterial strain
variation, host genetic variation plays a role in susceptibility
(Murray et al. 2011; Whipps et al. 2008); suggesting management strategies need to be tailored to balance biosecurity
efforts with value and susceptibility of zebrafish lines.
The challenges inherent in the control of mycobacteriosis
highlight the need for continued epidemiological studies
connecting Mycobacterium species and strains to their potential sources, empirical data on the impact of subclinical
infections, development of rapid and specific diagnostic
tests, and realistic management plans with proven efficacy
determined through controlled experimentation.
Mycobacteriosis in Zebrafish
Piscine mycobacteriosis is well recognized in marine and
freshwater fishes, with several reviews written on the topic
(Decostere et al. 2004; Frerichs 1993; Gauthier and Rhodes
2009; Parisot 1958). Reviews indicate the common occurrence
in many fish species, involvement of more than one species
of Mycobacterium in fish infections, ranging manifestation
of disease (although typically including granuloma formation),
potential for zoonotic transmission, and challenges for treatment
and control. These same themes can be applied to zebrafish.
Historically, mycobacteriosis in fishes was attributable
primarily to M. chelonae, M. fortuitum, and M. marinum
(Frerichs 1993). However, increased study and more refined
diagnostic methods (i.e., DNA sequencing) have identified
at least 16 described Mycobacterium species from fish
(Gauthier and Rhodes 2009; Whipps et al. 2007b). In zebrafish,
at least 6 described species of mycobacteria have been reported (Table 1). These infections were first documented by
Astrofsky and colleagues (2000) when M. abscessus, M. chelonae, and M. fortuitum were isolated from zebrafish exhibiting decreased survival and reproductive output. Subsequently,
Kent and colleagues (2004) identified several other species
from facilities experiencing different degrees of mortality.
Mycobacterium peregrinum and M. haemophilum were associated with severe disease. In facilities experiencing moderate
to minimal mortalities, M. chelonae and an M. chelonae–like
bacterium were isolated. Watral and Kent (2007) added
M. marinum to this growing list, isolating the bacterium from
zebrafish from a facility supplying fish to the research community and experiencing low to moderate levels of mortality.
Manifestation of mycobacterial disease in zebrafish is
broad ranging (Astrofsky et al. 2000; Kent et al. 2008). Externally, fish may present with nonspecific dermal lesions
(Figure 1), have raised scales, or have swollen abdomens.
Emaciation may occur (Figure 1) and is, in our experience,
very often associated with M. haemophilum infection
(Whipps et al. 2007a). Behaviorally, fish may swim erratically or be lethargic. Often, animals will show no signs of
disease (Kent et al. 2004; Whipps et al. 2007a). Internally,
granulomas may be visible throughout all tissues but primarily in the spleen, kidney, and liver. Diffuse systemic infections without prominent granulomas have been reported for
M. haemophilum (Whipps et al. 2007a) and M. marinum
(Ramsay et al. 2009b). Bacteria have been observed in the
ovaries, suggesting the potential for contamination of offspring, if not vertical transmission (Kent et al. 2004). Involvement of the swim bladder (aerocystitis) is not uncommon
(Whipps et al. 2008). Zebrafish are physostomus, and thus
the swim bladder is directly connected to the gastrointestinal
tract by a pneumatic duct. This connection provides a possible route of infection to the swim bladder. Bacteria may be
observed in the intestinal epithelium and within the lumen
(Whipps et al. 2007a), indicative of shedding across this surface and excretion in the feces.
Transmission of mycobacteria through ingestion has
been demonstrated in other fishes (Ross 1970) and is consistent with the intestine likely being the primary route of invasion (Harriff et al. 2007). Bacteria from infected animals
may be shed from skin lesions or the intestine (Noga 2010),
providing a continuous source of mycobacteria in affected
tanks. This highlights the importance of rapid removal of
dead fish to minimize transmission through cannibalization,
as well as removal of any moribund animals, which might
act as reservoirs of infection. The oral route of transmission
suggests food presents a risk for exposure. Testing for mycobacteria in feed at one facility (Whipps et al. 2008) yielded
negative results; however, Beran and colleagues (2006) reported mycobacteria from brine shrimp eggs, one of the
most commonly used feeds for zebrafish. The use of live
Table 1 Mycobacterium species known to infect zebrafish in research facilities
Species
Source
Mycobacterium abscessus
Mycobacterium chelonae
Mycobacterium chelonae–like
Mycobacterium fortuitum
Mycobacterium haemophilum
Mycobacterium marinum
Mycobacterium peregrinum
Astrofsky et al. (2000); Watral and Kent (2007)
Astrofsky et al. (2000); Kent et al. (2004); Whipps et al. (2008)
Kent et al. (2004); Whipps et al. (2007a)
Astrofsky et al. (2000)
Whipps et al. (2007b)
Watral and Kent (2007)
Kent et al. (2004)
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ILAR Journal
(2008) examined M. marinum isolates from fish and humans
from the same location and found identical pulsed field gel
electrophoresis cutting patterns from both hosts. These data
do not rule out a common source of infection, as opposed to
direct transmission, but illustrate that the same strains can
infect fish and humans. Human core body temperature
(37°C) is thought to limit the establishment and spread of
fish-associated Mycobacterium species infections to the
extremities, and, although isolates from zebrafish tend not to
grow at 37°C by plate culture, replication may be observed
at 37°C in macrophage culture or mouse footpad assays
(Harriff et al. 2008; Kent et al. 2006). To the best of our
knowledge, there are no documented cases of human mycobacteriosis associated with zebrafish handling.
Diagnostic Methods
Figure 1 (A, B) External lesions (arrows) associated with Mycobacterium marinum infection in zebrafish. (C) Severe emaciation
associated with Mycobacterium haemophilum infection.
feed may present some risk, but comprehensive screening is
required to evaluate this risk. Recognition of the role of surface biofilms in persistence of mycobacteria in a system and
as a potential source of infection is increasing.
It is important to note the zoonotic potential of Mycobacterium species. Transmission between fish species has been
demonstrated by feeding infected tissues to other species
(Ross 1970), and the same genetic strains of M. marinum
have been reported from zebrafish and hybrid striped bass
(Ostland et al. 2008). Mycobacterium marinum is of particular concern because it is known to infect humans. Such infections are associated with aquarium maintenance or handling
food fishes (Ang et al. 2000; Aubry et al. 2002). Swimming
or other direct contact with sea water (Jernigan and Farr
2000) is also associated with cases of M. marinum infection
in humans. Approximately 84% of infections in humans
have been associated with contact with home aquaria (Aubry
et al. 2002). Other species (Table 1) are potential opportunistic human pathogens (Brown-Elliott and Wallace 2002;
Whipps et al. 2007a). Whereas direct transmission from fish
to humans has not been confirmed, Ostland and colleagues
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2012
The reported nonspecific signs of disease or complete absence of clinical signs of disease dictates that mycobacteriosis in zebrafish be confirmed by an established diagnostic
method. Several tools are available, with relative benefits
and limitations. Traditionally, diagnosis has relied on histological examination of zebrafish sections stained with
Ziehl-Neelsen or Fite’s acid fast stain and this is the primary
technique recommended by the Zebrafish International Resource Center for routine surveillance. Observation of acidfast bacilli in tissue sections correlates well to results from
culture for M. chelonae (Whipps et al. 2008) and may have
improved detection for difficult to grow species such as
M. haemophilum (Whipps et al. 2007a) and M. marinum
(Ramsay et al. 2009b). Histology also has the advantage that
most of the major internal organs of a zebrafish can be examined in a single section. The most important disadvantage of
diagnosis using acid-fast stained sections is that the species
of bacterium cannot be readily identified. Touch imprints of
spleen, kidney, or liver stained with Kinyoun’s acid-fast stain
have this same limitation with regard to identification but
can be carried out quickly and are standard procedure in our
diagnostic screening. For severe diffuse infections, imprint
results correlate well to histology and culture (Whipps et al.
2007a), but in less severe cases tissue imprints are less consistent (C. Whipps, unpublished data). Both tissue sectioning
and touch imprints are important for diagnosis when the
Mycobacterium species present is difficult to grow.
Diagnosis of mycobacteria of fish in general is reviewed
well by Gauthier and Rhodes (2009); the following techniques relate to implementation in zebrafish. Culture of
mycobacteria from zebrafish is typically accomplished on
Middlebrook 7H10 agar supplemented with oleic acid, albumin, dextrose, and catalase or Lowenstein-Jensen slants
(Kent and Kubica 1985). Pretreatment with 1% cetyl pyridinium chloride prior to plating is suggested for cultures
from fish tissues and strongly recommended for environmental samples to minimize background growth. Cultures
are typically incubated at 28-30°C and monitored for growth
for 6 to 8 weeks. Rapid growers such as M. chelonae or
97
M. abscessus form visible colonies within 5 days. Mycobacterium marinum grows more slowly on Middlebrook 7H10
agar, forming colonies in 10 to 14 days. Alternatives to
Middlebrook 7H10 agar may be more appropriate for M.
marinum; for example, Ostland and colleagues (2008) used
Columbia with colistin and nalidixic acid agar supplemented
with 5% sheep blood. Mycobacterium haemophilum is the
epitome of a slow-growing species, forming visible colonies
in 6 to 8 weeks on Middlebrook 7H10 agar supplemented
with oleic acid, albumin, dextrose, and catalase and 60 μM
hemin (Whipps et al. 2007a). The lengthy incubation time
and specialized medium may contribute to the underdiagnosis of this important zebrafish pathogen. In all cases, any suspect colonies are tested with an acid-fast stain to verify their
presumptive identity as mycobacteria.
A limitation of culture is the sometimes prolonged incubation time required to identify the species. Although culture
remains a gold standard for a thorough investigation and
subsequent storage and cataloguing of species and strains,
more rapid methods of diagnosis and identification are required. Thus, DNA-based methods have risen to the forefront as diagnostic tools in zebrafish. For routine polymerase
chain reaction (PCR1) testing, the hsp65 primers of Selvaraju
and colleagues (2005) are effective and have the secondary
benefit of providing amplified fragments of adequate length
for sequencing and identification. The sensitivity of these
primers has not been determined in zebrafish. Speciesspecific tests are desirable for high-impact species such as
M. haemophilum and have been implemented in outbreak
investigations (Whipps et al. 2007a). PCR and restriction enzyme analysis (Talaat et al. 1997; Telenti et al. 1993) remains
a rapid and useful technique for detection and identification
of species. Quantitative PCR methods for detection of members of the genus Mycobacterium in general have been developed (Jacobs et al. 2009c; Zerihun et al. 2011) and are
currently under investigation for broad-scale use in zebrafish.
A desirable diagnostic test should be highly sensitive and
provide at least a species-level identification simultaneously.
High-resolution melting curve analysis of PCR products has
promise to quickly differentiate species and has been applied
to M. tuberculosis (Choi et al. 2010) and the M. avium complex (Castellanos et al. 2010). The continued challenge is an
adaptable diagnostic method that can readily accommodate
new strains and species as they become known.
Fresh tissues are preferred for culture, direct DNA
extractions, and PCR testing, but frozen and ethanol-fixed
tissues are also appropriate if culture is not absolutely necessary. PCR confirmation and species identification of mycobacteria from histological sections are less optimal but can be
done (Jacobs et al. 2009a; Loeschke et al. 2005; Pourahmad
et al. 2009; Zerihun et al. 2011). In fixed specimens, the duration
of fixation, method of decalcification, and thickness of sections all influence the successful detection of mycobacteria
by PCR. We first identified M. haemophilum as an important
1Abbreviation that appears ≥3x throughout this article: PCR, polymerase
chain reaction
98
cause of disease in zebrafish using PCR directly on tissues
(Kent et al. 2004). A common target is the small subunit
(SSU) rDNA sequence, a highly conserved gene that is useful for higher taxonomic comparisons. However, there are
a few species that have identical SSU rDNA sequences
(Tortoli 2003) and researchers have turned to other genes,
such as the ITS region of the rDNA, to better elucidate relationships of closely related Mycobacterium spp. (Roth et al.
2000). Taking this approach, we were able to identify differences in ITS sequence from two M. chelonae isolates from
zebrafish that had identical SSU rDNA sequences (Kent et al.
2004). The heat shock protein 65 (hsp65) gene is also used
for reliable mycobacterial species identifications (Kim et al.
2005; McNabb et al. 2006) and may differentiate between
species that SSU rDNA sequence analysis cannot (Ringuet
et al. 1999). We have found hsp65 to be the most rapid and
reliable gene for preliminary identification of fish mycobacteria, even able to subdivide strains of M. chelonae by the
variation in hsp65 (Whipps et al. 2008). Other potentially
useful genes that have been used less extensively for zebrafish
mycobacteria are erp (Jacobs et al. 2009a), rpoB, and sod
(Devulder et al. 2005).
Subdivisions within species can be accomplished to
some extent from DNA sequence data, but strain differentiation requires greater resolution and a pure culture (Whipps
et al. 2008). Pulsed field gel electrophoresis effectively resolved strains of M. marinum (Ostland et al. 2008) and M.
chelonae (Vanitha et al. 2003), but in our experience was
less useful for M. salmoniphilum, M. peregrinum, and M.
abscessus (C. Whipps, unpublished data). Furthermore, pulsed
field gel electrophoresis is excessively time consuming, and
more rapid methods of strain differentiation have been
adopted, specifically enterobacterial repetitive intergenic
consensus PCR (Sampaio et al. 2006). To supplement enterobacterial repetitive intergenic consensus PCR, other randomly amplified polymorphic DNA methods can also be
used (Zhang et al. 1997). Results are rapid, can be performed
in any lab with a thermocycler, and are straightforward to
interpret. This method has been applied to M. chelonae from
zebrafish in a single facility, distinguishing multiple strains
(Whipps et al. 2008), and we continue to use this method as
a routine part of our diagnostic screening of cultures from
both fish and environmental sources.
Variation in Mycobacterium Species
from Zebrafish
Six described Mycobacterium species have been reported
from zebrafish (Table 1). As much as mycobacteriosis is
caused by several different species, these species entities can
be further subdivided into strains, with their own unique
properties and challenges. Strain delineation is accomplished through biochemical analysis, DNA sequencing, and
DNA fingerprinting methods, and multiple strains of piscine
M. chelonae and M. marinum have been described (Ostland
et al. 2008; Whipps et al. 2008). Other entities that we cannot
ILAR Journal
ascribe to species are not listed here, with the exception of
an M. chelonae–like species we had originally identified as
M. chelonae (Kent et al. 2004), but subsequent phylogenetic
analyses demonstrated that it was a distinct entity and should
be described as a new species (Whipps et al. 2007b).
The manifestation of disease varies with species, and
each can be broadly categorized as either pathogen or opportunist. The reality, however, is a continuum between these
extremes, with M. chelonae, M. abscessus, M. fortuitum, and
M. peregrinum tending to be associated with low-level,
chronic disease, whereas M. marinum and M. haemophilum
are much more virulent, causing disseminated infections and
higher levels of mortality (Kent et al. 2004; Ostland et al.
2008; Watral and Kent 2007; Whipps et al. 2007a). The differences may be attributable to genetic elements associated
with virulence that are present in some species and strains
but not others (Harriff et al. 2008). Although not included in
the study by Harriff and colleagues (2008), this is likely the
case with M. haemophilum and M. marinum, as these species
tend to cause mortality in experimental exposure studies
(Figure 2) (Watral and Kent 2007; Whipps et al. 2007a).
Mycobacterium chelonae appears to be largely an opportunistic pathogen, recognizing that multiple strains exist and
may vary in their ability to cause disease (Whipps et al.
2008). Other species (M. abscessus, M. peregrinum) have
been associated with increased disease, but when fish are
challenged experimentally (Watral and Kent 2007) with
these same strains, they may produce largely subclinical infections (Figure 2). This suggests that species and strains
vary in their pathogenicity, and when less pathogenic species
are associated with an outbreak, the increased mortalities
may be not only related to the infection but also due to suboptimal environmental conditions.
Environmental conditions and the nutritional and immunological state of zebrafish are probably the most important variables affecting the pathogenesis of mycobacteriosis
when the cause is an opportunistic species. These conditions likely play a less significant role with overt pathogens
such as M. haemophilum and M. marinum, but suboptimal
environmental parameters will exacerbate these infections
to a greater degree. Ramsay and colleagues (2009a, 2009b)
evaluated the role of stress on cortisol level in zebrafish
and the progression of disease with fish exposed to either
M. chelonae or M. marinum. In the stressed zebrafish,
Ramsay and colleagues (2009b) observed increased prevalence of infection for M. chelonae, greater mortality in
Figure 2 Relative pathogenicity of Mycobacterium species isolated from zebrafish in colonies experiencing minimal mortalities or Mycobacteriumassociated outbreaks with significant mortality (OB). Mortality and histology were used as endpoints for all experiments. Data are summarized
from studies carried out by Watral and Kent (2007) and Whipps et al. (2007b).
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99
M. marinum–exposed fish, and an increased number of disseminated infections for both pathogens when compared
with unstressed controls. In striped bass, suboptimal diet
influenced the progression of disease associated with M.
marinum infections, with a greater likelihood of diffuse
systemic disease in treatments versus controls (Jacobs et al.
2009b). Different lines of zebrafish appear to be more or
less susceptible to infection based on the cross-sectional
study at a single facility (Murray et al. 2011; Whipps et al.
2008). Tobin and colleagues (2010) have identified at least
one locus in zebrafish important for M. marinum susceptibility,
and Hegedus and colleagues (2009) and van der Sar (2009)
characterized the transcriptome of infected zebrafish, suggesting that tools for comparison are available. A streamlined approach where multiple lines can be evaluated with
multiple species of mycobacteria will be necessary to identify the common and unique genetic drivers of susceptibility and disease.
Mycobacteria in Biofilms
The ability for at least some, if not all, piscine mycobacteria
species to persist in surface biofilms in aquatic systems
(Beran et al. 2006; Whipps et al. 2008) presents additional
challenges for interrupting the cycle of infection once it is
established. This highlights the role of a quarantine program
to minimize potential introductions of the more pathogenic
mycobacteria, regular monitoring of populations to remove
infected fish, and the routine cleaning and disinfection of
impacted tanks and equipment. Mycobacteria are hydrophobic and oligotrophic, requiring low levels of dissolved organic carbon (Falkinham 2009). As such, they readily adhere
to surfaces and are adapted to survival in “clean” water systems such as aquaria. The community of mycobacteria in
biofilms can be diverse (Schulze-Röbbecke et al. 1992), with
a variety of species having been isolated from aquaria
(Beran et al. 2006; Whipps et al. 2007a, 2008). Although many
of these species have never been reported from fish, those
that are found in zebrafish have also been reported from biofilms. Furthermore, genetic comparisons of Mycobacterium
isolates have revealed identical strains of M. chelonae in
zebrafish (Whipps et al. 2008) and M. marinum in pompano
Trachinotus carolinus (Yanong et al. 2010), as are found in
associated biofilms. The finding of the same strains in fish
and biofilms presents a question of which is the source and
which is the sink. The feeding habits of zebrafish and the
descriptive studies below suggest biofilms are indeed a
source of infection, although fish shedding bacteria are also
contributing to the biofilms.
In the aquatic environment, biofilms are found on all surfaces, and this biofilm and the detritus at the bottom of the
tank are thought to be the source of mycobacterial infection
in zebrafish. Zebrafish are thought to be generalist consumers (Lawrence 2007), primarily feeding in the water column
but also on the surface and substrate (Spence et al. 2008). In
our observations of fish in a sump at a large facility, zebrafish
100
hunt for benthic organisms and slow zooplankton along the
biofilm scaffolding. They are probably also eating bacteria,
slime molds, and protozoa. The larvae are especially active
grazers and hunters once they get to be about 8 to 10 mm.
These fish and larvae are probably selecting specific organisms but are likely consuming mycobacteria incidentally.
Given the suggested oral transmission route (Harriff et al.
2007), zebrafish may be infected by the oral route by grazing
on the microflora of the biofilm and detritus. The following
are our observations on mycobacteria in biofilms relating to
infections at two facilities.
Sump Fish Case Study: Facility A
Dense mature biofilms in the dirty sump yielded the highest concentrations of mycobacteria by acid-fast staining
and PCR testing, as compared with frequently cleaned,
minimal density sump tank biofilms. Sentinel fish (raised
from larvae stage) from tanks that were frequently cleaned
with minimal density biofilms have been mycobacteria
negative for 3 years. The sentinel fish from the dense mature biofilm tanks yielded at least 1% fish positive for mycobacteria on histology each year for the past 3 years. It
was determined that the species of mycobacteria causing
disease in the fish and the species of mycobacteria found
in the biofilm and detritus were the same, M. fortuitum.
Frequent elimination of detritus and minimization of the
biofilm by cleaning or tank changes should decrease fish
exposure to mycobacterial pathogens; this seems to be especially true when raising larval stages of fish. Mycobacterium fortuitum infection appears to be a manageable
disease but is not usually entirely eliminated from a large
recirculation zebrafish system.
Biofilm Monitoring: Facility B
An initial evaluation and continued monitoring of biofilms
was conducted in a facility that experienced morbidity and
mortality in zebrafish as a result of M. haemophilum infection. Mycobacteriosis was initially diagnosed by histologic
methods and later confirmed with PCR. Following the diagnosis of several cases of M. haemophilum infection, surface
biofilm sampling was conducted to determine the identity of
existing mycobacterial florae and to monitor this community
following disinfection and repopulation of the system (Table 2).
Initially, M. chelonae and M. abscessus were the only species isolated from the biofilms. Unlike our earlier studies of
M. haemophilum (Whipps et al. 2007a), we were unable to
detect this bacterium in the biofilms.
The aquatic housing system was depopulated and all
nonreplaceable system components were thoroughly disinfected with 1000 ppm bleach buffered to a pH of 7. Following disinfection, biofilm samples were evaluated from all
locations on the system and proven to be negative by both
culture and PCR (Table 2) before introducing larvae from
bleached disinfected embryos (25-50 ppm for 10 min).
ILAR Journal
Table 2 Ongoing biofilm sampling of a zebrafish system over 21 months
Date
System status
Location
Culture result
Species identificationa
September 9, 2009
System with fish
March 22, 2010
System with fish
November 29, 2010
System bleached,
no fish
System bleached and
scrubbed, no fish
System
reassembled,
no fish
Fish placed on
system
System with fish
Positive
Positive
Positive
Positive
Positive
All negative
Mycobacterium chelonae
M. chelonae
M. chelonae
Mycobacterium abscessus
M. chelonae
July 19, 2010
Gutters
Tank with fish
Gutters
Tank with fish
Sump
Gutters, tank,
sump
Gutters, tank,
sump
Gutters, tank,
sump
All negative
December 13, 2010
System with fish
December 27, 2010
System with fish
Gutters, tank,
sump
Gutters, tank,
sump
Gutter
Populated tank
August 10, 2010
October 8, 2010
November 15, 2010
January 10, 2011
System with fish
January 24, 2011
System with fish
February 7, 2011
System with fish
February 22, 2011
System with fish
March 7, 2011
System with fish
March 21, 2011
System with fish
All negative
All negative
All negative
Negative
Positive (multiple)
Sump
Gutter
Populated tank
Sump
Gutters, tank,
sump
Gutter
Positive (multiple)
Positive
Negative
Positive
All negative
Populated tank
Sump
Gutter
Populated tank
Sump
Gutter
Populated tank
Sump
Gutter
Populated tank
Sump
Positive (multiple)
Positive
Positive (multiple)
Positive
Positive (multiple)
Positive
Positive (multiple)
Positive (multiple)
Positive
Positive
Positive
Positive (multiple)
M. abscessus, Mycobacterium
phocaicum, Mycobacterium
gordonae
M. phocaicum, M. chelonae
M. gordonae
M. phocaicum
M. phocaicum, Mycobacterium
fortuitum
M. phocaicum, M. chelonae
M. phocaicum
M. chelonae, M. phocaicum
M. chelonae
Mycobacterium mucogenicum
M. phocaicum
M. chelonae, M. phocaicum
M. chelonae
M. chelonae
M. phocaicum
M. phocaicum
Swabs were used to collect surface biofilms from three locations within the system on a repeated basis. Biofilms were processed as described
by Whipps et al. (2008) and grown on MB7H10 medium supplemented with hemin at 28°C for 8 wk. Where “multiple” is noted, several colony
types were observed on the culture medium.
aSpecies identification based solely on BLAST searching of hsp65 DNA sequences. In some cases an equally likely match was obtained
(e.g., M. phocaicum and M. mucogenicum).
Less than 3 weeks after the introduction of larvae, a biofilm
sample from a populated tank was again PCR positive for
mycobacteria. By culture methods, mycobacteria remained
below detectable levels after fish were placed on the system
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2012
for approximately 6 weeks, at which time mycobacteria
were isolated from the populated tank and system sump. Continued biweekly sampling in subsequent months demonstrated colonization of the system by at least five species:
101
M. abscessus, M. chelonae, M. fortuitum, M. gordonae, and
M. phocaicum/mucogenicum. These species continued to be
found throughout the biweekly biofilm sampling until June
2011 when monitoring of biofilms stopped. Mycobacteriosis
was identified in an occasional fish following the clean-up
(M. chelonae), but M. haemophilum was not detected again.
Furthermore, there have been no indications of M. haemophilum outbreaks in this facility to date (September 2012). This
demonstrates several key points. First, the decontamination
was effective in killing the mycobacteria present in the system. Second, by culture, mycobacteria did not reach detectable levels until well after fish were stocked into the system,
suggesting that although mycobacteria are oligotrophic, the
organic load contribution of fish likely enhances growth and
establishment of biofilms. Finally, the Mycobacterium community was different before and after disinfection, with an M.
phocaicum–like organism dominating in more recent biofilm
samples. This may represent an early colonizer and transitional community, but M. phocaicum is not a bacterium that
has been identified from zebrafish and may occupy substrate
that prevents colonization of other species. These data also
suggest that, although biofilms may indeed be a source of
infection for fish, the initial colonization by the fish-specific
pathogens likely requires an animal source.
Control and Treatment
Recommendation for control of zebrafish diseases were recently reviewed (Kent et al. 2009), and the importance of
quarantine, disinfection, a functioning ultraviolet system,
and sentinel programs to monitor for disease were highlighted. These are broadly applicable to mycobacteria, and
aspects of these have been discussed (Astrofsky et al. 2000;
Kent et al. 2004; Whipps et al. 2008). Elimination of mycobacteria from a facility once it has been established
is challenging, and depopulation is often recommended
(Francis-Floyd and Yanong 1999). This may not be feasible
in all cases, and thus management of the endemic disease
is often the appropriate choice. Minimizing any potential
source of infection by cleaning tanks to reduce biofilms or
cleaning and upgrading ultraviolet sterilization for water
likely helps drive down free bacteria in the system. Removal
of sick animals and affected tanks rapidly will minimize
spread. The standard practice of bleaching eggs for 10 minutes at 50 ppm (Westerfield 2000) has unknown efficacy for
killing the many different strains of mycobacteria found in
zebrafish laboratories. Further complicating this matter is
that mycobacteria are known to be differentially susceptible to
disinfection, whether planktonic or in biofilms (Bardouniotis
et al. 2003; Steed and Falkinham 2006). Ferguson and colleagues (2007) highlighted the importance of adjustment to
pH 7 of the bleach solution as well as validation of concentration using a chlorine meter. Mainous and Smith (2005)
reported that 50 ppm bleach was ineffective at killing
M. marinum in culture after a 10-minute exposure, requiring
60 minutes for complete germicidal activity. Studies under
102
way in the Whipps laboratory using pH-adjusted bleach at
50 ppm for 10 minutes show complete germicidal effect for
M. chelonae cultures at 103, 104, and 105 colony-forming
units per milliliter. The efficacy of bleach on biofilms of
these same bacteria, such as those that may be present on
zebrafish eggs, is not known. Bacteria harbored within eggs
are protected from disinfection and present a risk for contamination of other eggs. Vaccination with extracellular mycobacterial products (Chen et al. 1996) and DNA vaccines
(Pasnik and Smith 2005) have demonstrated the efficacy
of these techniques in stimulating specific antibody production and eliciting an immune response. Similarly, vaccination
with attenuated mycobacteria shows some promise for protective immunity in zebrafish (Cui et al. 2010) and other aquaculture species (Kato et al. 2011).
Little is known about the efficacy of antibiotic treatment
for mycobacteriosis in fish because treatment in food fish
aquaculture is impractical due to high costs of antibiotics,
long treatment regimens, and concerns about use of these
powerful human drugs in fish destined for human consumption. Nevertheless, treatment of infected zebrafish may be
appropriate when extremely valuable strains or populations
are involved. Subclinical infections in valuable lines of
zebrafish may need to be treated to remove this confounding
factor from experiments planned for these animals or minimize the chances of vertical transmission when breeding
new fish for research. Mycobacteria are known to be susceptible to rifampicin and Kawakami and Kusuda (1990) reported that rifampicin, streptomycin, and erythromycin were
effective for reducing mortalities associated with a Mycobacterium sp. in cultured yellowtail (Seriola quinqueradiata).
In this study, however, only an initial dose at the time of exposure and a dose at 24 hours postexposure were given to the
fish. Although infection was not completely eliminated in
fish after 7 weeks, more regular treatments may have been
effective. In contrast, Hedrick and colleagues (1987) found
rifampicin treatment of M. marinum–infected striped bass
ineffective after 60 days of feeding the bass antibioticsupplemented feed. These differences across host species
suggest that results obtained from one fish species may not
be applicable to all species, and that treatment must be tested
specifically in zebrafish to determine if it is an effective
option.
Summary and Future Directions
The impacts of mycobacteria are clear when associated with
mortalities and decreased reproductive output in zebrafish.
What is less well understood is the impact of subclinical
infections and the influence of these infections in specific
areas of research. Such nonexperimental variation magnifies
the importance of evaluating the influences these infections
may have when zebrafish are used as models for studies on disease, immunology, ecotoxicology, and so on. The more serious
pathogens, M. haemophilum and M. marinum, are those of
greatest concern for disease outbreaks, but the opportunists still
ILAR Journal
present a significant concern because they often go unnoticed and their impacts are less well known. Complete elimination of mycobacteria from a large facility supplied with
recirculating water is probably not feasible given the ease
with which mycobacteria can colonize such systems. Through
strict biosecurity protocols, it is worthwhile to attempt to
exclude the more virulent pathogens that may colonize a
facility initially through the introduction of infected fish from
another facility where the pathogen is endemic. Elimination
of all mycobacteria in small-scale operations with flowthrough water may be possible when it is called for in quarantine or when using highly susceptible fish. Monitoring and
management of the disease are sure to be the focus of continued research that emphasizes mycobacterial ecology, biology, and pathogenesis in zebrafish so that management
actions can be better informed.
Acknowledgments
The authors are grateful to the three anonymous reviewers
for their helpful comments and suggestions. CM Whipps
thanks Hadi L. Jabbar (State University of New York College
of Environmental Science and Forestry) for bacterial isolation
and PCR on fish and biofilm samples. C Lieggi acknowledges the assistance of Yuri Igarashi for sample collection
and Aziz Toma for culturing and shipping all of the samples
originating at Weill Cornell Medical College.
References
Ang P, Rattana-Apiromyakij N, Goh CL. 2000. Retrospective study of Mycobacterium marinum skin infections. Int J Dermatol 39:343-347.
Astrofsky KM, Schrenzel MD, Bullis RA, Smolowitz RM, Fox JG. 2000.
Diagnosis and management of atypical Mycobacterium spp. infections
in established laboratory zebrafish (Brachydanio rerio) facilities. Comp
Med 50:666-672.
Aubry A, Chosidow O, Caumes E, Robert J, Cambau E. 2002. Sixty-three
cases of Mycobacterium marinum infection: Clinical features, treatment, and antibiotic susceptibility of causative isolates. Arch Intern
Med 162:1746-1752.
Baker D. 2003. Natural Pathogens of Laboratory Animals: Their Effects on
Research. Herndon VA: ASM Press.
Bardouniotis E, Ceri H, Olson ME. 2003. Biofilm formation and biocide
susceptibility testing of Mycobacterium fortuitum and Mycobacterium
marinum. Curr Microbiol 46:28-32.
Beran V, Matlova I, Dvorska L, Svastova P, Palik I. 2006. Distribution of
mycobacteria in clinically healthy ornamental fish and their aquarium
environment. J Fish Dis 29:383-393.
Brown-Elliott BA, Wallace RJ Jr. 2002. Clinical and taxonomic status of
pathogenic nonpigmented or late-pigmenting rapidly growing mycobacteria. Clin Microbiol Rev 15:716-746.
Castellanos E, Aranaz A, De Buck J. 2010. PCR amplification and highresolution melting curve analysis as a rapid diagnostic method for genotyping members of the Mycobacterium avium–intracellulare complex.
Clin Microbiol Infect 16:1658-1662.
Chen SC, Yoshida T, Adams A, Thompson KD, Richards RH. 1996. Immune response of rainbow trout to extracellular products of Mycobacterium spp. J Aquat Anim Health 8:216-222.
Choi GE, Lee SM, Yi J, Hwang SH, Kim HH, Lee EY, Cho EH, Kim JH,
Kim HJ, Chang CL. 2010. High-resolution melting curve analysis for
rapid detection of rifampin and isoniazid resistance in Mycobacterium
tuberculosis clinical isolates. J Clin Microbiol 48:3893-3898.
Volume 53, Number 2
2012
Cui Z, Samuel-Shaker D, Watral V, Kent ML. 2010. Attenuated Mycobacterium marinum protects zebrafish against mycobacteriosis. J Fish Dis
33:371-375.
Decostere A, Hermans K, Haesebrouck F. 2004. Piscine mycobacteriosis: A
literature review covering the agent and the disease it causes in fish and
humans. Vet Microbiol 99:159-166.
Devulder G, Pérouse de Montclos M, Flandrois JP. 2005. A multigene approach to phylogenetic analysis using the genus Mycobacterium as a
model. Int J Syst Evol Microbiol 55:293-302.
Ellett F, Lieschke GJ. 2010. Zebrafish as a model for vertebrate hematopoiesis. Curr Opin Pharmacol 10:563-570.
Falkinham JO. 2009. Surrounded by mycobacteria: Nontuberculous mycobacteria in the human environment. J Appl Microbiol 107:356-367.
Falkinham JO, Norton CD, LeChevallier MW. 2001. Factors influencing
numbers of Mycobacterium avium, Mycobacterium intracellulare, and
other mycobacteria in drinking water distribution systems. Appl Environ Microbiol 67:1225-1231.
Ferguson J, Watral V, Schwindt A, Kent ML. 2007. Spores of two fish microsporidia (Pseudoloma neurophilia and Glugea anomola) are highly
resistant to chlorine. Dis Aquat Org 76:205-214.
Francis-Floyd R, Yanong RPE. 1999. Mycobacteriosis in fish. Gainesville:
University of Florida, Institute of Food and Agricultural Sciences; Fact
Sheet VM-96.
Frerichs GN. 1993. Mycobacteriosis: Nocardiosis. In: Inglis V, Roberts RJ,
Bromage NR, eds. Bacterial Diseases of Fish. Oxford: Blackwell.
p 219-233.
Gauthier DT, Rhodes MW. 2009. Mycobacteriosis in fishes: A review. Vet J
180:33-47.
Harriff MJ, Bermudez LE, Kent ML. 2007. Experimental exposure of zebrafish (Danio rerio Hamilton) to Mycobacterium marinum and Mycobacterium peregrinum reveals the gastrointestinal tract as the primary
route of infection: A potential model for environmental mycobacterial
infection. J Fish Dis 30:587-600.
Harriff MJ, Wu M, Kent ML, Bermudez LE. 2008. Species of environmental mycobacteria vary in their ability to grow in human, mouse, and carp
macrophages, associated with the absence of mycobacterial virulence
genes observed by DNA microarray hybridization. Appl Environ Microbiol 74:275-285.
Hedrick RP, McDowell T, Groff J. 1987. Mycobacteriosis in cultured
striped bass from California. J Wildl Dis 23:391-395.
Hegedus Z, Zakrzewska A, Agoston VC, Ordas A, Rácz P, Mink M,
Spaink HP, Meijer AH. 2009. Deep sequencing of the zebrafish transcriptome response to mycobacterium infection. Mol Immunol 46:
2918-2930.
Jacobs JM, Howard DW, Rhodes MR, Newman MW, May EB, Harrell
RM. 2009a. Historical presence (1975–1985) of mycobacteriosis in
Chesapeake Bay striped bass Morone saxatilis. Dis Aquat Organ 85:
181-186.
Jacobs JM, Rhodes MR, Baya A, Reimschuessel R, Townsend H, Harrell
RM. 2009b. Influence of nutritional state on the progression and severity of mycobacteriosis in striped bass Morone saxatilis. Dis Aquat
Organ 87:183-197.
Jacobs J, Rhodes M, Sturgis B, Wood B. 2009c. Influence of environmental
gradients on the abundance and distribution of Mycobacterium spp. in a
coastal lagoon estuary. Appl Environ Microbiol 75:7378-7384.
Jernigan JA, Farr BM. 2000. Incubation period and sources of exposure for
cutaneous Mycobacterium marinum infection: Case report and review
of the literature. Clin Infect Dis 31:439-443.
Kato G, Kato K, Saito K, Pe Y, Kondo H, Aoki T, Hirono I. 2011. Vaccine
efficacy of Mycobacterium bovis BCG against Mycobacterium sp. infection in amberjack Seriola dumerili. Fish Shellfish Immunol 30:467472.
Kawakami K, Kusuda R. 1990. Efficacy of rifampicin, streptomycin and
erythromycin against experimental Mycobacterium infection in cultured yellowtail. B Jpn Soc Sci Fish 56:51-53.
Kent PT, Kubica GP. 1985. Public health mycobacteriology: A guide for the
level III laboratory. Atlanta: US Department of Health and Human Services, CDC.
103
Kent ML, Bishop-Stewart JK. 2003. Transmission and tissue distribution
of Pseudoloma neurophilia (Microsporidia) of zebrafish, Danio rerio
(Hamilton). J Fish Dis 26:423-426.
Kent ML, Matthews JL, Bishop-Stewart JK, Whipps CM, Watral V, Poort
M, Bermudez L. 2004. Mycobacteriosis in zebrafish (Danio rerio)
research facilities. Comp Biochem Physiol C Toxicol Pharmacol 138:
383-390.
Kent ML, Watral V, Wu M, Bermudez L. 2006. In vivo and in vitro growth
of Mycobacterium marinum at homoeothermic temperatures. FEMS
Microbiol Lett 257:69-75.
Kent ML, Spitsbergen JM, Matthews JM, Fournie JW, Westerfield M.
2008. Diseases of zebrafish in research facilities. Available online at
http://zebrafish.org/zirc/health/diseaseManual.php (accessed October 8,
2012).
Kent ML, Feist SW, Harper C, Hoogstraten-Miller S, Law JM, SánchezMorgado JM, Tanguay RL, Sanders GE, Spitsbergen JM, Whipps CM.
2009. Recommendations for control of pathogens and infectious diseases in fish research facilities. Comp Biochem Physiol C Toxicol Pharmacol 149:240-248.
Kent ML, Buchner C, Watral VG, Sanders JL, LaDu J, Peterson TS, Tanguay
RL. 2011. Development and maintenance of a specific pathogen free
(SPF) zebrafish research facility for Pseudoloma neurophilia. Dis Aquat
Org 95:73-79.
Kent ML, Harper C, Wolf JC. 2012. Documented and potential research
impacts of subclinical diseases in zebrafish. ILAR J 53:126-134.
Kim H, Kim SH, Shim TS, Kim MN, Bai GH, Park YG, Lee SH, Chae GT,
Cha CY, Kook YH, Kim BJ. 2005. Differentiation of Mycobacterium
species by analysis of the heat-shock protein 65 gene (hsp65). Int J Syst
Evol Microbiol 55:1649-1656.
Lawrence C. 2007. The husbandry of zebrafish (Danio rerio): A review.
Aqaculture 269:1-20.
Lesley R, Ramakrishnan L. 2008. Insights into early mycobacterial pathogenesis from the zebrafish. Curr Opin Microbiol 11:277-83.
Lieschke GJ, Currie PD. 2007. Animal models of human disease: Zebrafish
swim into view. Nat Rev Genet 8:353-367.
Loeschke S, Goldmann T, Vollmer E. 2005. Improved detection of mycobacterial DNA by PCR in formalin-fixed, paraffin-embedded tissues using thin sections. Pathol Res Pract 201:37-40.
Mainous ME, Smith SA. 2005. Efficacy of common disinfectants against
Mycobacterium marinum. J Aquat Anim Health 17:284-288.
McNabb A, Adie K, Rodrigues M, Black WA, Isaac-Renton J. 2006. Direct
identification of mycobacteria in primary liquid detection media by partial sequencing of the 65-kilodalton heat shock protein gene. J Clin Microbiol 44:60-66.
Murray KN, Bauer J, Tallen A, Matthews JL, Westerfield M, Varga ZM.
2011. Characterization and management of asymptomatic Mycobacterium infections at the Zebrafish International Resource Center. JAALAS
50:675-679.
Noga E. 2010. Fish Disease: Diagnosis and Treatment. Malden MA: Blackwell Publishing.
Norton W, Bally-Cuif L. 2010. Adult zebrafish as a model organism for
behavioural genetics. BMC Neurosci 11:90.
Ostland VE, Watral V, Whipps CM, Austin FW, St-Hilaire S, Westerman
ME, Kent ML. 2008. Biochemical, molecular, and virulence studies of
select Mycobacterium marinum strains in hybrid striped bass (Morone
chrysops x M. saxatilis) and zebrafish (Danio rerio). Dis Aquat Org
79:107-118.
Parisot TJ. 1958. Tuberculosis in fish: A review of the literature with a
description of the disease in salmonid fish. Bacteriol Rev 22:240-245.
Pasnik DJ, Smith SA. 2005. Immunogenic and protective effects of a DNA
vaccine for Mycobacterium marinum in fish. Vet Immunol Immunopathol 103:195-206.
Pourahmad F, Thompson KD, Adams A, Richards RH. 2009. Detection and
identification of aquatic mycobacteria in formalin-fixed, paraffinembedded fish tissues. J Fish Dis 32:409-419.
Ramsay JM, Feist GW, Varga ZM, Westerfield M, Kent ML, Schreck CB.
2009a. Whole-body cortisol response of zebrafish to acute net handling
stress. Aquaculture 297:157-162.
104
Ramsay JM, Watral V, Schreck CB, Kent ML. 2009b. Husbandry stress exacerbates mycobacterial infections in adult zebrafish, Danio rerio
(Hamilton). J Fish Dis 32:931-941.
Ringuet H, Akoua-Koffi C, Honore S, Varnerot A, Vincent V, Berche P,
Gaillard JL, Pierre-Audigier C. 1999. hsp65 Sequencing for identification of rapidly growing mycobacteria. J Clin Microbiol 37:852-857.
Ross AJ. 1970. Mycobacteriosis among Pacific salmonid fishes. In: Sniesko
SF, ed. A symposium on diseases of fishes and shellfishes. Washington:
American Fisheries Society. p 279-283.
Roth A, Reischl U, Streubel A, Naumann L, Kroppenstedt RM, Habicht M,
Fischer M, Mauch H. 2000. Novel diagnostic algorithm for identification of mycobacteria using genus-specific amplification of the 16S-23S
rRNA gene spacer and restriction endonucleases. J Clin Microbiol
38:1094-1104.
Sampaio JL, Viana-Niero C, de Freitas D, Hofling-Lima AL, Leao SC.
2006. Enterobacterial repetitive intergenic consensus PCR is a useful
tool for typing Mycobacterium chelonae and Mycobacterium abscessus
isolates. Diagn Microbiol Infect Dis 55:107-118.
Sanders J, Watral V, Kent M. 2012. Microsporidiosis in zebrafish research
facilities. ILAR J 53:106-113.
Schulze-Röbbecke R, Janning B, Fischer R. 1992. Occurrence of mycobacteria in biofilm samples. Tubercle Lung Dis 73:141-144.
Selvaraju SB, Khan IU, Yadav JS. 2005. A new method for species identification and differentiation of Mycobacterium chelonae complex based
on amplified hsp65 restriction analysis (AHSPRA). Mol Cell Probes
19:93-99.
Spence R, Gerlach G, Lawrence C, Smith C. 2008. The behaviour and
ecology of the zebrafish, Danio rerio. Biol Rev Camb Philos Soc
83:13-34.
Stanley KA, Curtis LR, Simonich SL, Tanguay RL. 2009 Endosulfan I and
endosulfan sulfate disrupts zebrafish embryonic development. Aquat
Toxicol 95:355-361.
Steed KA, Falkinham JO 3rd. 2006. Effect of growth in biofilms on chlorine
susceptibility of Mycobacterium avium and Mycobacterium intracellulare. Appl Environ Microbiol 72:4007-4011.
Talaat AM, Reimschuessel R, Trucksis M. 1997. Identification of mycobacteria infecting fish to the species level using polymerase chain reaction
and restriction enzyme analysis. Vet Microbiol 58:229-237.
Telenti A, Marchesi F, Balz M, Bally F, Böttger EC, Bodmer T. 1993.
Rapid identification of mycobacteria to the species level by polymerase chain reaction and restriction enzyme analysis. J Clin Microbiol 31:175-178.
Tobin DM, Vary JC Jr, Ray JP, Walsh GS, Dunstan SJ, Bang ND, Hagge
DA, Khadge S, King MC, Hawn TR, Moens CB, Ramakrishnan L.
2010. The lta4h locus modulates susceptibility to mycobacterial infection in zebrafish and humans. Cell 140:717-730.
Tortoli E. 2003. Impact of genotypic studies on mycobacterial taxonomy: The new mycobacteria of the 1990s. Clin Microbiol Rev 16:
319-354.
van der Sar AM, Appelmelk BJ, Vandenbroucke-Grauls CM, Bitter W.
2004. A star with stripes: Zebrafish as an infection model. Trends Microbiol 12:451-457.
van der Sar AM, Spaink HP, Zakrzewska A, Bitter W, Meijer AH. 2009.
Specificity of the zebrafish host transcriptome response to acute and
chronic mycobacterial infection and the role of innate and adaptive immune components. Mol Immunol 46:2317-2332.
Vanitha JD, Venkatasubramani R, Dharmalingam K, Paramasivan CN.
2003. Large-restriction-fragment polymorphism analysis of Mycobacterium chelonae and Mycobacterium terrae isolates. Appl Environ Microbiol 69:4337-4341.
Watral V, Kent ML. 2007. Pathogenesis of Mycobacterium spp. in zebrafish
(Danio rerio) from research facilities. Comp Biochem Physiol C Toxicol Pharmacol 145:55-60.
Westerfield M. 2000. The Zebrafish Book. A Guide for the Laboratory
Use of Zebrafish (Danio rerio), 4th ed. Eugene: University of Oregon
Press.
Whipps CM, Butler WR, Pourahmad F, Watral VG, Kent ML. 2007a. Molecular systematics support the revival of Mycobacterium salmoniphilum
ILAR Journal
(ex Ross 1960) sp. nov., nom. rev., a species closely related to
Mycobacterium chelonae. Int J Syst Evol Microbiol 57:25252531.
Whipps CM, Dougan ST, Kent ML. 2007b. Mycobacterium haemophilum
infections of zebrafish (Danio rerio) in research facilities. FEMS Microbiol Lett 270:21-26.
Whipps CM, Matthews JL, Kent ML. 2008. Distribution and genetic characterization of Mycobacterium chelonae in laboratory zebrafish (Danio
rerio). Dis Aquat Organ 82:45-54.
Volume 53, Number 2
2012
Yanong RP, Pouder DB, Falkinham JO 3rd. 2010. Association of mycobacteria in recirculating aquaculture systems and mycobacterial disease in
fish. J Aquat Anim Health 22:219-223.
Zerihun MA, Hjortaas MJ, Falk K, Colquhoun DJ. 2011. Immunohistochemical and Taqman real-time PCR detection of mycobacterial infections in fish. J Fish Dis 34:235-246.
Zhang Y, Rajagopalan M, Brown BA, Wallace RJ Jr. 1997. Randomly amplified polymorphic DNA PCR for comparison of Mycobacterium abscessus
strains from nosocomial outbreaks. J Clin Microbiol 35:3132-3139.
105