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Appl Environ Microbiol, June 1998, p. 2256-2261, Vol. 64, No. 6
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Mycobacterium avium Bacilli Grow Saprozoically in
Coculture with Acanthamoeba polyphaga and Survive within
Cyst Walls
Michael
Steinert,1,*
Kristin
Birkness,2
Elizabeth
White,3
Barry
Fields,1 and
Frederick
Quinn2
Respiratory Diseases Branch, Division of
Bacterial and Mycotic Diseases,1
Tuberculosis/Mycobacteriology Branch, Division AIDS, STD and TB
Laboratory Research,2 and
Molecular
Pathology and Ultrastructure Activity, Division of Viral and
Rickettsial Diseases,3 National Center for
Infectious Diseases, Centers for Disease Control and Prevention,
Atlanta, Georgia 30333
Received 1 December 1997/Accepted 17 February 1998
 |
ABSTRACT |
Protozoans are gaining recognition as environmental hosts for a
variety of waterborne pathogens. We compared the growth of Mycobacterium avium, a human pathogen associated with
domestic water supplies, in coculture with the free-living amoeba
Acanthamoeba polyphaga with the growth of M. avium when it was separated from amoebae by a 0.1-µm-pore-size
polycarbonate membrane (in a parachamber). Although viable mycobacteria
were observed within amoebal vacuoles, there was no significant
difference between bacterial growth in coculture and bacterial growth
in the parachamber. This suggests that M. avium is able to
grow saprozoically on products secreted by the amoebae. In contrast,
Legionella pneumophila, a well-studied intracellular
parasite of amoebae, multiplied only in coculture. A comparison of
amoebae infected with L. pneumophila and amoebae infected
with M. avium by electron microscopy demonstrated that there were striking differences in the locations of the bacteria within
amoebal cysts. While L. pneumophila resided within the cysts, M. avium was found within the outer walls of the
double-walled cysts of A. polyphaga. These locations may
provide a reservoir for the bacteria when environmental conditions
become unfavorable.
| |
INTRODUCTION |
The facultative intracellular
pathogen Mycobacterium avium is one of the primary health
threats to patients with AIDS. It can cause bacteremia and disseminated
multiorgan bacterial disease, including pulmonary infections of
immunocompetent individuals (10, 15). The interaction of
mycobacteria with host phagocytic cells likely is central to
mycobacterial pathogenesis. Potential virulence mechanisms against
phagocytic cells include prevention of the acidification of phagocytic
vesicles and limited fusion of the phagosomes with the endosomal and
lysosomal compartments that can lead to bacterial replication within
macrophages (12, 13, 18). Within phagocytic cells M. avium bacilli tend to occupy individual vacuoles and to release
superoxide dismutase (15) and the cell wall constituent
lipoarabinomannan into the cytoplasm and into other lysosomal vesicles
(34). Immunoelectron microscopy of macrophages infected with
M. avium indicates that the parasitophoric vacuolar membrane
possesses the late endosomal marker lysosome-associated membrane
protein 1 but lacks the vesicular proton-ATPase (34).
The epidemiology of M. avium has been compared to that of
Legionella pneumophila (8) because both organisms
are widespread in aquatic environments, including municipal drinking
water systems (9, 14, 16, 27, 32). Previous studies have
proposed that amoebae might be environmental hosts of certain
mycobacteria (4, 24). More recently, it has been shown that
M. avium survives intracellularly within Acanthamoeba
castellanii and interferes with the fusion of the lysosomal and
parasitophoric vacuoles. In addition, the growth of M. avium
in environmental amoebae results in increased virulence in the beige
mouse model (8). These observations are reminiscent of the
intracellular parasitism of L. pneumophila in the
trophozoites of a variety of free-living amoebae. Meanwhile, it is
well-documented that L. pneumophila multiplies within
amoebae and that Acanthamoeba cysts are able to protect
legionellae from certain disinfection procedures (17, 22).
Since the biology of mycobacteria and the biology of legionellae have
some significant similarities, we compared these two pathogens with
regard to their interactions with amoebae. We confirmed that M. avium survives intracellularly and that growth occurs in coculture
with amoebae. In contrast to L. pneumophila, M. avium exhibits saprophytic growth on products secreted by
Acanthamoeba cells. However, since the mycobacteria are
viable in trophozoites and cysts of Acanthamoebae, they might benefit
from this interaction under adverse conditions.
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MATERIALS AND METHODS |
Bacterial strains and growth conditions.
The M. avium serotype 4 strain used was a clinical isolate obtained from
an AIDS patient (19). The strain was cultured on Middlebrook
7H11 agar (Difco) supplemented with oleic acid, albumin, dextrose, and
catalase (BBL) for 10 days at 37°C. For amoeba infection assays,
M. avium was grown in Middlebrook 7H9 broth (Difco)
supplemented with albumin, dextrose, and catalase (BBL) to the late
logarithmic phase with slow shaking (50 rpm) for 8 days at 37°C in
the presence of 5% CO2. L. pneumophila
Philadelphia I JR32 was grown on buffered charcoal-yeast extract agar.
The plates were incubated at 37°C in the presence of 5%
CO2 for 3 days. Escherichia coli HB101 was maintained on Luria-Bertani agar at 37°C.
Acanthamoeba polyphaga.
Acanthamoeba polyphaga
ATCC 30872 was obtained from the American Type Culture Collection and
was maintained axenically at room temperature in PYG 712 broth [2%
proteose peptone, 0.1% yeast extract, 0.1 M glucose, 4 mM
MgSO4, 0.4 M CaCl2, 0.1% sodium citrate dihydrate, 0.05 mM
Fe(NH4)2(SO4)2 · 6H2O, 2.5 mM NaH2PO3, 2.5 mM
K2HPO3] as monolayers in 75-cm2
tissue culture flasks. Amoebae were suspended by tapping the flasks.
Cell counts were determined with a modified Fuchs-Rosenthal chamber.
Amoebae were subcultured at intervals of 10 days.
Infection of A. polyphaga monolayers and parachamber
experiments.
L. pneumophila and E. coli were
suspended in Acanthamoeba buffer (see below) and centrifuged
at the maximum speed in an Eppendorf microcentrifuge, and the pellets
were washed twice in Acanthamoeba buffer. Broth cultures of
M. avium were likewise centrifuged and washed twice in
buffer. After washing, M. avium and L. pneumophila were incubated in Acanthamoeba buffer for
10 days (33°C, 5% CO2) without shaking to deplete the
stored nutrients in the bacteria. This 10-day starvation procedure was
necessary since the residual growth rate of M. avium in
buffer was very high. After this step the bacterial suspensions were
vortexed for 1 min and adjusted in Acanthamoeba buffer to a
concentration of 108 cells/ml, as determined by optical
density at 600 nm.
Axenic cultures of A. polyphaga in the logarithmic phase
were centrifuged (200 × g, 15 min), and the pellets
were resuspended in Acanthamoeba buffer (PYG 712 medium
without proteose peptone, yeast extract, sodium citrate, and glucose).
After two repetitions of this washing step, the cells were adjusted to
a titer of 105 cells per ml in this buffer. One milliliter
of this amoebal suspension was pipetted into each well of a 24-well
microtiter plate (Costar, Cambridge, Mass.). Following 2 h of
incubation at 33°C, the amoeba monolayer was inoculated with 10 µl
of a bacterial suspension (105 bacteria/ml), which resulted
in a preparation containing 103 bacteria/ml (multiplicity
of infection, 10
2). The numbers of CFU were determined at
time zero and after 1, 7, and 14 days. In a control experiment the
multiplication of the bacteria in buffer without amoebae was
determined. To minimize evaporation, the plates were sealed in plastic
bags.
In the parachamber experiment the bacteria were washed as described
above, and 1 ml of a suspension containing 103 bacteria/ml
in Acanthamoeba buffer was pipetted into each well of a
24-well microtiter plate (Costar). After insertion of a parachamber (diameter, 6.5 mm; transwell pore size, 0.1 µm; Costar) into each well, 200 µl of an amoebal suspension containing 5 × 105 cells/ml in Acanthamoeba buffer was pipetted
into each transwell. Colony counts were determined at time zero and
after 1, 7, and 14 days. As described above for the
Acanthamoeba infection, control experiments in buffer were
performed and precautions to prevent evaporation were taken. In
addition, the separation of bacteria and amoebae by the
0.1-µm-pore-size polycarbonate membrane filter was confirmed by
viable cell counting and by light microscopy of the contents of the
lower chamber.
Localization of intracellular bacteria and viability testing of
M. avium.
One milliliter of an Acanthamoeba cell
suspension (105 cells/ml in Acanthamoeba buffer)
was inoculated with a minimal volume containing 107
bacteria (in Acanthamoeba buffer) to yield a multiplicity of infection of 100. At 1, 6, 24, and 48 h the amoeba monolayer was washed twice with 1 ml of phosphate-buffered saline to eliminate noningested bacteria. To localize intracellular bacteria, the cells
were processed for acid-fast staining and electron microscopy. For
electron microscopy the cells were fixed in 2.5% cacodylate-buffered glutaraldehyde, postfixed with 1% osmium tetroxide in 0.1 M cacodylate buffer (1 h), and embedded in Epon resin. Thin sections were stained with 2% uranyl acetate and lead citrate and examined with a
transmission electron microscope (Phillips Electronic Instruments,
Mahwah, N.J.) at 40 kV.
To determine the viability of intracellular bacteria, a Baclight
Live/Dead kit (Molecular Probes, Junction City, Oreg.) was
used as
described by the manufacturer. Briefly, after 2 days of
incubation at
33°C, infected acanthamoebae (see above) were mounted
on glass
slides. Saline (100 µl) was placed on the air-dried cells,
and 1 µl
of SYTO 9 nucleic acid stain and 1 µl of a propidium
iodide solution
were suspended in the saline. SYTO 9 is a nucleic
acid stain that
labels bacterial cells with green fluorescence;
propidium iodide, a red
fluorescent nucleic acid stain, penetrates
only bacteria with damaged
membranes and effectively competes
with SYTO 9 for nucleic acid binding
sites. Thus, damaged cells
were identified by red fluorescence, while
live cells were identified
by green fluorescence. The preparation was
incubated in the dark
for 15 min and then examined by fluorescence
microscopy.
Synchronous encystment of infected A. polyphaga
cells.
The experiment to study infection of A. polyphaga monolayers was set up as described above for the
coculture experiment except that the mycobacterial inoculum used was
larger (final concentration, 107 bacteria/ml). After
incubation for 24 h at 33°C, the supernatant was discarded, and
the amoeba monolayer was rinsed with encystment buffer [0.1 M KCl,
0.02 M tris(2-amino-2-hydroxymethyl)-1,3-propandiol, 8 mM
MgSO4, 0.4 mM CaCl2, 1 mM NaHCO3]
and then incubated in fresh encystment buffer at 33°C. After 3 days,
the cell suspension was centrifuged (1,000 × g, 20 min), and the pellet was resuspended in 3% (vol/vol) hydrochloric
acid. This acid treatment was sufficient to kill the remaining
trophozoites, immature cysts, and extracellular bacteria after 36 h. During the treatment the percentages of viable amoebae and bacteria
were determined by Trypan blue exclusion and plating on Middlebrook
7H11 agar, respectively. After the acid treatment the cysts were washed
three times with Acanthamoeba buffer. One-half of the sample
was processed for electron microscopy (see above), and the other half
was incubated in PYG medium at 33°C for 7 days. The excystment of the
cysts was examined by light microscopy, and the presence of viable
bacteria was determined by viable counting on Middlebrook 7H11 agar
plates.
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RESULTS |
Growth of bacteria in Acanthamoeba buffer.
Despite
intensive washing with buffer, unstarved mycobacteria showed
significant residual growth in buffer after 10 days of incubation
(Table 1). The increases in CFU per
milliliter were 72-fold for the unstarved bacilli and 6-fold for the
starved mycobacteria. In contrast, the number of CFU of L. pneumophila was almost unaffected by starvation (0.3-fold decrease
for unstarved legionellae and 0.7-fold decrease for starved
legionellae). The viability of unstarved E. coli decreased
0.049-fold. Although media and growth conditions are known to influence
virulence, there was no obvious difference in the initial uptake by
amoebae between starved and unstarved bacteria (data not shown).
Direct coculture and parachamber culture.
E. coli and
starved cells of M. avium and L. pneumophila were
tested for the ability to grow in coculture with A. polyphaga. The results of this growth assay are shown in Fig.
1. For M. avium the number of
bacteria increased from 1.5 × 103 CFU in the inoculum
to 3 × 105 CFU after 7 days and to 4 × 106 CFU after 14 days. In monolayers infected with L. pneumophila the number of bacteria increased from 5 × 102 CFU in the inoculum to 3 × 105 CFU
after 2 days to 2 × 107 CFU after 14 days. E. coli also multiplied in coculture; the number of bacteria
increased from 866 CFU to 1.6 × 106 CFU within 1 day.
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FIG. 1.
Growth of the M. avium serotype 4 strain,
L. pneumophila JR32, and E. coli HB101 in
cocultures with A. polyphaga. Each coculture experiment was
performed three times in Acanthamoeba buffer at 33°C. The
values shown are the mean numbers of CFU (± standard deviations)
determined at zero time and at 1, 7, and 14 days after coincubation was
started.
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Parachamber experiments were used to determine if the amoeba-associated
growth of the bacteria was due to intracellular parasitism
of
A. polyphaga (Fig.
2).
M. avium and
E. coli showed approximately
the same growth
kinetics as those observed in the direct coculture
experiment. In
contrast,
L. pneumophila showed no increase in
CFU when it
was separated from the amoebae by the 0.1-µm-pore-size
polycarbonate
membrane. Due to the low-nutrient-content environment
the number of
amoebal cysts increased during the course of the
experiment.
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FIG. 2.
Growth of the M. avium serotype 4 strain,
L. pneumophila JR32, and E. coli HB101 when they
were separated from A. polyphaga by a 0.1-µm-pore-size
polycarbonate membrane (parachamber). Each parachamber experiment was
performed three times in Acanthamoeba buffer at 33°C. The
values shown are the mean numbers of CFU (± standard deviations)
determined at zero time and at 1, 7, and 14 days after inoculation.
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Intracellular survival.
Acid-fast staining, Gimenez staining,
and transmission electron microscopy were used to demonstrate bacterial
ingestion by A. polyphaga during coculture. These studies
confirmed previous observations that great numbers of legionellae are
found intracellularly inside a single vacuole. The numbers of
legionellae increase over time, and finally the bacteria fill the
entire host cell. In contrast, the numbers of intracellular M. avium cells were much lower (1 to 20 bacteria per vacuole), and
the bacteria were found in several vacuoles (one to six vacuoles per
cell) (Fig. 3). The viabilities of
intracellular bacteria could be confirmed by using differential live-dead fluorescence staining (Fig. 4).
The percentage of infected amoebae increased over time, while the
number of intracellular mycobacteria within a single host cell remained
constant after 2 days of cocultivation. This observation suggests that
the primary mechanism of mycobacterial growth in coculture is not
intracellular. The very few amoebae that contained E. coli
cells contained one to three bacteria in a single vacuole.
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FIG. 3.
Transmission electron micrograph of M. avium
serotype 4 bacilli within cytoplasmic vesicles of an A. polyphaga trophozoite after 2 days of coincubation. Bar = 1 µm.
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FIG. 4.
Differential live-dead fluorescence staining of M. avium serotype 4 bacilli within vesicles of A. polyphaga trophozoites after 2 days of coincubation. Green
fluorescence indicates live bacterial cells that have an intact
membrane.
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M. avium within Acanthamoeba cysts.
The ability of L. pneumophila to survive within the cysts of
A. polyphaga was previously demonstrated by Kilvington and
Price (22). To determine whether M. avium and
E. coli have similar capabilities, the encystment of amoebae
after coculture with these species was induced by incubation in
encystment buffer. Six hours after cyst induction the trophozoites
rounded up, and encystment of a trophozoite appeared to be complete
after 18 h. After 3 days about 92% of the trophozoites produced
mature cysts (as determined by the presence of a double wall). The
upper temperature limit that permits encystment is 40°C, but optimal
numbers of infected cysts were found at 33°C. In order to kill the
remaining trophozoites, immature cysts, and extracellular bacteria, a
treatment with hydrochloric acid (3%, vol/vol) was performed.
Examination of thin sections by transmission electron microscopy
revealed double-walled cysts containing one to nine mycobacteria within
the amoeba outer cell wall (Fig. 5).
E. coli was not found within cysts. Our observations that
L. pneumophila was within the cysts but was not between the cell walls of the cysts were consistent with the description given in
the previous study. After 7 days in PYG medium the
Acanthamoeba cysts were able to excyst and replicate.
Legionellae and mycobacteria (at lower numbers), but not E. coli, were culture positive on growth plates after the excystment
of the amoebae.
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FIG. 5.
Transmission electron micrograph of a mature A. polyphaga cyst containing M. avium serotype 4 bacilli
(arrow) within the double cell wall (note that the outer cell wall is
divided and surrounds the bacterial cells). Bar = 1 µm.
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DISCUSSION |
The importance of protozoans in soil and water ecosystems has been
recognized for several decades, and the relevance of predatory protozoans in the control of bacterial populations is widely
acknowledged (11, 20, 30, 31). However, the potential role
of protozoans as reservoirs for human pathogens has only recently
received adequate attention (2, 4, 17, 25). The best-studied
example of a protozoan-bacterial pathogen interaction is the
intracellular parasitism of L. pneumophila. Adaptation of
L. pneumophila to parasitism of free-living amoebae might
have led to the ability of this environmental bacterium to infect human
lung macrophages (4). This observation led to the concepts
that selection for resistance to digestion by predatory protozoans is a
driving force in the evolution of pathogenic environmental bacteria and
that protists are the "missing link between ecology and pathology" (4, 23, 25).
Since M. avium has been isolated from habitats where amoebae
normally feed on bacteria (14, 26, 29, 32), we studied the
interaction of A. polyphaga with M. avium. Our
results show that M. avium does not behave in a manner
analogous to L. pneumophila. Since L. pneumophila
only grows intracellularly within amoebae, the parasitism of
legionellae seems to be much more specific and more highly evolved than
the simple growth enhancement of mycobacteria and E. coli by
their association with amoebae. In pure buffer (parachamber
experiments) M. avium and E. coli were able to
grow as free-living saprophytes on products secreted by A. polyphaga. Although we demonstrated that the growth of M. avium in direct cocultures and the growth of M. avium
in parachamber experiments are similar, we cannot eliminate the
possibility that intracellular multiplication occurs. Consistent with
results published recently (8), we found that M. avium is able to survive in the intracellular environment of
amoebae, and consequently, these bacteria may receive nutrients and
grow within the protozoans. Considering that M. avium is a
slowly growing bacterium that lives in a habitat where amoebae feed on
bacteria, survival in a hostile intracellular environment may be an
important advantage.
Although there are important differences in the interactions of amoebae
with legionellae and mycobacteria, intracellular survival provides a
possible evolutionary explanation for how a saprozoic-saprophytic or
parasitic environmental organism can acquire the ability to survive
within human macrophages. M. avium and L. pneumophila are known to inhibit phagolysosomal vacuole fusion in
amoebae and macrophages (5, 8, 13, 18, 21). Other defense mechanisms, such as bacterial toxin production, the presence of protective outer membrane structures, and the reduction of vacuole acidification by the bacteria, might also contribute to resistance in
phagocytic cells. A comparison of the intracellular activities that
occur in protozoans after ingestion of bacteria showed that these
activities are very similar to the activities observed in macrophages
(1, 3, 5, 7, 8, 25, 33).
It is generally accepted that engulfed bacteria may benefit from the
protective coat conferred by protozoans (23). The presence of mycobacteria in domestic water supplies and the results of disinfection studies in which the authors tested various concentration of sodium hypochlorite suggest that chlorination has little effect on
mycobacteria (6, 9, 27). Previous studies have demonstrated that cysts of A. polyphaga can contain viable L. pneumophila cells which are protected from disinfection
(22). Recovery of M. avium from HCl-treated cysts
showed that the exploitation of this amoebal differentiation event is
not restricted to L. pneumophila. Therefore, we suggest that
the resistance of infected amoebal cysts to biocidal agents may
additionally interfere with disinfection of domestic water supplies
contaminated with mycobacteria. However, in contrast to L. pneumophila, M. avium is located within the double
walls of the cysts, which are known to be composed largely of
polysaccharides (one-third is cellulose) (28). The numbers
of bacterial cells in the cysts are much lower than the numbers of
legionellae, and it is unclear how mycobacteria are distributed to this
location.
To control and prevent the dissemination of nontuberculosis
mycobacterial infections, it is important to focus on the ecology of
the bacterial environment. Although no data are available, protozoans
like Hartmannella and Naegleria spp. very likely
exhibit interactions with mycobacteria identical to those observed with acanthamoebae. The results of these and other studies in which pathogens were enhanced by associations with protozoans and perhaps biofilms indicate that there is a need for further research on the
physiological ecology of these pathogens (16).
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ACKNOWLEDGMENT |
This work was supported by grant Ste 838/1-1 from the Deutsche
Forschungsgemeinschaft.
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FOOTNOTES |
*
Corresponding author. Mailing address: Respiratory
Diseases Branch, Division of Bacterial and Mycotic Diseases, Centers
for Disease Control and Prevention, 1600 Clifton Road, Atlanta, GA 30333. Phone: (404) 639-0855. Fax: (404) 639-4215. E-mail:
zma7{at}cdc.gov.
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Abstract
Introduction
