Washington State University, Department of
Microbiology, Pullman, Washington 99164-4233
Received 27 January 2000/Returned for modification 14 February
2000/Accepted 1 April 2000
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INTRODUCTION |
Coxiella burnetii, the
causative agent of Q fever, is an obligate intracellular organism that
replicates within the phagolysosome of host cells. The phagolysosome is
a harsh environment where C. burnetii is exposed to
degradative proteases, reactive oxygen species, and a pH below 4.8 (13, 19). Despite the inability of C. burnetii to
replicate under any known in vitro conditions, some metabolic processes
can be supported in vitro where the pH appears to be a critical factor.
Thus, C. burnetii can transport and incorporate glucose,
glutamate (11), and proline (14) and synthesize
nucleic acids (9) and proteins (28) at a mildly acidic, but not at a neutral, pH.
By 48 h postinfection, vacuoles containing C. burnetii
(VCB) appear to be typical phagolysosomes. Antibodies to lysosomal membrane proteins label the membranes of VCB (13). The
lysosomal enzyme, acid phosphatase, as well as fluid phase markers used to label lysosomes, are found inside VCB (1, 8). There is evidence to suggest, however, that C. burnetii is able to
modify its environment. The respiratory burst that accompanies
phagolysosomal fusion is significantly reduced during C. burnetii infection of J774A.1 cells (6).
C. burnetii has two cell variants, the large-cell variant
(LCV) and the small-cell variant (SCV), both of which are infectious (30). The morphological differences between these two
variants have been carefully described (5, 8, 20, 22). The
LCV has greater metabolic activity and is more sensitive to
environmental stresses than the SCV, while the environmentally stable
SCV has a thicker peptidoglycan layer, has more condensed nuclear
material and, as the name implies, is smaller in size. On the basis of these studies and the evidence that C. burnetii in the
extracellular environment can remain infectious for more than a year
(31), it has been suggested that the infective variant in
natural aerosols is primarily the SCV and that infection is initiated
when phagocytic cells internalize C. burnetii contained in
inhaled aerosols. In this model, exposure to the phagolysosomal
environment activates C. burnetii metabolism and replication
ensues. Based on electron microscopic studies, it has been proposed
that intracellular C. burnetii goes through a typical
bacterial growth cycle, with an increase in the relative number of LCVs
as the population enters log phase (20). Then, as the
stationary phase is approached, there is an increase in the number of
SCVs, and LCVs occasionally divide asymmetrically producing a
spore-like form (21). The bacteria are released from the
host cell as a result of host cell lysis or possibly exocytosis, and
these "naturally released" C. burnetii infect other host cells.
The present study considers this model by exploiting the recent
discovery of an SCV-specific C. burnetii protein, ScvA. It has been shown that when C. burnetii variants are separated
on a density gradient that antibody to ScvA binds only to the more dense SCV (12). Using naturally released C. burnetii to infect host cells, we found that the transition from
SCV to LCV takes place immediately following uptake and that in vitro
this transition takes place most rapidly at a pH higher than that
expected in the phagolysosome. In addition, C. burnetii is
able to delay phagolysosomal fusion, perhaps to facilitate this
transition from SCV to LCV.
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MATERIALS AND METHODS |
Organisms and cell lines.
J774A.1 cells, a continuous murine
macrophage-like cell line (TIB-67; American Type Culture Collection,
Rockville, Md.), were used as host cells in all experiments. Cells were
maintained in RPMI 1640 medium (Gibco BRL, Grand Island, N.Y.)
supplemented with 10% heat-inactivated fetal bovine serum (FBS) and
100 U of penicillin and 100 U of streptomycin per ml at 37°C and 5%
CO2. Nine Mile Phase I C. burnetii
(27) was propagated in persistently infected J774A.1 cells
maintained as described above without antibiotics. C. burnetii cells were harvested from the supernatants of cultures by
differential centrifugation, first at 550 × g for 10 min to remove cell debris and then at 15,000 × g for
1 h to pellet bacteria. The bacterial numbers were determined with
a Klett-Summerson photoelectric colorimeter (Klett Manufacturing Co.,
Inc., New York, N.Y.) using a no. 42 filter (23).
Quantitation with a photoelectric colorimeter is an approximation due
to variable amounts of cell debris in the final pellet. C. burnetii were inactivated by a 12-h incubation with vigorous
shaking in 2% paraformaldehyde-2% glutaraldehyde in
phosphate-buffered saline (PBS; 0.1 M NaPO4, 0.1 M NaCl
[pH 7.4]). Salmonella enterica serovar Enteritidis, used
as a biodegradable particle, was grown in Luria-Bertani broth and
inactivated by incubation with gentamicin at 250 µg/ml, in addition
to 25 µg of gentamicin per ml added to the tissue culture media to
ensure inactivation.
Electron microscopy.
A total of 5 × 105
J774A.1 cells were seeded onto 15-mm Thermonox coverslips (Fisher
Scientific, Santa Barbara, Calif.) in the wells of a 24-well Falcon
plate (VWR Scientific Products, Seattle, Wash.) and allowed to adhere
for 12 h. Media were removed, and J774A.1 cells were incubated
with C. burnetii at an approximate multiplicity of infection
(MOI) of 1,000:1 in RPMI 1640-10% FBS. After 15 min C. burnetii was removed by washing three times with RPMI 1640 prewarmed to 37°C. Cells were fixed at 0, 15, 30, 60, and 120 min in
Immunofix {0.5% gluteraldehyde, 2% paraformaldehyde, 0.1 M PIPES
[piperazine-N,N'-bis(2-ethanesulfonic acid)]}
for 1 h at 25°C, after which fixative was replaced with 0.1 M
PIPES buffer. Thermonox coverslips were transferred to glass vials for further processing. Samples were dyhydrated through an ethanol series
and infiltrated with L. R. White resin. To embed the monolayer, Thermonox coverslips were inverted on 7- by 7-mm Histoprep disposable base molds (Fischer, Pittsburgh, Pa.) and cured overnight at 60°C. The Thermonox coverslips were removed from the hardened resin by
heating, and J774A.1 cells were preserved as a monolayer on the surface
of the resin block. Thin sections were taken from the block face and
mounted on Formvar-coated nickel grids for immunogold labeling.
Immunogold labeling of ScvA was conducted by blocking samples for
1 h in TBST (10 mM Tris, 250 mM NaCl, 0.3% Tween 20 [pH 8.2])-1% bovine serum albumin (BSA), followed by a 1 h of
incubation in rabbit anti-ScvA polyclonal sera (kindly provided by R. Heinzen) diluted 1:25 in TBST-BSA. Samples were then washed three times for 15 min in TBST-BSA and incubated for 1 h with goat anti-rabbit antibody which was conjugated to 10- or 20-nm gold particles. Samples
were again washed twice with TBST-BSA, twice with TBST, and once with
double-distilled H2O. Sections were stained with uranyl
acetate-KMnO4 for 10 min and examined with a JEOL 1200 EX
transmission electron microscope.
To label phagolysosomes with thorium dioxide, J774A.1 cells were seeded
at 5 × 105 cells on 15-mm Thermonox coverslips as
described above. Cells were incubated for 6 h in thorium dioxide
(kindly provided by O. Baca) diluted in RPMI 1640-10% fetal calf
serum to label the lysosomes. This was followed by a 12-h chase period
to allow internalization of all thorium dioxide. These cells were
incubated with Nine Mile Phase I C. burnetii or inactivated
C. burnetii at an MOI of approximately 1,000:1 for 1 h
or inactivated serovar Enteritidis at 100:1 for 30 min. Inocula were
removed by washing three times with prewarmed RPMI 1640, and the media
were replaced.
At specified time points, inoculated J774A.1 cells were fixed for
electron microscopy in 4% paraformaldehyde-2% glutaraldehyde-0.1 M
sodium cacodylate at 4°C for 12 h. Fixative was washed off with 0.1 M cacodylate buffer (pH 7.5) and Thermonox coverslips were processed as described above. Samples were embedded in Epon, inverted on Histoprep disposable base molds, and cured overnight at 70°C. Thin
sections were mounted on Formvar-coated copper grids, stained with 2%
uranyl acetate and lead citrate, and examined with a Hitachi 600 transmission electron microscope.
To label phagolysosomes with acid phosphatase stain, J774A.1 cells were
seeded on Thermonox coverslips as described above. J774A.1 cells were
inoculated with viable or inactivated C. burnetii at an MOI
of approximately 1,000:1 or 0.8-µm-diameter latex beads (Sigma, St.
Louis, Mo.). Inocula were removed by washing three times with warmed
RPMI. At each time point samples were fixed in 1.5%
glutaraldehyde-0.1 M sodium cacodylate buffer [pH 7.4] for 15 min.
Fixative was removed, and samples were washed three times for 2 min
each time in Tris-malate buffer (0.2 M Tris, 0.2 M maleic acid, with
0.8 g of NaOH/100 ml added [pH was adjusted to 5.0 with 10 N
NaOH]). Samples were incubated in the reaction mix (one part 1.25%
-glycerophosphate [pH 5.0], one part H2O, one part
Tris-malate buffer, and two parts 0.2% lead nitrate [added drop by
drop to avoid precipitation] mixed immediately before use) for 1 h with shaking at 37°C. Samples were then washed three times for 2 min in Tris-malate buffer and fixed for 12 h at 4°C in 3%
glutaraldehyde-1% sucrose-0.2 M sodium cacodylate buffer (pH 7.4)
(7). Samples were prepared and embedded for electron microscopy as above.
Activation and metabolic labeling.
Activation of C. burnetii was achieved by incubating 109 C. burnetii in 0.5 ml in acid activation buffer (32 mM
KPO4, 15 mM NaCl, 152.2 mM KCl, 100 mM glycine, 5 mM
L-glutamate, 20 µM L-proline), adjusted to pH
7.0, 5.5, or 4.5 at 37°C for 2 h. To verify that the pH did not
change during the incubations, the pH was monitored immediately before
and after all incubations. Pelleted C. burnetii were
embedded in 10 µl of 4% low-melting-point agarose and processed for
immunolocalization and electron microscopy as described above.
Approximately 5 × 109 C. burnetii were
incubated in 250 µl of acid activation buffer at pH 4.5, 5.5, or 7.0 with 50 µCi of [35S]methionine-[35S]cysteine (Express;
NEN, Boston, Mass.) and 50 µg of cyclohexamide per ml to prevent
incorporation by residual host cells. For each experiment the harvested
C. burnetii cells were divided equally into three samples
(approximately 109 C. burnetii cells/sample).
After a 16-h incubation, C. burnetii cells were washed three
times in PBS to remove the extracellular [35S]methionine-[35S]cysteine and denatured
by boiling in Laemmli sample buffer. The counts per minute (cpm) in the
denatured samples, from three independent experiments, were determined
by scintillation counting. To visualize incorporation, proteins were
resolved by sodium dodecyl sulfate-polyacrylamide electrophoresis
(SDS-PAGE) (17) using a minigel apparatus (Bio-Rad,
Hercules, Calif.). Gels were loaded with equal numbers of bacteria per
lane which was confirmed by staining with Coomassie brilliant blue R250
(Bio-Rad). Autoradiographs of dried gels were analyzed with an Image
Master scanning densitometer and software (Pharmacia Biotech, Uppsala, Sweden).
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RESULTS |
Kinetics of ScvA degradation following internalization.
J774A.1 cells infected with a mixture of SCV and LCV C. burnetii were examined at 2 and 6 h postinfection by
transmission electron microscopy (TEM). The morphology of the C. burnetii in the inoculum, as well as samples viewed at 2 h
postinfection, indicate that these organisms are primarily SCVs. By
6 h postinfection, internalized C. burnetii cells
appear to be primarily LCVs (Fig. 1).
Infected host cells and the inoculum were prepared for immunoelectron microscopy, and the sections were incubated with anti-ScvA polyclonal sera followed by colloidal gold labeling. To quantitate the apparent decrease in SCVs following internalization, we monitored the decrease in ScvA, a protein specific to the SCVs of C. burnetii,
using anti-ScvA polyclonal sera and colloidal gold. There was a
reduction in the ScvA/SCV ratio from an average of 4.1 to 2.2 ScvA/SCV
during the 2 h immediately after internalization. In addition, we
found that the percentage of LCV C. burnetii, as defined by
the complete lack of gold label, increased from 25% in the inoculum to
83% during this time period (Fig. 2).
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FIG. 1.
(A) TEM image showing a typical vacuole in J774A.1 cells
infected with C. burnetii at 2 h postinfection. (B)
J774A.1 cells infected with C. burnetii, in the same
experiment as panel A, at 6 h postinfection. At least 20 vacuoles
were examined at each time point. Bar, 100 nm.
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FIG. 2.
In vivo experiments show the decrease in ScvA label
after infection with viable C. burnetii. ScvA, a
small-cell-specific protein, was labeled with anti-ScvA polyclonal
sera. The percentage of LCV ( , as defined by the absence of any
anti-ScvA polyclonal serum label) increases over the 2 h, while at
the same time the average number of ScvA/SCV ( ) decreases. The data
were obtained from a minimum of 50 C. burnetii organisms for
the inocula and at each time point shown.
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Effect of pH on ScvA.
When C. burnetii is
internalized the endosome is acidified by vacuole ATPases to pH 5.5 (32) and, after phagolysosomal fusion, the pH drops still
further to pH 4.5 (19, 24). The pH is one element of the
vacuolar environment that could initiate the decrease in ScvA;
therefore, we monitored the effect of pH on ScvA in vitro. Identical
samples of C. burnetii were incubated at 37°C for 2 h
in an acid activation buffer with the pH adjusted to 7.0, 5.5, or 4.5. The samples, as well as an untreated control, were then fixed for
immunoelectron microscopy, and ScvA was labeled with colloidal gold.
The average number of gold particles (ScvA label)/C. burnetii cell was determined for each treatment in four
independent experiments. The number of gold particles/C.
burnetii incubated at a pH of 7.0 or 4.5 was found to be not
statistically different from untreated controls. The decrease in ScvA
label seen with incubation at pH 5.5 was statistically significant in
all experiments (P = 0.001), with an average decrease
of 42% after 2 h (Fig. 3). This
decrease is less than the decrease in ScvA label seen in vivo during
the same time period (an average decrease in ScvA of 71%). This
suggests that either some of the SCV are degraded in vivo or that the
decrease in ScvA is not only a response to pH.
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FIG. 3.
C. burnetii were incubated for 2 h in
acid activation buffers at pH 7.0, 5.5, or 4.5. Samples were labeled
with anti-ScvA polyclonal serum and examined by electron microscopy.
The ScvA/C. burnetii ratio was compared to that found in an
untreated control. At least 200 C. burnetii organisms were
examined for each treatment in four independent experiments, with error
bars indicating the standard deviation. Only treatment at pH 5.5 yielded a ScvA/C. burnetii ratio that was significantly
different as compared by Student t tests, with a reduction
of 42% and a 95% confidence interval (P  0.001).
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Effect of pH on internalization of
[35S]methionine-[35S]cysteine.
To
further explore the effect of pH, C. burnetii was incubated
for 16 h in acid activation buffers containing cyclohexamide and
[35S]methionine-[35S]cysteine under the
same conditions as described above. The 35S internalized by
C. burnetii was determined for three experiments (Fig.
4). There was significantly more
35S in organisms incubated at pH 5.5 than at pH 4.5 or 7.0. However, at pH 4.5, the estimated pH of the phagolysosome, there was
2.5 times as much 35S as in organisms incubated at pH 7.0. Proteins were resolved by electrophoresis, with an equal number of
organisms being loaded onto each lane (Fig.
5). Proteins labeled with
[35S]methionine-[35S]cysteine during acid
activation were visualized by autoradiography. Although there were no
obvious differences in protein expression, incubation at pH 5.5 resulted in an increase in protein synthesis by C. burnetii.
This increased synthesis and the observed decrease in ScvA in response
to pH may be necessary for the establishment of infection.
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FIG. 4.
Approximately 5 × 109 C. burnetii were incubated for 16 h in acid activation buffers
at a pH of 7.0, 5.5, or 4.5 with
[35S]methionine-[35S]cysteine and
cycloheximide. Internalization of labeled amino acids was determined by
scintillation counting. Bars indicate the average cpm internalized
during incubation for three experiments, with error bars indicating the
standard deviation. Internalized cpm for each treatment were
significantly different from the other two by Student t
tests, with a confidence interval of 95% (P 0.05).
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FIG. 5.
Approximately 109 C. burnetii
were incubated in acid activation buffers with pH set to 7.0, 5.5, or
4.5 for 16 h with
[35S]methionine-[35S]cysteine and
cycloheximide. C. burnetii organisms were denatured in
Laemmli buffer, and proteins were separated by SDS-PAGE. (A) Coomassie
blue-stained image with equal numbers of C. burnetii per
lane. (B) Autoradiograph of the same gel. Samples incubated at pH 4.5 (lane 1), 5.5 (lane 2), 7.0 (lane 3) are shown. (C) Comparison of
proteins synthesized during incubations at pH 4.5 and 5.5. Autoradiographs were exposed for different lengths of time. Lane 4, containing sample incubated at pH 4.5, was exposed for 72 h, and
lane 5, containing sample incubated at pH 5.5, was exposed for 24 h.
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Phagolysosomal fusion.
Most intracellular pathogens are able
to modify their intracellular environments. The possibility that
C. burnetii is also able to affect phagolysosomal fusion was
investigated here. Two methods, thorium dioxide labeling and acid
phosphatase staining, were used to visualize phagolysosomal fusion, and
both techniques yielded very similar results. Viable and
formaldehyde-inactivated C. burnetii were used in both of
these experiments. There are limitations to both of these methods.
During thorium dioxide labeling, an extended chase period after
labeling is necessary for complete internalization of label in order to
avoid false-positive results found when extracellular label is taken up
with the inoculum. During the chase period, new lysosomes may form that
do not contain the marker. Acid phosphatase staining is occasionally
variable, resulting in little or no staining of some cells
(7). Using thorium dioxide or acid phosphatase, we obtained
maxima of 70 and 90% labeled VCB, respectively.
Lysosomes of J774A.1 cells were labeled with thorium dioxide, a
fluid-phase marker, and then labeled cells were inoculated with viable
or inactivated C. burnetii or with inactivated
Salmonella serovar Enteritidis. The thorium dioxide is
delivered to vacuoles when phagolysosomal fusion occurs (Fig. 6A and
B). By 1 h
postinoculation, thorium dioxide was found in 27% of the vacuoles
containing viable C. burnetii and 53% of the vacuoles
containing inactivated C. burnetii. The percentage of
labeled vacuoles containing viable C. burnetii began to
increase at 6 h postinoculation, and by 24 h it had reached
51%, while 66% was seen for inactivated C. burnetii. The
percentage of labeled VCB was compared to the labeling of vacuoles
containing inactivated serovar Enteritidis, a biodegradable control. At
1 h postinoculation 60% of the vacuoles that contained serovar
Enteritidis also contained thorium dioxide, and by 2 h postinoculation the serovar Enteritidis was completely degraded, indicating a rapid rate of phagolysosomal fusion and that the degradative processes of the J774A.1 cells were intact (Fig.
7A).
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FIG. 6.
(A) TEM image showing J774A.1 cells labeled with thorium
dioxide are shown at 2 h postinfection with viable C. burnetii. Two SCVs of C. burnetii, indicated by small
arrows, are visible within unfused vacuoles. Larger arrows indicate
lysosomes labeled with thorium. Bar, 500 nm. (B) Thorium dioxide is
visible within a vacuole containing an LCV of C. burnetii
6 h after infection. A small arrow indicates C. burnetii, and larger arrows indicate the thorium dioxide label.
Bar, 500 nm. (C) Acid phosphatase stain showing lysosomes wrapping
around C. burnetii at 4 h postinfection. Bar, 100 nm.
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FIG. 7.
Results of two experiments to determine the rate of
phagolysosomal fusion. (A) The lysosomes of J774A.1 cells were labeled
with the fluid-phase marker, thorium dioxide, and subsequently
inoculated with inactivated serovar Enteritidis ( ), inactivated
C. burnetii ( ), or viable C. burnetii ( ).
Vacuoles containing both inoculant and thorium dioxide were considered
to be fused. When cells were inoculated at 0 h, the percentage was
assumed to be zero. (B) Phagolysosomal fusion visualized by
histochemical staining of the lysosomal enzyme, acid phosphatase.
J774A.1 cells were inoculated with latex beads (), inactivated
C. burnetii (), or viable C. burnetii (),
and the percent fusion was determined as described above. At least 50 vacuoles were counted for each sample at each time point. Fusion was
determined at 24 h postinfection with thorium dioxide label, and
it was found that the fusion of vacuoles containing viable C. burnetii was slightly less than that seen for vacuoles containing
inactivated C. burnetii.
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The percentage of fused vacuoles was also monitored by determining the
presence of acid phosphatase. After inoculation of J774A.1 cells with
viable or inactivated C. burnetii or with latex beads, cells
were stained for acid phosphatase, a lysosomal enzyme that is delivered
to phagosomes when fusion takes place (Fig. 6C). Similar to results
seen with thorium labeling, at 1 h postinoculation 31% VCB were
labeled, while 52% of the vacuoles containing inactivated C. burnetii were labeled for acid phosphatase (Fig. 7B). At 1.5 h postinoculation, 70% of vacuoles containing latex beads, a
nondegradable control, were labeled, and this percentage remained
constant throughout the remainder of the experiment. Taken together,
the results of both of these methods clearly demonstrate that vacuoles
containing viable C. burnetii exhibit significantly reduced
phagolysosomal fusion during the first 6 h postinfection.
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DISCUSSION |
Although the ability of C. burnetii to proliferate
within the harsh environment of the phagolysosome is well established, little is known about processes that allow this intracellular parasite
to colonize this niche. The present study examines C. burnetii immediately after its internalization by the mouse
macrophage cell line J774A.1 and correlates pH to changes observed in
this pathogen. Previous investigators have shown that while
phagolysosomal fusion is necessary for C. burnetii
proliferation, the parasitopherous phagolysosome may be modified, as
demonstrated by a reduced oxidative burst (6). We show here
for the first time that phagolysosomal fusion is delayed by viable
C. burnetii but not by inactivated organisms. This raises
the question: if C. burnetii requires the phagolysosomal
environment for proliferation, why would phagolysosomal fusion be delayed?
We found that when J774A.1 cells were infected with a mixture of LCVs
and SCVs, within 6 h postinfection many vacuoles contained primarily LCVs, suggesting that this transition begins immediately after internalization during the time that phagolysosomal fusion is
delayed. The transition from SCV to LCV observed by electron microscopy
was quantitated using serum to an SCV-specific protein, ScvA. A
decrease in ScvA was clearly demonstrated in vivo and in vitro in
response to pH. ScvA has been shown to bind DNA, and it has been
proposed that it might play a part in the DNA condensation and
protection in the SCV (12). During the transition from SCV to LCV, DNA condensation and protection would no longer be necessary.
Several in vitro studies using C. burnetii mechanically
released from an intracellular environment have established its
acidophilic nature. The incorporation of glutamate and the
incorporation and catabolism of glucose are highest at a pH close to
4.5, while the catabolism and transport of glutamate and the transport
of glucose are maximal at pH 3 (11). Proline transport is
also maximal at pH 3 (14). Because it has been shown that
naturally released C. burnetii differs from mechanically
released organisms in DNA and protein synthesis (9, 28), we
have used naturally released organisms in this study to mimic the
population of C. burnetii available to infect cells in vivo.
In vitro incorporation assays were carried out to compare C. burnetii metabolism at the endosomal pH of 5.5 (32) and
at the phagolysosomal pH of 4.5 (19). We found that pH 5.5 is optimum for in vitro internalization of
[35S]methionine-[35S]cysteine by C. burnetii. To mimic the transition from the endosomal pH to the
phagolysosomal pH, C. burnetii was activated for 1 or 3 h at either pH 5.5 or 4.5 and then labeled with
[35S]methionine-[35S]cysteine at pH 4.5 for
2 h. There was no difference in incorporation or in proteins
expressed between samples activated at pH 5.5 and those activated at pH
4.5 (data not shown). These results may reflect the limitations of this
in vitro system or it is possible that, in vivo, C. burnetii
protein synthesis is reduced at pH 4.5. The in vitro pH optimum for
metabolic activities found here introduces the possibility that
C. burnetii may initially prefer a slightly less acidic environment.
Morphological differences between C. burnetii LCV and SCV
are suggestive of the morphological differences seen in other
gram-negative bacteria, when organisms in log phase are compared to
those in stationary phase. In both cases there is an increase in
lipopolysaccharide (4, 15), a thickening of the
peptidoglycan layer (3, 18), an increase in the protein
cross-linking of the outer membrane to the peptidoglycan (3,
29), and a decrease in outer membrane proteins (2, 3).
These comparisons suggest that the SCV may be similar to other
gram-negative bacteria in the stationary phase and that the LCV
morphology may be similar to that of gram-negative bacteria in the log
phase. In addition to these morphological differences it has been shown
that, when exposed to adverse conditions such as starvation, E. coli (26), Vibrio cholerae (10),
Salmonella serovar Enteritidis (26),
Campylobacter jejuni (25), Legionella pneumophila (16) and other organisms require a
resuscitation period before they resume proliferation. Release into the
extracellular environment by lysed host cells may impose stresses on
C. burnetii such that it also requires resuscitation prior
to proliferation in the phagolysosome.
This study reports for the first time a delay in phagolysosomal fusion
induced by viable C. burnetii upon infection of J774A.1 cells. Accompanying this delay is a transition from a population of
mixed LCVs and SCVs to one that is composed largely of LCVs. These
observations suggest that the current model of the C. burnetii developmental cycle should be modified to one in which
the transition from SCV to LCV begins in the endosome prior to
phagolysosomal fusion. A more complete understanding of the C. burnetii developmental cycle will lead to the development of a
means to interrupt this cycle and the subsequent control of C. burnetii infection.
We thank Michael Konkel for his technical advice and for his
assistance in the preparation of the manuscript. We thank Robert Heinzen for his generous gift of the anti-ScvA sera. We also thank to
Christine Davitt, Valerie Lynch-Holm, Vincent Franceschi, and the staff
at the Electron Microscopy Center at Washington State University.
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Abstract
Introduction
This paper is dedicated to the memory of Louis P. Mallavia.
