INFECTION AND IMMUNITY, Nov. 2006, p. 6108–6117
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Vol. 74, No. 11
Mycobacterium marinum Infection of Adult Zebrafish Causes Caseating
Granulomatous Tuberculosis and Is Moderated by Adaptive Immunity†
Laura E. Swaim,1‡ Lynn E. Connolly,2‡* Hannah E. Volkman,3Olivier Humbert,1§
Donald E. Born,4and Lalita Ramakrishnan1,2,5
Departments of Microbiology,1Medicine,2Immunology,5and Pathology4and Molecular and
Cellular Biology Graduate Program,3University of Washington, Seattle, Washington 98195
Received 5 June 2006/Returned for modification 6 July 2006/Accepted 31 July 2006
The zebrafish, a genetically tractable model vertebrate, is naturally susceptible to tuberculosis caused by
Mycobacterium marinum, a close genetic relative of the causative agent of human tuberculosis, Mycobacterium
tuberculosis. We previously developed a zebrafish embryo-M. marinum infection model to study host-pathogen
interactions in the context of innate immunity. Here, we have constructed a flowthrough fish facility for the
large-scale longitudinal study of M. marinum-induced tuberculosis in adult zebrafish where both innate and
adaptive immunity are operant. We find that zebrafish are exquisitely susceptible to M. marinum strain M.
Intraperitoneal injection of five organisms produces persistent granulomatous tuberculosis, while the injection
of ?9,000 organisms leads to acute, fulminant disease. Bacterial burden, extent of disease, pathology, and host
mortality progress in a time- and dose-dependent fashion. Zebrafish tuberculous granulomas undergo caseous
necrosis, similar to human tuberculous granulomas. In contrast to mammalian tuberculous granulomas,
zebrafish lesions contain few lymphocytes, calling into question the role of adaptive immunity in fish tuber-
culosis. However, like rag1 mutant mice infected with M. tuberculosis, we find that rag1 mutant zebrafish are
hypersusceptible to M. marinum infection, demonstrating that the control of fish tuberculosis is dependent on
adaptive immunity. We confirm the previous finding that M. marinum ?RD1 mutants are attenuated in adult
zebrafish and extend this finding to show that ?RD1 predominantly produces nonnecrotizing, loose macro-
phage aggregates. This observation suggests that the macrophage aggregation defect associated with ?RD1
attenuation in zebrafish embryos is ongoing during adult infection.
Human tuberculosis, caused by Mycobacterium tuberculosis,
results in a variety of outcomes that range from long-term
asymptomatic infection to active disease (9, 14). The hallmark
lesion of tuberculosis is the granuloma, an organized collection
of differentiated macrophages: large epithelioid cells with
tightly interdigitated cell membranes linking adjacent cells to-
gether (1). In addition, human tuberculous granulomas often
contain lymphocytes and extracellular matrix and can undergo
central necrosis that correlates with the gross appearance of
caseous necrosis (1, 14). Necrotizing granulomas containing
organisms are seen in active human tuberculosis (14, 15, 53).
Chronic asymptomatic tuberculous infection of humans is
characterized by fibrotic and calcified granulomas that may or
may not contain viable bacteria (35). Necrosis likely plays an
important role in tuberculosis morbidity and transmission, but
the host and bacterial determinants required for its formation
remain poorly understood (14).
Several animal models are used to study the pathogenesis
and immunology of M. tuberculosis infection, each of which has
its advantages and drawbacks (20). The commonly used mouse
model produces poorly organized macrophage aggregates that
do not undergo necrosis but has the advantage of abundant
immunological and genetic tools and techniques (20). Guinea
pigs and rabbits produce necrotizing granulomas (2, 15, 16, 29),
but very few immunological and molecular reagents are avail-
able for these animals, which are also more expensive to main-
tain. Nonhuman primates appear to most faithfully mimic hu-
man tuberculosis (8), but their use is constrained by ethical and
cost issues. These differences notwithstanding, cell-mediated
adaptive immunity plays a critical role in the control of human
tuberculosis and in all experimental mammalian models of
tuberculosis where this has been assessed (21, 34).
The relatively rapidly growing human and animal pathogen
Mycobacterium marinum, a close genetic relative of M. tuber-
culosis (49), is used to study the pathogenesis of tuberculosis
(11, 14, 39, 46, 55). M. marinum causes systemic granulomatous
infections and disease in its natural hosts, ectotherms such as
fish and frogs, and peripheral chronic granulomatous disease
(fish tank granulomas) in humans (10, 11, 19). Inoculation of
the leopard frog produces a long-term, asymptomatic infection
with well-defined, nonnecrotizing granulomas that harbor bac-
teria (5, 41). In contrast, experimental inoculation of goldfish,
zebrafish, striped bass, or medaka produces necrotizing gran-
ulomas and ultimately lethal disease (7, 11, 12, 24, 39, 40, 48,
53, 54, 59).
The zebrafish is a commonly used laboratory animal for the
study of development and disease (4, 50). Zebrafish are genet-
ically tractable hosts: transgenic animals and random and tar-
geted mutants can be created (14, 27, 31, 36, 58), its genome
sequence is nearing completion (43), and microarray analysis
using mRNA extracted from whole fish can be readily carried
* Corresponding author. Mailing address: Box 357242, University of
Washington, Seattle, WA 98195. Phone: (206) 221-6367. Fax: (206)
616-1575. E-mail: email@example.com.
† Supplemental material for this article may be found at http://iai
‡ L.E.S. and L.E.C. contributed equally to this work.
§ Present address: Molecular and Cellular Biology Graduate Pro-
gram, University of Washington, Seattle, WA 98195.
out (30). Chemical and drug screens are also easily performed
using this organism (37, 38). Zebrafish become infected with
M. marinum in fish facilities (57) and also develop disease upon
experimental inoculation (11, 12, 22, 23, 40, 54). A zebrafish
embryo model of infection has been used to characterize the
earliest events following mycobacterial infection in which mac-
rophage aggregates with key features of tuberculous granulo-
mas form (18). Because adaptive immunity is not yet operant
at this early stage of development, the interactions of innate
immunity with mycobacteria and their virulence determinants
can be dissected in isolation (13, 52, 56).
Here, we describe a large-scale longitudinal analysis of M.
marinum infection of adult zebrafish using the sequenced hu-
man strain M (42). Similar to a previous study using a fish-
derived strain (40), we find a dose- and time-dependent mor-
tality that is accompanied by increased bacterial burden and
progressive granulomatous disease. In this study, we used in-
ocula that are 2 to 5 logs lower than those reported previously
by other groups (7, 22, 23, 40, 54) to demonstrate the exquisite
sensitivity of the zebrafish to infection by M. marinum strain M.
Although zebrafish possess adaptive immunity similar to that
of mammals (50, 52), its role in controlling tuberculosis has not
previously been determined. By infecting mutant zebrafish de-
ficient in rag1 (58), a recombinase required for the develop-
ment of functional T and B cells, we demonstrate that adaptive
immunity plays a critical role in controlling zebrafish tubercu-
losis as it does in mammalian tuberculosis (44). Finally, we find
that an attenuated bacterial strain lacking the RD1 locus,
which encodes a novel bacterial secretion system and displays
a macrophage aggregation defect in zebrafish embryos (56),
exhibits a similar defect in adult animals. This finding suggests
that the RD1 locus plays similar roles throughout infection
both in the context of innate immunity alone and in the pres-
ence of an adaptive immune response.
MATERIALS AND METHODS
Bacterial strains and growth conditions. Mycobacterium marinum strains used
for this study were derived from a human clinical isolate, strain M (ATCC
BAA-535), and were grown at 33°C in Middlebrook 7H9 broth (Difco) supple-
mented with 0.5% bovine serum albumin, 0.005% oleic acid, 0.2% glucose, 0.2%
glycerol, 0.085% sodium chloride, and 0.05% Tween 80 or on Middlebrook 7H10
agar (Difco) supplemented with 0.5% bovine serum albumin, 0.005% oleic acid,
0.2% glucose, 0.2% glycerol, and 0.085% sodium chloride. Infected fish homog-
enates were plated onto 7H10 agar supplemented with amphotericin B (10
mg/liter), polymyxin B (20 mg/liter), trimethoprim (20 mg/liter), and carbenicillin
(50 mg/liter) to avoid contamination with normal flora. Cultures used in infec-
tions were grown to an optical density at 600 nm of 1.0 and maintained at ?80°C
in 1-ml aliquots with 10% glycerol. CC1, a wild-type (WT) strain (12), and ?RD1
(56), an attenuated strain, have been described previously.
Fish husbandry and genotyping. Zebrafish husbandry has been detailed pre-
viously by Cosma et al. (11). Briefly, the fish were reared in recirculating fish
systems obtained from Aquatic Habitats (Apopka, FL) and transferred to a
flowthrough fish system for the infection experiment. The flowthrough system
prevents reinfection by allowing water to pass continuously through the tanks.
Up to 12 fish were kept in each 9-liter tank, and tanks were maintained under
conditions compliant with laboratory standards outlined by the Institutional
Animal Care and Use Committee (water temperature of ?28°C, pH of ?7.4, and
conductivity of ?1,500 ?S).
Wild-type (family AB) founder zebrafish were originally obtained from David
Raible’s fish facility at the University of Washington. Heterozygous rag1 mutant
zebrafish (family Tu ¨bingen) obtained from Artemis Pharmaceuticals (Ko ¨ln, Ger-
many) (58) were spawned to produce isogenic siblings that were predicted to be
approximately one-quarter mutant:one-half heterozygous:one-quarter wild type
for rag1. Their genotype was determined at 3 to 6 months of age using DNA
obtained from a tail clip procedure (57). Briefly, a small piece of fin was clipped
from the fish and incubated in lysis buffer (100 mM Tris-HCl, 200 mM NaCl,
0.2% sodium dodecyl sulfate, 5 mM EDTA, 100 ?g/ml proteinase K) overnight
at 55°C. DNA was precipitated from the supernatant with isopropanol and then
resuspended in water. To screen for the point mutation R797 that gives rise to a
truncated Rag1 protein (58), the DNA was amplified by PCR with primers seq2F
(5?-CCCTGACAGCCATCTTGGGAC-3?) and seq2R (5?-CGTGTCATCAGC
CGACGAGC-3?). The 600-bp PCR product obtained was sequenced using the
seq2F primer. The resulting sequence chromatograms were analyzed for the
expected single nucleotide polymorphism (thymine-to-cytosine replacement) at
one or both alleles.
Zebrafish infections and bacterial quantification. After lightly anesthetizing
fish with 0.1% 3-aminobenzoic acid ethyl ester (tricaine), they were infected by
intraperitoneal injection with 50 ?l of thawed bacterial stocks that were diluted
in phosphate-buffered saline (PBS) to obtain the desired inoculum. Bacterial
CFU contained in the injected inoculum were confirmed at the time of infection
by plating onto 7H10 agar. To assess the bacterial burden from whole fish at
various time points postinfection, the fish were first bathed in 1.5 mg/ml kana-
mycin sulfate for 45 min at 27°C to inhibit contamination with normal flora.
Following terminal anesthesia in 0.5% tricaine, fish were weighed and homoge-
nized (model TH-115; Omni International, Marietta, GA) in 1.5 ml PBS in
sterile polystyrene tubes. The homogenates were brought to a volume of 2.5 ml
with PBS, and 10-fold serial dilutions were plated. Four to eight fish were plated
for each condition per time point, and the resulting bacterial counts or CFU were
Histology and microscopy. After terminal anesthesia, fish were fixed in
Dietrich’s fixative (57) for 72 h, transferred into 70% ethanol, and then sent to
Histo-Tec Laboratories (Hayward, CA) for paraffin embedding, sectioning, and
staining. Serial sagittal sections were used for hematoxylin and eosin and mod-
ified Ziehl-Neelson staining in all cases and additionally for Masson’s modified
trichrome (collagen) and Van Kossa (calcium) staining in some cases. Two to five
fish from each infection dose at each time point were examined. Sections were
examined under a Nikon E600 microscope (Tokyo, Japan), and images were
collected with a digital photo camera (model DKC-5000; Sony, Tokyo, Japan)
and produced using Metamorph software (Molecular Devices Corporation,
Sunnyvale, CA). Final images were transferred to Adobe (San Jose, CA) Pho-
toshop 7.0 to adjust levels and brightness (12, 56).
For a quantitative comparison of ?RD1 and wild-type lesions, a blinded
reviewer scored 121 granulomas from four wild-type-infected fish at 16 weeks
and 126 granulomas from three ?RD1-infected fish at 16 weeks in all tissues
(kidney, liver, mesentery, pancreas, gonads, spleen, and head). All granulomas
were assessed for the presence of necrosis and residence in a multicentric lesion.
Nonnecrotizing lesions were further analyzed to assess the compactness of the
macrophage association. The data were analyzed using contingency tables, as-
signing ?RD1-infected fish as the experimental group and wild-type-infected fish
as the control group, to determine the relative risk (RR) and 95% confidence
intervals (CI). Significance was determined by Fisher’s exact test.
Statistical analysis. Statistics were calculated using GraphPad Instat version
3.05 and Stata version 9.0. For the analysis of the bacterial growth curves, the
data were found to not be normally distributed despite log transformation, so
nonparametric tests (Mann-Whitney test for comparisons of two groups and
Kruskal-Wallis test for comparisons between three or more groups) were used to
determine significance. Medians were calculated using all fish, including those
with bacterial burdens below the limit of detection.
M. marinum causes chronic, progressive disease of zebrafish
in a dose-dependent fashion. Adult zebrafish were injected
with three doses of M. marinum or with PBS (mock), and
survival and disease progression were assessed in two different
cohorts of injected fish. Similar to previously published find-
ings (40), a dose-dependent mortality was noted over the 16-
week monitoring period (Fig. 1A). Most of the fish infected
with 8,970 bacteria died within 2 weeks, whereas only 44% of
the fish infected with 5 CFU of M. marinum died within the
16-week monitoring period. The fish infected with 60 CFU
exhibited an intermediate mortality, with 83% being dead by
16 weeks. Only 11% of the mock-injected fish died during the
16-week period. Death of infected fish was preceded by re-
VOL. 74, 2006M. MARINUM INFECTION OF ADULT ZEBRAFISH6109
duced feeding and weakened swimming for approximately 1
week. The fish either were often listless at the bottom of the
tank or exhibited listless piping, remaining at the surface, con-
stantly opening and closing their mouths in an attempt to
increase gas exchange (32). Dying fish also frequently had
external red lesions on the trunk ventral to the lateral line and
in close proximity to internal organs, ascites, and exophthal-
mia, the latter two signs being indicators of kidney failure and
osmoregulatory stress (47).
Disease progression was assessed in the second cohort of
infected fish by serial bacterial counts (Fig. 1B to D) and
histopathology at 1, 8, and 11 days for the highest dose and 1,
2, 4, 8, and 16 weeks postinfection for the two lower doses. The
small size of the zebrafish allowed a determination of total
bacterial counts in the whole animal without separating organs
(40). For all doses, bacterial counts increased to reach a pla-
teau of 6 to 7 logs at times postinfection that correlated in-
versely with the infecting dose (Fig. 1B and C). A few of the
survivors had overt signs of disease, while the others ap-
peared healthy. Of the fish that survived to the 16-week time
point, two in the group infected with 5 CFU had counts that
were below 3.4 logs, the level of detection in these experi-
ments, while two other fish (one from each group) had
counts that were markedly lower than those of the other fish
FIG. 1. Host death correlates with bacterial burden and disease dissemination. (A) Dose-dependent survival of adult zebrafish infected with
wild-type M. marinum. Three separate tanks, each containing 12 fish at the indicated doses, were monitored for mortalities over a 16-week time
period. No significant tank effects were detected, and the data from the three tanks were pooled for further analysis. A Kaplan-Meier curve was
calculated for each dose as well as for mock-injected animals. Compared to mock-injected animals, each dose showed a significant difference in
survival as calculated by log-rank test (P ? 0.001). The Cox proportional hazards model was used to quantify group differences compared to
mock-injected fish and confirmed significant differences in survival at each dose with the following hazard ratios (HR), 95% confidence intervals
(CI), and P values: for the 5-CFU group, the HR was 4.85, the 95% CI was 1.6 to 14.5, and the P value was ?0.005; for the 60-CFU group, the
HR was 14.27, the 95% CI was 5.0 to 40.8, and the P value was ?0.0001; for the 8.6 ? 103-CFU group, the HR was 86.49, the 95% CI was 28.6
to 261.5, and the P value was ?0.0001. (B and C) Dose-dependent growth of M. marinum in adult zebrafish. Fish in B were inoculated with 5 CFU
(■) or 60 CFU (F) of WT M. marinum, while fish in C were inoculated with 8,970 CFU (Œ), and bacterial counts were enumerated at the indicated
time points. The limit of detection in these experiments (3.4 log CFU per fish) is indicated by the dashed line, and the number of fish falling below
this limit at each time point is designated by a symbol underneath this line. Multiple fish at a given time point having the same bacterial counts
are shown as a white numeral within the symbols (■, F, or Œ). Median CFU are shown as overlying bar graphs (white for the 5-CFU group, dark
gray for the 60-CFU group, and light gray for 8,970-CFU group). Significant differences in median bacterial loads at each time point were
determined using the Mann-Whitney test (**, P ? 0.05). (D) Fish infected with either 5 CFU (■) or 60 CFU (F) of M. marinum were killed at
the indicated time periods, and the median number of tissues containing granulomas was determined. Each individual fish is represented as a single
point, and the medians are shown as overlying bar graphs (white for the 5-CFU group and dark gray for the 60-CFU group). Multiple fish at a
given time point having the same number of tissues involved are shown as a white numeral within the symbols (■ or F). Tissues examined included
liver, kidney, pancreas, gonads, and mesentery. The significance in the differences in the extent of granulomatous disease between the two doses
was determined using the Mann-Whitney test (**, P ? 0.05).
6110SWAIM ET AL. INFECT. IMMUN.
(Fig. 1B). Because these fish were sampled from the surviv-
ing population, which was likely enriched for fish that may
have been misinjected, it was probable that these fish were
never infected with M. marinum. This notion was further
supported by the finding that three of the surviving fish in
the 5-CFU group showed no evidence of active or healing
infection by histology, while the remainder of the fish in
both groups showed extensive granulomatous disease (data
not shown). The two fish with very low bacterial counts may
have been initially uninfected and may have become in-
fected later from dying fish in the same tank, although we
cannot rule out the possibility that they were in the process
of clearing the infection. The median bacterial counts be-
tween 4, 8, and 16 weeks of infection in the 60-CFU group
and 8 and 16 weeks in the 5-CFU group were not signifi-
cantly different by analysis of variance (ANOVA). Similarly,
the bacterial burden in the 8,970-CFU group did not change
significantly between 8 and 11 days and was not significantly
different from the final counts achieved in the 5- and 60-
CFU groups, as determined by ANOVA, confirming that the
bacterial counts plateau to a fixed level, regardless of the
The extent of M. marinum granulomatous disease is time-
and dose-dependent. The small size of the zebrafish permitted
a histological assessment of the progression and dissemination
of the infection in the entire fish by examining sagittal sections
in which the organs were well represented. Serial sections
could be used for different histological stains of the same
granulomas. In the 5- and 60-CFU groups, the extent of gran-
ulomatous pathology increased over time and correlated with
progressive mortality and an increase in bacterial burden (Fig.
1A, C, and D). At 1 week, granulomas were evident in only one
fish in each group, whereas at later time points, granulomas
were readily detected in most fish. Granulomas were fre-
quently found in the mesentery, pancreas, gonads, kidney, and
liver but rarely in the head, branchial arches, and spleen and
never in the muscle or skin.
The extent of disease was scored by assessing the number of
regions and/or organs involved by granulomatous disease in
each fish. The number of regions involved increased with time
for the 60-CFU dose (Fig. 1D). By week 4, there was a signif-
icant difference in the extent of disease between the 5- and
60-CFU doses, mirroring the differences in bacterial CFU and
host survival between these doses at this time point (Fig. 1B).
Owing to mortalities by the 8-week time point, only two fish
were available for histological analysis in the 5-CFU group,
and none were available in the 60-CFU group. These two fish
both had five regions involved, similar to the extent of disease
seen in the 60-CFU group at 4 weeks. This increase in disease
burden in the fish that received the lower dose again mirrors
the pattern in the bacterial counts, with increasing bacterial
burden correlating with disease dissemination (Fig. 1B and D).
Consistent with the accelerated mortality and bacterial counts
seen in the fish infected with 8,970 CFU, this inoculum pro-
duced extensive disease involving most regions by 8 days
postinfection (data not shown).
Progression of granuloma pathology is also both time- and
dose-dependent. A detailed analysis of progression of granu-
loma pathology was performed for the fish infected with 5
CFU. For the first 2 weeks after infection, nonnecrotizing
lesions were predominant, and early macrophage aggregates
with cytoplasmic and nuclear debris, likely representing early
necrosis, were seen (Fig. 2A and B). By 16 weeks, approxi-
mately equal numbers of well-organized but nonnecrotizing
and necrotizing granulomas were present (Fig. 2C to F). The
clear areas in Fig. 2C likely reflect either the beginnings of
cellular breakdown to form necrotic zones or less interdigita-
tion of the cells. An example of an early necrotic focus is shown
in Fig. 2E and F, where cellular remnants are seen in the
partially necrotic area but not in the completely necrotic area.
The bacteria were found predominantly in necrotic areas and
were in greater numbers in fully necrotic than in partially
necrotic zones (Fig. 2F). As a result, necrotizing lesions had far
greater numbers of bacteria than did nonnecrotizing granulo-
mas (Fig. 2, compare B and D to F and H). At 16 weeks,
necrotic lesions predominated, and thin-walled lesions with
prominent necrosis were found (Fig. 2G and H), as has been
previously reported for human granulomas in the setting of
active disease (53). Also prominent at this time point were
multicentric granulomas (Fig. 2I and J), which again were
similar to lesions found in late-stage human tuberculosis (53).
Progression to necrotizing infection was also dose dependent,
appearing by day 8 in the 60- and 8,970-CFU groups and being
predominant at the highest dose at both days 8 and 11 (see Fig.
S1 in the supplemental material).
Despite extensive examination of numerous fish at all doses
and time points, we found no lesions similar to those seen
during chronic asymptomatic tuberculous infection of humans
as determined by Von Kossa staining for the presence of cal-
cification and trichrome staining for the presence of fibrosis
(data not shown). A feature specific to fish and frogs was the
accumulation of pigment in the later stages of disease (Fig. 2I
and J). This pigment is likely associated with melanomacro-
phages. Melanomacrophages are capable of phagocytosing and
eliminating bacterial pathogens in other fish diseases (6), but
their potential role in mycobacterial disease pathogenesis re-
mains unexplored (5, 33, 48).
Similar to frog and goldfish lesions, zebrafish M. marinum
granulomas contained few lymphocytes as judged by hematox-
ylin and eosin staining (compare Fig. 2A, C, and E). This
feature is in contrast to M. tuberculosis granulomas in mice,
humans, and nonhuman primates and M. marinum granulomas
in humans (14). The abundance of lymphocytes does not ap-
pear to be related to the presence of necrosis, as zebrafish,
goldfish, and frog M. marinum granulomas all show few lym-
phocytes, yet the former two hosts readily form necrotizing
Zebrafish lacking rag1 are hypersusceptible to M. marinum
infection. Cell-mediated adaptive immunity plays a critical pro-
tective role in mammalian tuberculosis (21), and the paucity of
lymphocytes in zebrafish granulomas suggested that the role of
the adaptive immune response to mycobacteria might be less
important in this host. The recent availability of a targeted rag1
knockout fish (58) allowed us to test the role of lymphocytes in
protection against mycobacterial infection in the fish. Fish
lacking rag1 do not undergo VDJ recombination and thus lack
functional T and B cells (58). We infected isogenic rag1?/?,
rag1?/?, and rag1?/?zebrafish siblings with M. marinum and
compared their survival rates and disease pathologies. The
genotype of each fish was confirmed prior to infection, and the
VOL. 74, 2006 M. MARINUM INFECTION OF ADULT ZEBRAFISH6111
FIG. 2. Zebrafish tuberculosis disease progression is characterized by progressive necrosis and pigment deposition with large numbers of
bacteria found within necrotic granulomas. Adult zebrafish were infected with 5 CFU of wild-type M. marinum, and histology was assessed at 2
weeks (A and B), 8 weeks (C, D, E, and F), and 16 weeks (G, H, I, and J) postinfection with hematoxylin and eosin (A, C, E, G, and I) and
6112 SWAIM ET AL.INFECT. IMMUN.
rag1 mutant fish raised in our facility appeared healthy but did
not spawn as well as their wild-type or heterozygote siblings.
In four experiments using fish spawned at different times and
a range of M. marinum infection doses (8 to 54 CFU), rag1?/?
fish showed significantly increased mortality compared to their
rag1?/?and rag1?/?siblings (Fig. 3). Fish that died had ex-
tensive granulomatous disease that was the likely the cause of
their death. This observation shows that, similar to mammalian
hosts infected with M. tuberculosis, adaptive immunity plays a
critical role in protection against pathogenic mycobacteria in
A small cohort of isogenic rag1?/?, rag1?/?, and rag1?/?
zebrafish siblings was infected with 25 CFU of M. marinum to
compare the pathologies at early time points. Infected fish
were examined at 6 weeks, and all showed evidence of granu-
loma formation. Similar to heterozygous and wild-type fish,
rag1 mutant fish formed both nonnecrotizing and necrotizing
lesions (data not shown). While the number of fish available
for this assessment was too small to make quantitative com-
parisons of the histopathology of rag1 mutant and wild-type
fish, these preliminary data suggest that granuloma formation
and necrosis are independent of adaptive immunity.
M. marinum ?RD1 attenuation in adult zebrafish is associ-
ated with defects in granuloma formation. To further validate
our model, we next sought to determine if bacterial mutants
that are attenuated in other model systems are similarly atten-
uated in adult zebrafish. M. tuberculosis strains with mutations
in the RD1 region are attenuated for growth and virulence in
a mouse infection model (26, 28, 45), and RD1-deficient M.
marinum strains are attenuated in frogs and zebrafish (22, 56).
In the zebrafish embryo infection model, macrophages infected
with ?RD1 show an impaired ability to aggregate (56). We
wished to determine if this impaired macrophage aggregation
correlated with the attenuation of the ?RD1 mutant when
infection was initiated in the context of adaptive immunity. We
also wished to address the possibility that macrophage lineages
could be distinct in adult and embryonic fish so that the defects
observed in the embryo might be specific to bacterial interac-
tions with embryonic macrophages.
Fish were infected with 1,318 CFU of M. marinum ?RD1 or
50 CFU of wild-type M. marinum, and host survival was mon-
itored over a 16-week period. Similar to previous findings (22),
?RD1 caused significantly delayed mortality despite its 26-
fold-higher infection dose (Fig. 4A), underscoring the severe
attenuation of the ?RD1 strain previously reported in a variety
of animal models. The bacterial burden of the ?RD1-infected
survivors at 16 weeks reached a median of 5.71 logs (range,
5.32 to 7.23 logs) and, by ANOVA, was found to be not sig-
nificantly different from the plateau of bacterial counts at-
tained for all three doses of wild-type infection described
above (Fig. 1B and C). This result is in contrast to a previous
report that showed lower bacterial counts in fish infected with
RD1 mutant bacteria than in fish infected with wild-type bac-
teria at early time points (22) but is in agreement with the
observation that mice infected with ?RD1 M. tuberculosis
eventually reach high bacterial burdens if observed long
enough, with some resultant host mortality (45).
?RD1 infection of 1-day-old zebrafish embryos results in
delayed macrophage aggregation, and this macrophage aggre-
gation defect persists in fish reared to 1 month of age as
evidenced by loose, poorly formed, nonnecrotic aggregates
(56). To determine if the attenuation phenotype of the ?RD1
strain seen in adult fish was also associated with these histo-
logical differences, individual wild-type and ?RD1 granulomas
were scored in a blinded fashion for the presence of necrosis,
whether they were solitary granulomas or in multicentric ag-
gregates, and, in the case of nonnecrotizing granulomas,
whether the cellular association was loose or compact (Fig. 4B,
C, and D). Because the cohort of fish infected with wild-type
bacteria at the same time as the ?RD1 strain did not survive to
be included in this analysis, we instead compared the ?RD1
lesions to lesions from the 16-week survivors of the wild-type-
infected fish described above. Although the two groups of fish
Ziehl-Neelson (B, D, F, H, and J) staining of serial sections. (A and B) Early, loosely associated, nonnecrotic lesions (arrows) with early cellular
breakdown in the gonad (teste) composed of an epithelioid macrophage infiltrate surrounding scattered mycobacteria. The inset in B shows a
higher-magnification view of acid-fast bacteria residing within macrophages. (C and D) Well-organized, nonnecrotizing granuloma in the liver with
clear areas between macrophages reflecting either early necrosis or less interdigitation of the macrophages. (E and F) Necrotizing granuloma in
the liver with areas of partial (dashed lines) and complete (arrow) necrosis and acid-fast bacilli (arrowheads) residing predominantly within the
necrotic areas. (G and H) Late-stage lesion in the liver with a thin, well-defined macrophage border (arrow) and prominent central necrosis
(dashed lines). Note the high bacterial numbers present in the central, necrotic core in H (arrowhead). (I and J) Multicentric necrotic lesions in
the head kidney during late-stage disease with extensive pigment deposition (arrowheads). Scale bars, 50 ?m. Magnifications, ?200 for A, B, I,
and J and ?400 for all other panels.
FIG. 3. Zebrafish lacking rag1 are hypersusceptible to infection by
M. marinum. Twelve rag1 wild-type, heterozygous, or mutant fish were
infected with 8 CFU of WT bacteria and reared in separate tanks.
Each tank was monitored for mortalities over a 16-week period. A
Kaplan-Meier curve was calculated for each group of fish, and com-
pared to rag1 WT or heterozygous fish, rag1 mutant animals showed a
significant difference in survival as calculated by log-rank test (P ?
0.002). The Cox proportional hazards model was used to quantify the
differences between groups and confirmed significant differences in
survival between mutant and wild-type fish (HR ? 2.7; 95% CI ? 1.6
to 4.7; P ? 0.0001) and mutant and heterozygous fish (HR ? 3.2; 95%
CI ? 2.1 to 4.8; P ? 0.0001). There was no significant difference in
survival between WT and heterozygous fish (HR ? 0.86; 95% CI ?
0.46 to 1.63; P ? 0.654).
VOL. 74, 2006 M. MARINUM INFECTION OF ADULT ZEBRAFISH6113
were from different initial cohorts, they were bred from the
same original AB stocks and were analyzed at the same time
point after infection (16 weeks), at which time all fish also had
similar median bacterial burdens. This analysis showed that the
proportions of nonnecrotizing and solitary granulomas were
significantly higher in ?RD1-infected fish than in fish infected
with wild-type bacteria (Fig. 4B). In addition, the proportion of
loosely aggregated nonnecrotic lesions was also significantly
higher in ?RD1-infected fish (Fig. 4B and D). These observa-
tions suggest that the RD1 locus influences the formation of
tight macrophage aggregates during both embryonic and adult
infection and plays a role in granuloma necrosis during adult
The construction of a flowthrough zebrafish facility has al-
lowed us to conduct detailed longitudinal studies of the course
and pathology of M. marinum infection in large numbers of
adult zebrafish. Our study of M. marinum-induced tuberculosis
in the zebrafish highlights features shared with active human
tuberculosis (53). The hallmark lesion of active tuberculosis,
FIG. 4. ?RD1 is attenuated in adult zebrafish and produces a greater proportion of solitary, loosely associated, nonnecrotic granulomas than
does infection with WT M. marinum. (A) Survival of adult zebrafish infected with WT or ?RD1. Fifteen fish were injected with 50 CFU of WT
or 1,318 CFU of ?RD1 bacteria and were reared in separate tanks. Each tank was monitored for mortalities over a 16-week time period. A
Kaplan-Meier curve was calculated for each group of fish, and compared to WT-infected fish, ?RD1-infected fish showed a significant difference
in survival as calculated by the log-rank test (P ? 0.0012). The Cox proportional hazards model was used to quantify the difference between WT-
and ?RD1-infected fish and confirmed significantly increased survival in the ?RD1-infected fish, with an HR of 0.1592, a 95% CI of 0.04 to 0.57,
and a P value of 0.005. (B) ?RD1 granulomas are more likely to be solitary, nonnecrotizing, and loosely associated than are WT lesions. A total
of 121 WT and 126 ?RD1 granulomas in infected fish at 16 weeks were scored in a blinded fashion for the presence of necrosis and residence in
a multicentric lesion. The subset of nonnecrotic lesions was further assessed for the nature of cellular association (compact or loose). Differences
in the proportion of granulomas falling into each group were assessed using contingency tables to calculate RR and 95% CI, assigning ?RD1 as
the experimental group and the wild type as the control group. Significance was determined using Fisher’s exact test (??, P ? 0.05). Values obtained
for nonnecrotic lesions were an RR of 1.850, a 95% CI of 1.394 to 2.456, and a P value of ?0.0001; values obtained for solitary lesions were an
RR of 1.939, a 95% CI of 1.557 to 2.415, and a P value of ?0.0001; values obtained for loose lesions were an RR of 1.337, a 95% CI of 1.041 to
1.718, and a P value of 0.0111. (C and D) Histopathology of adult zebrafish infected for 16 weeks with either 100 CFU of WT (C) or 1,318 CFU
of ?RD1 (D) bacteria. Median bacterial loads at the time of examination were not significantly different (P ? 0.17 by Mann-Whitney test).
Hematoxylin and eosin staining of pancreatic lesions is shown. Arrows indicate compact, tightly interdigitated epithelioid macrophages in C, while
arrowheads in D show loosely associated macrophage aggregates with minimal epithelioid transformation. Scale bars, 50 ?m. Magnification, ?100.
6114SWAIM ET AL.INFECT. IMMUN.
necrotizing granulomas with abundant bacteria in the necrotic
areas (14, 15, 53), is the predominant lesion at late stages of
zebrafish disease. We took advantage of the first targeted
zebrafish mutant, deficient in rag1, to demonstrate that lympho-
cytes play the same critical role in controlling tuberculosis in
zebrafish and mammals and extended the finding that bacterial
mutants lacking the RD1 locus are attenuated in adult fish by
showing that this attenuation is associated with the formation
of loose macrophage aggregates and a paucity of necrotic le-
Adult zebrafish are exquisitely sensitive to infection with M.
marinum strain M; inoculation of 5 CFU virtually always re-
sults in chronic granulomatous disease. The mortality rate,
early bacterial burdens, and progression of granulomatous pa-
thology are dose dependent, making the zebrafish an ideal
animal model to test the virulence of bacterial mutants. Dif-
ferent M. marinum strains exhibit different virulences in this
model system (54). Human-derived isolates tend to be more
virulent for fish and form a distinct genetic cluster, suggesting
a genetic basis for this increased virulence. Our work lends
further support to this notion, as similar studies using a fish-
adapted strain (ATCC 927) required much higher doses of
bacteria to achieve similar disease states (40).
By using a range of inocula, we found that ?9,000 bacteria
produce acute, fulminant disease. The fish died within 10 days,
with extensive disease and high bacterial burdens. These data
extend previous reports where much higher doses of M. mari-
num M caused rapid death (22, 23). In contrast, we and others
found that lower inocula produce chronic disease (40), similar
to the chronic wasting mycobacterioses of aquarium fish rec-
ognized by tropical fish culturists. The dose required to cause
disease during natural infection is unknown, but it is likely to
come from water contaminated by infected fish or from bio-
films in the environment or system equipment, making it likely
to be low. The 5-CFU dose may simulate a natural disease
course, producing a chronic, long-term infection despite ulti-
mately achieving bacterial burdens and extent of granuloma-
tous disease similar to those of fish infected with higher doses.
A similar relationship between bacterial dose and disease out-
come has previously been described for goldfish and medaka
infected with M. marinum (7, 48). It is likely that high doses
overwhelm the immune system rapidly, whereas the low doses
allow the generation of partially protective innate and adaptive
The presence of necrosis in the zebrafish granulomas distin-
guishes them from M. marinum granulomas in the leopard frog
as well as M. tuberculosis granulomas in the commonly used
mouse model of infection (5, 20, 53). In this regard, they are
similar to M. marinum infection of other fish species (7, 24, 48)
and humans (3) and to M. tuberculosis infection of humans,
nonhuman primates, rabbits, and guinea pigs (2, 8, 15, 16, 29,
53). Therefore, the presence of necrosis in tuberculous gran-
ulomas appears to be at least partially determined by host
factors. The progression of necrosis to thin-walled cavities full
of necrotic material mirrors that found in human tuberculosis
(53). In contrast, leopard frogs are also highly susceptible to M.
marinum infection (?10 CFU establishes infection), but even
doses as high as 7 logs lead to long-term asymptomatic infec-
tion rather than active disease (41). Infected frogs also do not
develop necrotic granulomas even with high inocula. It is pos-
sible that the absence of necrosis indicates an increased resis-
tance to infection, although many other species-specific differ-
ences in immunity likely exist. There is considerable debate
and uncertainty about the role and nature of necrosis, the
physiological and replicative state of the bacteria residing
within necrotic lesions, and their potential contribution to the
antibiotic tolerance that makes tuberculosis so difficult to treat
(14, 25). Little is known about the host and bacterial factors
that contribute to the formation of caseous necrosis, and the
genetic tractability of both the zebrafish and M. marinum
should help to address this question. Indeed, our results show
that the RD1 locus influences the formation of necrosis.
In contrast to human and some nonhuman models, zebrafish
do not appear to clear pathogenic mycobacterial infection,
even at low infecting doses. No significant evidence of healing,
such as fibrosis or calcification, was seen in the many infected
animals that we examined (data not shown). Combined with
previous studies of M. marinum pathogenesis in fish and frogs
(22, 40, 41, 48, 54), our results suggest that the zebrafish is a
good model system for the study of active, caseating tubercu-
losis rather than latent disease, while the frog provides a good
model for indefinitely asymptomatic infection mimicking latent
Unlike mammalian tuberculous granulomas, zebrafish le-
sions have very few lymphocytes. In this way, they are similar to
frog and goldfish granulomas (5, 48) and distinct from M.
marinum-induced human granulomas (3). Based on these ob-
servations, the number of lymphocytes in tuberculous granu-
lomas is likely to be host mediated rather than due to differ-
ences between M. marinum and M. tuberculosis in promoting
lymphocyte infiltration of granulomas. Zebrafish have T lym-
phocytes that populate and exit the thymus in ways similar to
those of mammals (27, 52). Indeed, in silico analysis of the
zebrafish genome suggests that zebrafish share the same T-
lymphocyte subsets as mammals (52). Fluorescence-activated
cell sorter analysis of zebrafish peripheral blood showed that
lymphocytes are present, albeit at reduced ratios compared to
those seen in mice and humans (51).
Given the paucity of lymphocytes in granulomas, it was im-
portant to determine if they played a functional role in the
control of zebrafish tuberculosis. rag1 mutant fish die more
rapidly from M. marinum infection, similar to rag1 mutant mice
infected with M. tuberculosis (44). These rag1 mutant fish do
form granulomas, even necrotic ones. Thus, we conclude that
adaptive immunity does play an important role in the protec-
tion against mycobacterial disease in the fish, suggesting that
the few lymphocytes in zebrafish granulomas are functionally
equivalent to the many lymphocytes found in mammals. These
data corroborate our studies of zebrafish embryos showing that
granuloma formation occurs as a result of bacterial interac-
tions with innate immunity (18). Similar to M. tuberculosis
growth kinetics in mice, rabbits, or guinea pigs (17, 34), M.
marinum grows exponentially over the first few weeks after
infection, followed by a leveling of bacterial numbers. This
plateau in bacterial burden correlates with the onset of the
adaptive immune response in mammalian hosts (17, 34), sug-
gesting that the onset of the adaptive immunity in fish is sim-
ilarly responsible for curtailing bacterial growth. Indeed, the
likely reason for the higher mortality observed in the rag1
mutant fish is a failure to control bacterial growth, despite
VOL. 74, 2006 M. MARINUM INFECTION OF ADULT ZEBRAFISH 6115
grossly normal granuloma formation. More detailed studies
will be required to determine the impact of adaptive immunity
on the kinetics of bacterial growth as well as granuloma for-
mation and the maintenance of granuloma structure.
Finally, we confirm the finding that M. marinum strains lack-
ing the RD1 locus are attenuated in adult zebrafish (22) as they
are in embryos and adult frogs (56). A previous report using
much higher infection doses (103to 105CFU), shorter obser-
vation periods, and different RD1 mutants than the one used in
our study showed lower bacterial counts and longer survival in
zebrafish infected with the mutant bacteria (22). Our extended
analysis reveals that, at least for the mutant used in our studies,
bacteria lacking RD1 ultimately reach a similar plateau as
wild-type bacteria and that this increased bacterial burden is
associated with some host mortality. These results are similar
to those obtained for M. tuberculosis ?RD1 infection of mice,
where high organism burdens are ultimately reached with
some resultant host mortality (45), and have important impli-
cations for the use of strains lacking the RD1 region as vaccine
candidates (45). Low-dose (5- to 50-CFU) infections with the
various RD1 mutant bacteria will help to clarify whether the
differences in doses, observation times, and/or the actual na-
ture of the RD1 mutants account for the different outcomes
observed in the two zebrafish studies.
In previous studies, we have taken advantage of the optical
transparency of zebrafish embryos to determine the steps at
which bacterial virulence determinants impact infection (13,
56). Infection of embryos with the ?RD1 mutant results in
delayed macrophage aggregation, suggesting that this locus
promotes granuloma formation. ?RD1-infected adult fish
form a greater proportion of nonnecrotizing, solitary, and
loose aggregates than do fish infected with wild-type M. mari-
num, suggesting that macrophage aggregation is impaired even
in the context of adult-lineage macrophages and adaptive im-
munity. Based on our embryo work, we had proposed that the
lack of RD1 resulted in delayed kinetics of granuloma forma-
tion (56). In the adult fish, we saw predominantly solitary,
loose aggregates with a few well-formed and even necrotic
granulomas. These findings also suggest a kinetic defect in
granuloma formation rather than an absolute defect that is
associated with an attenuation in bacterial virulence. These
kinetic differences appear to have a significant impact on vir-
ulence, as even much higher inocula of the ?RD1 mutant take
much longer to plateau to wild-type levels and cause limited
death of the infected host. Taken together, these observations
suggest that mycobacterial virulence factors such as RD1 con-
tribute to pathogenesis by altering the kinetics of the host
response throughout the course of infection.
In conclusion, this study extends the findings of others and
lays the groundwork for the use of the adult zebrafish to study
a variety of aspects of tuberculosis ranging from immunology
to pathogenesis and drug tolerance in a facile, small animal
model of active necrotizing disease.
We thank Lance Squires and Josh Brown of Aquatic Habitats for
their assistance with designing the flowthrough system, Artemis Phar-
maceuticals for providing the rag1 zebrafish strain, Allan Decamp for
help with statistical analysis of the survival assays, Jose ´ de la Torre for
assistance in assembling figures and discussions regarding statistical
analysis, Mark Troll and Christine Cosma for advice regarding statis-
tical analysis, Robin Lesley for help with figure design and technical
assistance, and members of the Ramakrishnan laboratory for discus-
L.E.S. dedicates this paper to the memory of Marsha Landolt,
former director of the School of Aquatic and Fisheries Sciences and
Dean of the Graduate School of the University of Washington.
This work was supported by National Institutes of Health grants
RO1 AI036396 and RO1 AI54503 and a Burroughs Wellcome award
to L.R. L.E.C. was supported by a Pfizer Pharmaceuticals postdoctoral
fellowship in infectious diseases, and H.E.V. was supported by an
American Heart Association predoctoral fellowship.
1. Adams, D. O. 1976. The granulomatous inflammatory response. A review.
Am. J. Pathol. 84:164–191.
2. Balasubramanian, V., E. H. Wiegeshaus, and D. W. Smith. 1994. Mycobac-
terial infection in guinea pigs. Immunobiology 191:395–401.
3. Bartralot, R., R. M. Pujol, V. Garcia-Patos, D. Sitjas, N. Martin-Casabona,
P. Coll, A. Alomar, and A. Castells. 2000. Cutaneous infections due to
nontuberculous mycobacteria: histopathological review of 28 cases. Compar-
ative study between lesions observed in immunosuppressed patients and
normal hosts. J. Cutan. Pathol. 27:124–129.
4. Barut, B. A., and L. I. Zon. 2000. Realizing the potential of zebrafish as a
model for human disease. Physiol. Genomics 2:49–51.
5. Bouley, D. M., N. Ghori, K. L. Mercer, S. Falkow, and L. Ramakrishnan.
2001. Dynamic nature of host-pathogen interactions in Mycobacterium
marinum granulomas. Infect. Immun. 69:7820–7831.
6. Brattgjerd, S., and O. Evensen. 1996. A sequential light microscopic and
ultrastructural study on the uptake and handling of Vibrio salmonicida in
phagocytes of the head kidney in experimentally infected Atlantic salmon
(Salmo salar L.). Vet. Pathol. 33:55–65.
7. Broussard, G. W., and D. G. Ennis. 2006. Mycobacterium marinum produces
long-term chronic infections in medaka: a new animal model for studying
human tuberculosis. Comp. Biochem. Physiol., in press.
8. Capuano, S. V., III, D. A. Croix, S. Pawar, A. Zinovik, A. Myers, P. L. Lin,
S. Bissel, C. Fuhrman, E. Klein, and J. L. Flynn. 2003. Experimental My-
cobacterium tuberculosis infection of cynomolgus macaques closely resembles
the various manifestations of human M. tuberculosis infection. Infect. Im-
9. Casanova, J. L., and L. Abel. 2002. Genetic dissection of immunity to my-
cobacteria: the human model. Annu. Rev. Immunol. 20:581–620.
10. Clark, H. F., and C. C. Shepard. 1963. Effect of environmental temperatures
on infection with Mycobacterium marinum (Balnei) of mice and a number of
poikilothermic species. J. Bacteriol. 86:1057–1069.
11. Cosma, C. L., J. M. Davis, L. E. Swaim, H. Volkman, and L. Ramakrishnan.
Zebrafish and frog models of Mycobacterium marinum infection. In Current
Protocols in Microbiology, in press.
12. Cosma, C. L., O. Humbert, and L. Ramakrishnan. 2004. Superinfecting
mycobacteria home to established tuberculous granulomas. Nat. Immunol.
13. Cosma, C. L., K. Klein, R. Kim, D. Beery, and L. Ramakrishnan. 2006.
Mycobacterium marinum Erp is a virulence determinant required for cell wall
integrity and intracellular survival. Infect. Immun. 74:3125–3133.
14. Cosma, C. L., D. R. Sherman, and L. Ramakrishnan. 2003. The secret lives
of the pathogenic mycobacteria. Annu. Rev. Microbiol. 57:641–676.
15. Dannenberg, A. M., Jr. 1993. Immunopathogenesis of pulmonary tubercu-
losis. Hosp. Pract. 28:51–58.
16. Dannenberg, A. M., Jr. 2001. Pathogenesis of pulmonary Mycobacterium
bovis infection: basic principles established by the rabbit model. Tuberculosis
17. Dannenberg, A. M., Jr., and F. M. Collins. 2001. Progressive pulmonary
tuberculosis is not due to increasing numbers of viable bacilli in rabbits, mice
and guinea pigs, but is due to a continuous host response to mycobacterial
products. Tuberculosis (Edinburgh) 81:229–242.
18. Davis, J. M., H. Clay, J. L. Lewis, N. Ghori, P. Herbomel, and L. Ramakrish-
nan. 2002. Real-time visualization of Mycobacterium-macrophage interac-
tions leading to initiation of granuloma formation in zebrafish embryos.
19. Decostere, A., K. Hermans, and F. Haesebrouck. 2004. Piscine mycobacte-
riosis: a literature review covering the agent and the disease it causes in fish
and humans. Vet. Microbiol. 99:159–166.
20. Flynn, J. L. 2006. Lessons from experimental Mycobacterium tuberculosis
infections. Microbes Infect. 8:1179–1188.
21. Flynn, J. L., and J. Chan. 2001. Immunology of tuberculosis. Annu. Rev.
22. Gao, L. Y., S. Guo, B. McLaughlin, H. Morisaki, J. N. Engel, and E. J.
Brown. 2004. A mycobacterial virulence gene cluster extending RD1 is re-
quired for cytolysis, bacterial spreading and ESAT-6 secretion. Mol. Micro-
23. Gao, L. Y., M. Pak, R. Kish, K. Kajihara, and E. J. Brown. 2006. A myco-
6116SWAIM ET AL.INFECT. IMMUN.
bacterial operon essential for virulence in vivo and invasion and intracellular
persistence in macrophages. Infect. Immun. 74:1757–1767.
24. Gauthier, D. T., M. W. Rhodes, W. K. Vogelbein, H. Kator, and C. A.
Ottinger. 2003. Experimental mycobacteriosis in striped bass Morone saxa-
tilis. Dis. Aquat. Organ. 54:105–117.
25. Grosset, J. 2003. Mycobacterium tuberculosis in the extracellular compart-
ment: an underestimated adversary. Antimicrob. Agents Chemother. 47:
26. Guinn, K. M., M. J. Hickey, S. K. Mathur, K. L. Zakel, J. E. Grotzke, D. M.
Lewinsohn, S. Smith, and D. R. Sherman. 2004. Individual RD1-region
genes are required for export of ESAT-6/CFP-10 and for virulence of My-
cobacterium tuberculosis. Mol. Microbiol. 51:359–370.
27. Langenau, D. M., A. A. Ferrando, D. Traver, J. L. Kutok, J. P. Hezel, J. P.
Kanki, L. I. Zon, A. T. Look, and N. S. Trede. 2004. In vivo tracking of T cell
development, ablation, and engraftment in transgenic zebrafish. Proc. Natl.
Acad. Sci. USA 101:7369–7374.
28. Lewis, K. N., R. Liao, K. M. Guinn, M. J. Hickey, S. Smith, M. A. Behr, and
D. R. Sherman. 2003. Deletion of RD1 from Mycobacterium tuberculosis
mimics bacille Calmette-Guerin attenuation. J. Infect. Dis. 187:117–123.
29. McMurray, D. N. 2003. Hematogenous reseeding of the lung in low-dose,
aerosol-infected guinea pigs: unique features of the host-pathogen interface
in secondary tubercles. Tuberculosis (Edinburgh) 83:131–134.
30. Meijer, A. H., F. J. Verbeek, E. Salas-Vidal, M. Corredor-Adamez, J.
Bussman, A. M. van der Sar, G. W. Otto, R. Geisler, and H. P. Spaink. 2005.
Transcriptome profiling of adult zebrafish at the late stage of chronic tuber-
culosis due to Mycobacterium marinum infection. Mol. Immunol. 42:1185–
31. Nasevicius, A., and S. C. Ekker. 2000. Effective targeted gene ‘knockdown’ in
zebrafish. Nat. Genet. 26:216–220.
32. Noga, E. J. 1996. Fish disease: diagnosis and treatment. Mosby-Year Book,
St. Louis, Mo.
33. Noga, E. J., J. F. Wright, and L. Pasarell. 1990. Some unusual features of
mycobacteriosis in the cichlid fish Oreochromis mossambicus. J. Comp.
34. North, R. J., and Y. J. Jung. 2004. Immunity to tuberculosis. Annu. Rev.
35. Opie, E. L., and J. D. Aronson. 1927. Tubercle bacilli in latent tuberculous
lesions and in lung tissue without tuberculous lesions. Arch. Pathol. Lab.
36. Patton, E. E., and L. I. Zon. 2001. The art and design of genetic screens:
zebrafish. Nat. Rev. Genet. 2:956–966.
37. Peterson, R. T., and M. C. Fishman. 2004. Discovery and use of small
molecules for probing biological processes in zebrafish. Methods Cell Biol.
38. Peterson, R. T., S. Y. Shaw, T. A. Peterson, D. J. Milan, T. P. Zhong, S. L.
Schreiber, C. A. MacRae, and M. C. Fishman. 2004. Chemical suppression
of a genetic mutation in a zebrafish model of aortic coarctation. Nat. Bio-
39. Pozos, T. C., and L. Ramakrishnan. 2004. New models for the study of
Mycobacterium-host interactions. Curr. Opin. Immunol. 16:499–505.
40. Prouty, M. G., N. E. Correa, L. P. Barker, P. Jagadeeswaran, and K. E.
Klose. 2003. Zebrafish-Mycobacterium marinum model for mycobacterial
pathogenesis. FEMS Microbiol. Lett. 225:177–182.
41. Ramakrishnan, L., R. H. Valdivia, J. H. McKerrow, and S. Falkow. 1997.
Mycobacterium marinum causes both long-term subclinical infection and
acute disease in the leopard frog (Rana pipiens). Infect. Immun. 65:767–773.
42. Sanger Institute. 12 October 2005, revision date. Mycobacterium marinum
Sequencing Project. [Online.] http://www.sanger.ac.uk/Projects/M_marinum/.
43. Sanger Institute. 20 July 2006, revision date. The Danio rerio Sequencing
Project. [Online.] http://www.sanger.ac.uk/Projects/D_rerio/.
44. Saunders, B. M., H. Briscoe, and W. J. Britton. 2004. T cell-derived tumour
necrosis factor is essential, but not sufficient, for protection against Myco-
bacterium tuberculosis infection. Clin. Exp. Immunol. 137:279–287.
45. Sherman, D. R., K. M. Guinn, M. J. Hickey, S. K. Mathur, K. L. Zakel, and
S. Smith. 2004. Mycobacterium tuberculosis ?RD1 is more virulent than M.
bovis BCG in long-term murine infection. J. Infect. Dis. 190:123–126.
46. Stamm, L. M., and E. J. Brown. 2004. Mycobacterium marinum: the gener-
alization and specialization of a pathogenic mycobacterium. Microbes Infect.
47. Stoskopf, M. K. 1993. Fish medicine. W. B. Saunders Company, Philadel-
48. Talaat, A. M., R. Reimschuessel, S. S. Wasserman, and M. Trucksis. 1998.
Goldfish, Carassius auratus, a novel animal model for the study of Mycobac-
terium marinum pathogenesis. Infect. Immun. 66:2938–2942.
49. Tonjum, T., D. B. Welty, E. Jantzen, and P. L. Small. 1998. Differentiation
of Mycobacterium ulcerans, M. marinum, and M. haemophilum: mapping of
their relationships to M. tuberculosis by fatty acid profile analysis, DNA-
DNA hybridization, and 16S rRNA gene sequence analysis. J. Clin. Micro-
50. Traver, D., P. Herbomel, E. E. Patton, R. D. Murphey, J. A. Yoder, G. W.
Litman, A. Catic, C. T. Amemiya, L. I. Zon, and N. S. Trede. 2003. The
zebrafish as a model organism to study development of the immune system.
Adv. Immunol. 81:253–330.
51. Traver, D., B. H. Paw, K. D. Poss, W. T. Penberthy, S. Lin, and L. I. Zon.
2003. Transplantation and in vivo imaging of multilineage engraftment in
zebrafish bloodless mutants. Nat. Immunol. 4:1238–1246.
52. Trede, N. S., D. M. Langenau, D. Traver, A. T. Look, and L. I. Zon. 2004. The
use of zebrafish to understand immunity. Immunity 20:367–379.
53. Ulrichs, T., and S. H. Kaufmann. 2006. New insights into the function of
granulomas in human tuberculosis. J. Pathol. 208:261–269.
54. van der Sar, A. M., A. M. Abdallah, M. Sparrius, E. Reinders, C. M.
Vandenbroucke-Grauls, and W. Bitter. 2004. Mycobacterium marinum
strains can be divided into two distinct types based on genetic diversity and
virulence. Infect. Immun. 72:6306–6312.
55. van der Sar, A. M., B. J. Appelmelk, C. M. Vandenbroucke-Grauls, and W.
Bitter. 2004. A star with stripes: zebrafish as an infection model. Trends
56. Volkman, H. E., H. Clay, D. Beery, J. C. Chang, D. R. Sherman, and L.
Ramakrishnan. 2004. Tuberculous granuloma formation is enhanced by a
mycobacterium virulence determinant. PLoS Biol. 2:1946–1956.
57. Westerfield, M. 2000. The zebrafish book. A guide for the laboratory use of
zebrafish (Danio rerio). University of Oregon Press, Eugene, Oreg.
58. Wienholds, E., S. Schulte-Merker, B. Walderich, and R. H. Plasterk. 2002.
Target-selected inactivation of the zebrafish rag1 gene. Science 297:99–102.
59. Wolf, J. C., and S. A. Smith. 1999. Comparative severity of experimentally
induced mycobacteriosis in striped bass Morone saxatilis and hybrid tilapia
Oreochromis spp. Dis. Aquat. Organ. 38:191–200.
Editor: J. L. Flynn
VOL. 74, 2006 M. MARINUM INFECTION OF ADULT ZEBRAFISH6117