Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Matsuo, J. Articles by Yamaguchi, H. Search for Related Content PubMed PubMed Citation Articles by Matsuo, J. Articles by Yamaguchi, H. Agricola Articles by Matsuo, J. Articles by Yamaguchi, H.

 Previous Article  |  Next Article 

Applied and Environmental Microbiology, October 2008, p. 6397-6404, Vol. 74, No. 20
0099-2240/08/$08.00+0     doi:10.1128/AEM.00841-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Novel Parachlamydia acanthamoebae Quantification Method Based on Coculture with Amoebae

Junji Matsuo,¹ Yasuhiro Hayashi,¹ Shinji Nakamura,² Marie Sato,¹ Yoshihiko Mizutani,³ Masahiro Asaka,⁴ and Hiroyuki Yamaguchi^1*

Department of Medical Laboratory Sciences, Faculty of Health Sciences,¹ Department of Gastroenterology and Hematology, Graduate School of Medicine, Hokkaido University, Sapporo, Hokkaido 060-0812, Japan,⁴ Division of Biomedical Imaging Research,² Department of Human Pathology, Juntendo University Graduate School of Medicine, Tokyo 113-8421, Japan³

Received 8 April 2008/ Accepted 18 August 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Parachlamydia acanthamoebae, belonging to the order Chlamydiales, is an obligately intracellular bacterium that infects free-living amoebae and is a potential human pathogen. However, no method exists to accurately quantify viable bacterial numbers. We present a novel quantification method for P. acanthamoebae based on coculture with amoebae. P. acanthamoebae was cultured either with Acanthamoeba spp. or with mammalian epithelial HEp-2 or Vero cells. The infection rate of P. acanthamoebae (amoeba-infectious dose [AID]) was determined by DAPI (4',6-diamidino-2-phenylindole) staining and was confirmed by fluorescent in situ hybridization. AIDs were plotted as logistic sigmoid dilution curves, and P. acanthamoebae numbers, defined as amoeba-infectious units (AIU), were calculated. During culture, amoeba numbers and viabilities did not change, and amoebae did not change from trophozoites to cysts. Eight amoeba strains showed similar levels of P. acanthamoebae growth, and bacterial numbers reached ca. 1,000-fold (10⁹ AIU preculture) after 4 days. In contrast, no increase was observed for P. acanthamoebae in either mammalian cell line. However, aberrant structures in epithelial cells, implying possible persistent infection, were seen by transmission electron microscopy. Thus, our method could monitor numbers of P. acanthamoebae bacteria in host cells and may be useful for understanding chlamydiae present in the natural environment as human pathogens.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
During the last decade, chlamydiae, which are obligate intracellular bacterial pathogens, have been reclassified as the order Chlamydiales, which includes four families: Chlamydiaceae, Parachlamydiaceae, Waddliaceae, and Simkaniaceae (8). The Chlamydiaceae are well known to have a broad range of distribution in animals and humans and to be causative agents of human diseases (6, 34, 37, 50, 61). This family includes two major human pathogens: Chlamydophila pneumoniae is a causal agent of common respiratory infection and is also suspected of being involved in some chronic diseases, such as atherosclerosis (9), and Chlamydia trachomatis is responsible for sexually transmitted disease and preventable blindness (57). Chlamydiaceae species can infect most cultured human cells, including epithelial cells, endothelial cells, monocytes/macrophages, and lymphocytes (1, 22, 30, 53). They have a specific developmental cycle with alteration between an extracellular spore-like form, the elementary body (EB), and an intracellular metabolically active but noninfectious form, the reticulate body (RB) (43, 44, 54). The biological features of these chlamydiae, with respect to interaction with host cells and cell trafficking via the endocytic pathway, have well been studied (2, 14, 31, 58, 60, 63). Important to this research are reliable quantification methods to accurately determine the number of viable cells, such as the inclusion-forming unit (IFU) assay, as described by Ripa and Mardh (51).

Parachlamydiaceae, Waddliaceae, and Simkaniaceae species have recently been recognized as chlamydiae that show a wide distribution range in the natural environment, such as in rivers and in soil (15). All these species can survive within free-living amoebae, which are considered a reservoir for protecting bacteria from multiple environmental stresses (3, 17, 20, 29, 49, 56). Parachlamydia acanthamoebae and Simkania negevensis have been associated with lower respiratory tract infections (5, 38, 42), and Waddlia chondrophila, which was originally isolated from an aborted bovine fetus, is considered a potential abortigenic agent (17). In particular, there is accumulating evidence supporting the pathogenic role of P. acanthamoebae in humans (5, 15). Several studies have reported that parachlamydial DNA was detected by PCR in mononuclear cells of sputa and in bronchoalveolar lavage samples of a patient with bronchitis (16, 49). Other studies have suggested that P. acanthamoebae might be an agent of community-acquired pneumonia in human immunodeficiency virus-infected patients and in organ transplant recipients receiving immunosuppressive therapy (10, 11). P. acanthamoebae might also be a common agent of inhalation pneumonia (23, 24). Thus, P. acanthamoebae is emerging as a potential agent of respiratory tract infections. Moreover, recent work indicates that P. acanthamoebae possesses a specific developmental cycle consisting of the EB, the RB, and the crescent body (29) and that P. acanthamoebae can enter and multiply within human macrophages, pneumocytes, and lung fibroblasts and then induce apoptosis (13, 25, 27). A biological assay for determining the number of P. acanthamoebae bacteria inside host cells has been already established (27, 28). This method, which uses the mean number of bacteria per target cell or the highest dilution values that can still induce amoeba lysis, might be useful to roughly assess changes of bacterial numbers, but it is difficult to accurately determine the numbers of viable bacteria in cultures. Thus, for understanding the growth properties of P. acanthamoebae within mammalian cells as well as in amoebae, we present here a novel quantification method for P. acanthamoebae based on coculture with amoebae as well as an IFU assay suitable for Chlamydiaceae.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Eukaryotic cells.
Free-living amoebae (Acanthamoeba castellanii C3 and environmental isolates of Acanthamoeba spp.) and mammalian epithelial cells (human epithelial cell line HEp-2, derived from an epidermoid carcinoma of human larynx, and Vero cells, derived from the kidney of a normal adult African green monkey) were used for the present study. A. castellanii C3 (ATCC 50739) and HEp-2 cells (RCB 1889) were purchased from the American Type Culture Collection and the Riken Cell Bank, respectively. The eight strains of environmental Acanthamoeba spp. were isolated from soil and river samples from Sapporo City, Japan. Amoebae were maintained in broth including 0.75% peptone, 0.75% yeast extract, and 1.5% glucose (PYG) at 30°C (59). The epithelial cells were maintained in 10% fetal calf serum-Dulbecco modified Eagle medium at 37°C in 5% CO₂.

Bacteria.
P. acanthamoebae Bn₉ (ATCC VR-1476) was purchased from the American Type Culture Collection. The bacteria were propagated in the amoeba cell culture system according to methods described previously (29). In brief, the infected cells were harvested on day 7 and disrupted by freeze-thawing. After centrifugation at 180 x g for 5 min to remove cell debris, bacteria were concentrated by high-speed centrifugation at 3,500 x g for 30 min. The bacterial pellet was resuspended in sucrose-phosphate-glutamic acid buffer including 0.2 M sucrose, 3.8 mM KH₂PO₄, 6.7 mM Na₂HPO₄, and 5 mM L-glutamic acid (pH 7.4) and then stored at –80°C until used. The infective progeny was determined by the procedure described below.

Quantification procedure for infective progeny using coculture with amoebae.
After freeze-thawing, each sample containing viable P. acanthamoebae was serially diluted from 10⁰ to 10^–7 with PYG broth and incubated with A. castellanii (10⁴ or 10⁵ per well) in PYG broth with or without cycloheximide (200 µg/ml) on 96-well plates for 2 days. The infection rate of P. acanthamoebae for amoebae (amoeba-infectious dose [AID]) in each well was determined by microscopy at a magnification of x100, following DAPI (4',6-diamidino-2-phenylindole) staining. Ten fields were randomly selected for this assessment. The AIDs for a sample were plotted as a logistic sigmoid dilution curve using statistical software (KaleidaGraph 3.6; Hulinks, Tokyo, Japan). For logistic fitting, y = y_bottom + ( y_top – y_bottom)/[1 + (x/AID₅₀)^slope], as a function of the four-parameter logistic model described previously, was introduced (19). Because the AID values for each well are logistically varied in a range from 0 to 100%, the logistic function formula was simplified by plugging in y_bottom = 0 (0%) and y_top = 1 (100%), as follows: y = 1/[1 + (x/AID₅₀)^slope]. x and y show the value of the 10-fold dilution rate (the values vary from 10⁰ to 10⁷), and the amoeba-infectious rate (the values vary from 0% to 100%) corresponds to each value of the 10-fold dilution. Namely, the formula y = 1/[1 + (x/AID₅₀)^slope] with both sloped values of x and y logically draws a specific sigmoid curve via statistical software and shows a dilution rate corresponding to the mid-value of amoeba-infectious rate, AID₅₀. Finally, the viable bacterial numbers in cultures, defined as amoeba-infectious units (AIU), can be determined based on the value of AID₅₀.

Infection procedure for monitoring bacterial growth.
A 24-well plate with HEp-2 or Vero cells (5 x 10⁵/well) suspended in 10% fetal calf serum-Dulbecco modified Eagle medium or a six-well plate with amoebae (5 x 10⁵/well) suspended in PYG broth was infected with 5 x 10⁶ cells of P. acanthamoebae (multiplicity of infection of 1) by centrifugation at 700 x g for 1 h (the number of bacteria was determined by AIU assay). After centrifugation, cultures were resuspended in each medium and incubated for 10 days at 30°C in a normal atmosphere (for amoebae) or at 37°C under 5% CO₂ (for HEp-2 cells and Vero cells). During 10 days of culture, cells were regularly collected for determining cell numbers, assessing morphological changes and bacterial location in cells by electron microscopy, and determining viable bacterial numbers according to the method described above.

Cell counts.
Cell suspensions of infected amoeba cultures were collected at 0, 1, 4, 7, and 10 days after infection. Infected mammalian cells were detached from plates using cell dissociation solution, trypsin-EDTA (Sigma), and were collected. Cell suspensions were then used to determine viable cell counts by the trypan blue dye exclusion method.

Fluorescent in situ hybridization (FISH).
Oligonucleotide probes Bn₉658 (5'-TCC GTT TTC TCC GCC TAC-3'), specific for P. acanthamoebae 16S rRNA (3), and EUK516 (5'-ACC AGA CTT GCC CTC C-3'), targeted to eukaryote 18S rRNA (3), were used to confirm the location of bacteria in host cells. The Bn₉658 probe and the EUK516 probe were labeled with Alexa Fluor 350 (Invitrogen Corporation, Carlsbad, CA) and Alexa Fluor 488 (Invitrogen Corporation), respectively. Hybridizations were performed for 90 min at 46°C, according to methods described previously (20).

TEM.
For transmission electron microscopy (TEM), cells were immersed in a fixative containing 3% glutaraldehyde in 0.1 M phosphate-buffered saline, pH 7.4, for 24 h at 4°C. After a brief wash with phosphate-buffered saline, they were processed for alcohol dehydration and embedding in Epon 813, as described previously (64). Ultrathin sections of cells were stained with lead citrate and uranium acetate before being viewed by electron microscopy.

Statistical analysis.
Statistical analysis was performed with the F test (variation on values for AID₅₀) or with the unpaired Student t test (comparison on numbers of bacteria in culture). A P value of less than 0.05 was considered significant.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Establishment of quantification procedure.
Our experimental method is illustrated in Fig. 1. The method consisted of freeze-thawing of the sample (to release bacteria from host cells), infection and coculture with amoebae (to determine AIDs), and drawing a sigmoid curve by a fitting process with a logistic function formula (for calculating AIU).

View larger version (27K): [in this window] [in a new window]

FIG. 1. Brief summary of the experimental method.

The freeze-thawing method consistently ruptured mammalian cells and amoebae in the trophozoite stage but not cysts (data not shown). The optimal concentration of cycloheximide for keeping the number of amoebae constant for 5 days after infection without cytotoxicity was 200 µg/ml (data not shown). Figure 2A and B show representative images of DAPI-stained amoebae, with or without bacterial infection. The infected amoebae were easily distinguished from uninfected amoebae 2 days after infection. At this time, there was no difference in the size or the number of inclusions among infected amoebae. To confirm the DAPI staining data, FISH analysis with probes specific to P. acanthamoebae 16S rRNA was performed. As shown in Fig. 2C, D, and E, several specific clusters of spots, indicating inclusions, were observed in amoebae, indicating that the DAPI staining results were truly reflective of bacterial growth within amoebae. Secondary infection, via amoeba rupture, would make the accurate determination of bacterial numbers very difficult. Therefore, we determined the culture period that was less affected by secondary infection. Figure 3 shows a representative variation of DAPI-stained amoebae, with or without bacterial infection, during culture for up to 5 days. We observed no fluorescence particles outside amoebae for up to 2 days after infection. Thereafter, the number of particles outside amoebae increased with time, indicating that cell ruptures could cause secondary infection. At the same time, the number of amoebae decreased 3 days after infection, probably due to amoeba lysis (data not shown). Thus, we concluded that secondary infection in culture via amoeba rupture was minimal for up to 2 days after infection.

View larger version (24K): [in this window] [in a new window]

FIG. 2. Representative images of DAPI staining (A and B) and of FISH detection (C to E) of P. acanthamoebae-infected amoebae. (A and B) Amoebae infected with (A) or without (B) P. acanthamoebae were stained with DAPI at 2 days after infection. Magnification, x400. (C to E) Hybridizations with Alexa Fluor 488-labeled EUK516, which targets eukaryotic 18S rRNA (C) (green), and Alexa Fluor 350-labeled Bn9658, which is specific for P. acanthamoebae 16S rRNA (D) (purple). Merged images are shown in panel E. Magnification, x1,000.

View larger version (22K): [in this window] [in a new window]

FIG. 3. Representative variation of DAPI-stained amoebae with or without P. acanthamoebae infection. Magnification, x100.

Because the detection limit of the method can theoretically change according to the number of amoebae seeded in wells, we confirmed whether a change in the number of amoebae seeded in wells adversely affected AID₅₀ values. A sample was serially diluted and used to inoculate plates containing amoebae adjusted to either 10⁴ or 10⁵ per well. Then, AID values were determined following observation of DAPI staining, and two sigmoid curves were drawn based on the AID values. By comparing these sigmoid curves, we noted a rigorous declination with a log between the two sigmoid curves drawn through the fitting process (Fig. 4A). Thus, the numbers of amoebae seeded in wells were faithfully reflected in the change of infectious rates, depending on the concentration of bacteria in a sample. For drawing the sigmoid curve, a four-parametric logistic function, which is theoretically a symmetric curve with a cross-border value of AID₅₀, was introduced (19). This approach has the advantage that the curve can be drawn by limited plots rising initially. As shown in Fig. 4B, the variation of AID₅₀ that arose from different plot series was found. However, by plugging four consecutive plots into the function, including a value of more than 20% of the AID, the variance among the values of AID₅₀ was minimal. Thus, the established method defined as the AIU assay was a simple and accurate method to quantify infective progeny of P. acanthamoebae in host cells.

View larger version (20K): [in this window] [in a new window]

FIG. 4. Sigmoid curves drawn by a fitting process for determining AID50 (A) and the variation of AID50 derived from different plot series by plugging into the four-parametric logistic function (B). (A) The AIDs compliant with a diluted series of a sample were separately determined by using each plate containing amoebae adjusted to either 104 (closed circles) or 105 (open circles) per well, and two sigmoid curves were drawn by a fitting process with a four-parametric logistic function. Dilution rates equivalent to AID50, indicated by using the 104-per-well plate and the 105-per-well plate, were 2.616 ± 0.058 and 1.595 ± 0.043, respectively. There was rigorous declination with a log between the two sigmoid curves drawn through the fitting process. (B) The data represent the averages of AID50 ± standard deviations. *, P < 0.05 (variance) versus dilution series of 100 to 10–7.

Growth of P. acanthamoebae within amoebae and mammalian epithelial cells.
To determine quantitatively the growth of P. acanthamoebae within amoebae and mammalian epithelial cells, the number of bacterial infective progenies was assessed by the AIU assay for up to 10 days of culture. As shown in Fig. 5A, the numbers of bacteria in culture with amoebae increased significantly during the first 4 days of cultivation with numbers of bacteria reaching approximately 1,000-fold the initial number. There was no difference between the strains of amoebae that were used. Thereafter, bacterial numbers in the cultures remained constant. In contrast, no increase of bacterial numbers in the cultures with HEp-2 or Vero mammalian cells was observed, nor did the number of bacteria in the bacterial culture alone change significantly (Fig. 5B). In addition, the bacterial infection did not show any significant cytotoxicity to mammalian cells during culture, as determined by trypan blue dye exclusion (data not shown).

View larger version (25K): [in this window] [in a new window]

FIG. 5. Numbers of viable P. acanthamoebae bacteria in cultures of amoebae (A) and mammalian cells (B) for up to 10 days after infection. The number of bacteria was assessed by the AIU assay. The data represent the average AIU ± standard deviations. *, P < 0.05 versus values for bacteria alone.

Morphological analysis.
To confirm the accuracy of the AIU assay, the amoebae and mammalian epithelial cells infected with bacteria were examined by TEM. Figure 6 shows representative photo images of infected amoebae and mammalian cells. Several inclusions were present in amoeba infected with bacteria a day after inoculation, and the inclusions frequently contained dividing RBs (Fig. 6A). Four days after inoculation, enlarged inclusions were seen and the inclusions mostly contained many mature EBs (Fig. 6B). Figure 6A (arrows) also shows crescent bodies in inclusions, which are a specific stage in the developmental cycle of P. acanthamoebae, as reported by Greub and Raoult (29). In contrast, no typical inclusions were observed in mammalian cells infected with bacteria (Fig. 6C and D). However, RB-like structures, which might represent aberrant or persistent forms, were found in infected HEp-2 cells (Fig. 6C). In infected Vero cells, bacteria wrapped in membranes were seen, indicating an ongoing digestion of bacteria in vesicles (Fig. 6D). Thus, these results supported the findings obtained by the AIU assay that P. acanthamoebae grew normally in amoebae, but not in mammalian cells, such as HEp-2 and Vero cells.

View larger version (179K): [in this window] [in a new window]

FIG. 6. TEM images showing the ultrastructure of P. acanthamoebae-infected amoebae (A and B) and mammalian cells (C and D). (A) P. acanthamoebae-infected amoebae at 1 day after infection. The square shows an inclusion containing dividing RBs. Arrows indicate crescent bodies, which are a specific stage in the developmental cycle of P. acanthamoebae. (B) Amoebae infected with bacteria at 4 days after infection. The square shows an inclusion containing mature EBs. (C) HEp-2 cells infected with bacteria at 1 day after infection. Arrows show RB-like structures, which may represent aberrant or persistent forms. (D) Vero cells infected with bacteria at 1 day after infection. Bars, 500 nm.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Several assay systems for determining the number of P. acanthamoebae bacteria inside host cells have been established (10, 12, 13, 26, 27, 28, 45). The biological method originally proposed by Greub et al. is based on the mean number of bacteria per target cell or the highest dilution of bacteria that, via coculture with amoebae, led to complete amoeba lysis (27, 28). This quantification method has been widely used for analyzing antibiotic susceptibility, growth properties, and intracellular trafficking of P. acanthamoebae in host cells (13, 26, 27, 45). Recent work has demonstrated a real-time PCR assay for the specific detection of P. acanthamoebae DNA in samples (10, 12). However, because these assay systems are unlikely to accurately determine the number of viable bacteria in cultures, further studies are needed to develop a simple and accurate method to quantify P. acanthamoebae, analogous to the CFU assay for common bacteria. Thus, despite several quantification methods, the gold standard method for measuring infectious chlamydia progeny has traditionally been the IFU assay, based on counting large inclusions formed in cells (51). However, because P. acanthamoebae does not form typical inclusions within host human cells and inclusions formed in infected amoebae defined by fluorescence microscopic observation are relatively small compared with those in HEp-2 cells infected with C. pneumoniae and C. trachomatis, a simple method based on the IFU assay is not suitable (15). We therefore established an alternative method based on the rate of infection of amoebae, analogous to plaque-forming ability commonly used in the quantification of virus. The calculation of an index value of 50% infective rate has usually been performed according to the Reed and Muench method (52). However, these methods are required to use a wide range of values from 0% to 100%, and the steps for determining the values were formidable. Therefore, a four-parameter logistic function, as described previously (19), was introduced to the logistic fitting process. The drawing of logistic sigmoid dilution curves and the determination of AID₅₀ were performed by statistical software with the function in general use (e.g., KaleidaGraph).

The freeze-thawing step is the easiest method to release bacteria from cells such as amoebae. Because it is known that amoebae easily shift from trophozoite to cyst form, which is a resistant form to prevent damage from environmental stress and which is difficult to destroy by physical damage such as that from detergent, ultrasonic treatment, and freeze-thawing (36), we selected amoeba strains that would resist changing from trophozoite to cyst form during the 10-day culture period. As a result, amoebae were ruptured efficiently in more than ca. 95% of the cells. Meanwhile, several amoeba strains (Acanthamoebae spp.) in our stock collection isolated from the natural environment easily became cysts over short culture periods, and bacterial release from such cysts was very difficult to achieve. Moreover, although the presence of P. acanthamoebae bacteria as cysts in the natural environment is not supported by the fact that P. acanthamoebae bacteria were not found within cysts of acanthamoebae (29), it is possible that amoeba cysts harboring bacteria may be present in the natural environment. Thus, to apply this method to quantify the number of P. acanthamoebae bacteria within amoebae in environmental samples such as soil and water, it is necessary to include an additional short culture step to induce the change from cysts to trophozoites before freeze-thawing.

Our method requires that only amoebae infected with bacteria be detected; therefore, the culture period is critical. Thus, we carefully examined an optimal culture period without secondary infection. As a result, the infected amoebae were clearly distinguished from uninfected amoebae at 2 days after inoculation, and the rupture of amoebae with secondary infection was minimal for up to 2 days after inoculation. In the present study, we used the C3 strain (ATCC 50739) of amoebae as host cells for determining AIDs, because the strain could be easily purchased and its traits were well characterized (3, 7, 41, 47, 48). However, if other strains of amoebae are used, it may be necessary to determine an optimal culture period without secondary infection.

The detection limit of the method is dependent on the number of amoebae seeded in each well of the 96-well plate. Because two different numbers (10⁴ and 10⁵) of amoebae were seeded, the detection limit of each set was 2 x 10⁴ AIU and 2 x 10⁵ AIU, respectively. Even when the probable bacterial number in a sample is low, the bacterial number can be theoretically determined by our method using amoebae seeded at lower levels. However, seeding at low levels requires plates with smaller wells to maintain infectious efficiency.

The monitoring of P. acanthamoebae in the amoeba culture by the AIU assay showed that the numbers of bacterial infective progeny in amoebae increased rapidly for up to 4 days after infection and that the bacterial numbers in the culture remained constant during the remaining period. In order to confirm replication and growth within amoebae, TEM studies were performed at 1 day or 4 days after infection. This showed vacuoles containing RBs, EBs, and/or crescent bodies in amoebae, indicating bacterial proliferation and growth. Thus, these morphological traits were in good agreement with previous reports (15, 29), indicating that the culture condition used was optimal and that the values determined by the AIU assay were reflected in growth properties of P. acanthamoebae within amoebae.

It is well known that acanthamoebae are distributed in a broad range of environments, such as in rivers and in soil (4, 35, 36, 40, 46, 55). Acanthamoeba species have been classified into 15 different genotypes (T1 to T15) (21, 32, 33, 62). It has been reported that the T4 genotype, rather than the other genotypes, was frequently present in the natural environment and that 90% of acanthamoeba isolates belong to the T4 genotype, which is the most likely genotype to express strong virulence against humans (36, 39). These findings suggest that there is a possible variation for supporting growth of P. acanthamoebae among genotypes. However, we found no difference among traits of infective progeny between isolates (data not shown). Thus, the genotypes of Acanthamoeba species may be unrelated to P. acanthamoebae growth.

Serological studies suggest that P. acanthamoebae, as well as other chlamydiae, such as Waddliaceae and Simkaniaceae species, is associated with lower respiratory tract infections and that humans are commonly exposed to this pathogen (15). Although morphological studies also showed that P. acanthamoebae could enter and multiply within human pneumocytes (A549 cells), lung fibroblasts (HEL cells), and macrophages (peripheral blood macrophages) (25, 27, 29), the accurate numbers of P. acanthamoebae bacteria within human cells remain to be determined. Although it has been reported that Vero cells sustain P. acanthamoebae replication, the growth in Vero cells is controversial because complete data are not available (3, 45). Therefore, P. acanthamoebae growth within mammalian cells, such as in HEp-2 cells and Vero cells, which are representative for epithelial cells, was monitored by the AIU assay. However, no increase of bacterial numbers in the culture of either cell line was found during the culture period, nor was there an increase of bacteria in the culture with bacteria alone. In order to confirm whether P. acanthamoebae can enter and multiply within such cells, TEM studies were performed. No typical inclusion-containing bacteria in infected cells were observed; however, unique structures in the cytoplasm of HEp-2 cells infected with bacteria, but not in that of uninfected cells, were observed. Such structures had a plasma membrane and were not surrounded by vesicles, like typical inclusions formed by chlamydiae. Interestingly, the traits observed appeared to be extremely similar to the morphological finding of chlamydiae in Drosophila melanogaster S2 cells, as reported by Elwell and Engel (18). This suggests that unique structures observed in the infected cells may be abnormal bodies defined as a persistent form that have differentiated from EBs and that are unable to form infectious progeny. This may explain the results obtained by the AIU assay, which showed unchanged bacterial numbers in culture. The reasons for persistent infection are unknown, but it may be speculated that mammalian cells respond to P. acanthamoebae infection and produce cytokines, which may to some degree control bacterial growth in the cells, even though a lack of cytokine response in human macrophages infected with P. acanthamoebae has been reported (25). In addition, bacteria wrapped in membranes in infected Vero cells were seen, indicating an ongoing digestion of bacteria in vesicles. This finding may imply that P. acanthamoebae can enter but not grow inside Vero cells.

We present a method to quantify the infective progeny of P. acanthamoebae, based on coculture with amoebae. The method is a promising tool for monitoring exact numbers of P. acanthamoebae bacteria within host cells, comparable to the IFU assay available for chlamydiae such as C. pneumoniae and C. trachomatis. We also believe that this method would be suitable for other chlamydiae present in the natural environment such as Waddliaceae and Simkaniaceae species. Thus, the method established in this study may be useful for understanding the dynamism of these chlamydiae with respect to potential human pathogen behavior.

 
This work was supported by a grant-in-aid for scientific research from the Waksman Foundation and the Asahi Glass Foundation of Japan.

We thank Gilbert Greub (Institute of Microbiology, Faculty of Biology and Medicine, University of Lausanne, Lausanne, Switzerland) for providing P. acanthamoebae DNA that we used to perform preliminary experiments.

 
* Corresponding author. Mailing address: Department of Medical Laboratory Sciences, Faculty of Health Sciences, Hokkaido University, Nishi-5 Kita-12 Jo, Kita-ku, Sapporo, Hokkaido 060-0812, Japan. Phone and fax: 81-11-706-3326. E-mail: hiroyuki{at}med.hokudai.ac.jp

Published ahead of print on 29 August 2008.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Airenne, S. H., H. M. Surcel, H. Alakärppä, K. Laitinen, J. Paavonen, P. Saikku, and A. Laurila. 1999. Chlamydia pneumoniae infection in human monocytes. Infect. Immun. 67:1445-1449.[Abstract/Free Full Text]
2
2. Al-Younes, H. M., T. Rudel, and T. F. Meyer. 1999. Characterization and intracellular trafficking pattern of vacuoles containing Chlamydia pneumoniae in human epithelial cells. Cell. Microbiol. 1:237-247.[CrossRef][Medline]
3
3. Amann, R., N. Springer, W. Schönhuber, W. Ludwig, E. N. Schmid, K. Müller, and R. Michel. 1997. Obligate intracellular bacterial parasites of Acanthamoebae related to Chlamydia spp. Appl. Environ. Microbiol. 63:115-121.[Abstract]
4
4. Armstrong, M. 2000. The pathogenesis of human Acanthamoeba infection. Infect. Dis. Rev. 2:65-73.
5
5. Birtles, R. J., T. J. Rowbotham, C. Storey, T. J. Marrie, and D. Raoult. 1997. Chlamydia-like obligate parasite of free-living amoebae. Lancet 29:925-926.
6
6. Bodetti, T. J., and P. Timms. 2000. Detection of Chlamydia pneumoniae DNA and antigen in the circulating mononuclear cell fraction of humans and koalas. Infect. Immun. 68:2744-2747.[Abstract/Free Full Text]
7
7. Bowers, B., and E. D. Korn. 1968. The fine structure of Acanthamoeba castellanii. J. Cell Biol. 39:95-111.[Abstract/Free Full Text]
8
8. Bush, R. M., and K. D. E. Everett. 2001. Molecular evolution of the Chlamydiaceae. Int. J. Syst. Evol. Microbiol. 51:203-220.[Abstract]
9
9. Campbell, L. A., and C. C. Kuo. 2003. Chlamydia pneumoniae and atherosclerosis. Semin. Respir. Infect. 18:48-54.[CrossRef][Medline]
10
10. Casson, N., J. M. Entenza, N. Borel, A. Pospischil, and G. Greud. 2008. Murin model of pneumonia caused by Parachlamydia acanthamoebae. Microb. Pathog. 42:92-97.
11
11. Casson, N., and G. Greub. 2006. Resistance of different Chlamydia-like organisms to quinolones and mutations in the quinoline resistance-determining region of the DNA gyrase A- and topoisomerase-encoding gene. Int. J. Antimicrob. Agents 27:541-544.[CrossRef][Medline]
12
12. Casson, N., K. M. Posfay-Barbe, A. Gervaix, and G. Greub. 2008. New diagnostic real-time PCR for specific detection of Parachlamydia acanthamoebae DNA in clinical samples. J. Clin. Microbiol. 46:1491-1493.[Abstract/Free Full Text]
13
13. Casson, N., N. Medico, J. Bille, and G. Greub. 2006. Parachlamydia acanthamoebae enters and multiples within pneumocytes and human fibroblasts. Microb. Infect. 8:1294-1300.[CrossRef][Medline]
14
14. Coombes, B., and J. B. Mahony. 2002. Identification of MEK- and phosphoinositide 3-kinase-dependent signaling as essential events during Chlamydia pneumoniae invasion of HEp-2 cells. Cell. Microbiol. 4:447-460.[CrossRef][Medline]
15
15. Corsaro, D., and G. Greub. 2006. Pathogenic potential of novel chlamydiae and diagnostic approaches to infections due to these obligate intracellular bacteria. Clin. Microbiol. Rev. 19:283-297.[Abstract/Free Full Text]
16
16. Corsaro, D., D. Venditti, and M. Valassina. 2002. New parachlamydial 16S rDNA phylotypes detected in human clinical samples. Res. Microbiol. 153:563-567.[Medline]
17
17. Dilbeck, P. M., J. F. Evermann, T. B. Crawford, A. C. Ward, C. W. Leathers, C. J. Holland, C. A. Mebus, L. L. Logan, F. R. Rurangirwa, and T. C. McGuire. 1990. Isolation of a previously undescribed rickettsia from an aborted bovine fetus. J. Clin. Microbiol. 28:814-816.[Abstract/Free Full Text]
18
18. Elwell, C., and J. N. Engel. 2005. Drosophila melanogaster S2 cells: a model system to study Chlamydia interaction with host cells. Cell. Microbiol. 7:725-739.[CrossRef][Medline]
19
19. Finney, D. J. 1983. Response curves for radioimmunoassay. Clin. Chem. 29:1763-1766.
20
20. Fritsche, T. R., M. Horn, M. Wagner, R. P. Herwig, K. Schleifer, and R. K. Gauton. 2000. Phylogenetic diversity among geographically dispersed Chlamydiales endosymbionts recovered from clinical and environmental isolates of Acanthamoeba spp. Appl. Environ. Microbiol. 66:2613-2619.[Abstract/Free Full Text]
21
21. Gast, R. J. 2001. Development of an Acanthamoeba specific reverse dot-blot and the discovery of a new ribotype. J. Eukaryot. Microbiol. 48:609-615.[CrossRef][Medline]
22
22. Gaydos, C. A., J. T. Summersgill, N. N. Shnet, J. A. Ramirez, and T. C. Quinn. 1996. Replication of Chlamydia pneumoniae in vitro in human macrophages, endothelial cells, and aortic artery smooth muscle cells. Infect. Immun. 64:1614-1620.[Abstract]
23
23. Greub, G., P. Berger, L. Papazian, and D. Raoult. 2003. Parachlamydiaceae as rare agents of pneumonia. Emerg. Infect. Dis. 9:755-756.[Medline]
24
24. Greub, G., I. Boyadjiev, B. La Scola, D. Raoult, and C. Martin. 2003. Serological hint suggesting that Parachlamydiaceae are agents of pneumonia in polytraumatized intensive care patients. Ann. N. Y. Acad. Sci. 990:311-319.[CrossRef][Medline]
25
25. Greub, G., B. Desnues, D. Raoult, and J. Mega. 2005. Lack of microbicidal response in human macrophages infected with Parachlamydia acanthamoebae. Microb. Infect. 7:714-719.[Medline]
26
26. Greub, G., J. Mege, J. Gorvel, D. Raoult, and S. Méresse. 2005. Intracellular trafficking of Parachlamydia acanthamoebae. Cell. Microbiol. 7:581-589.[CrossRef][Medline]
27
27. Greub, G., J. Mega, and D. Raoult. 2003. Parachlamydia acanthamoeba enters and multiplies within human macrophages and induces their apoptosis. Infect. Immun. 71:5979-5985.[Abstract/Free Full Text]
28
28. Greub, G., B. La Scola, and D. Raoult. 2003. Parachlamydia acanthamoeba is endosymbiotic or lytic for Acanthamoeba polyphaga depending on the incubation temperature. Ann. N. Y. Acad. Sci. 990:628-634.[Medline]
29
29. Greub, G., and D. Raoult. 2002. Crescent bodies of Parachlamydia acanthamoeba and its life cycle within Acanthamoeba polyphaga: an electron micrograph study. Appl. Environ. Microbiol. 68:3076-3084.[Abstract/Free Full Text]
30
30. Haranaga, S., H. Yamaguchi, H. Friedman, S. Izumi, and Y. Yamamoto. 2001. Chlamydia pneumoniae infects and multiplies in lymphocytes in vitro. Infect. Immun. 69:7753-7759.[Abstract/Free Full Text]
31
31. Heinzen, R. A., M. A. Scidmore, D. D. Rockey, and T. Hackstadt. 1996. Differential interaction with endocytic and exocytic pathways distinguish parasitophorous vacuoles of Coxiella burnetii and Chlamydia trachomatis. Infect. Immun. 64:796-809.[Abstract]
32
32. Hewett, M. K., B. S. Robinson, P. T. Monis, and C. P. Saint. 2003. Identification of a new Acanthamoeba 18S rRNA gene sequence type corresponding to the species Acanthamoeba jacobsi Sawyer, Nerad and Visvesvara, 1992 (Lobosea: Acanthamoebidae). Acta Protozool. 42:325-329.
33
33. Horn, M., T. R. Fritsche, R. K. Gautom, and M. Wagner. 1999. Novel bacterial endosymbionts of Acanthamoeba spp. related to the Paramecium caudatum symbiont Caedibacter caryphilus. Environ. Microbiol. 1:357-367.[CrossRef][Medline]
34
34. Jackson, M., N. White, P. Giffard, and P. Timms. 1999. Epizootiology of Chlamydia infections in two free-range koala populations. Vet. Microbiol. 65:255-264.[CrossRef][Medline]
35
35. Khan, N. A. 2003. Pathogenesis of Acanthamoeba infections. Microb. Pathog. 34:277-285.[CrossRef][Medline]
36
36. Khan, N. A. 2006. Acanthamoeba: biology and increasing importance in human health. FEMS Microbiol. Rev. 30:564-595.[CrossRef][Medline]
37
37. Kutlin, A., P. M. Roblin, S. Kumar, S. Kohlhoff, T. Bodetti, P. Timms, and M. R. Hammerschlag. 2007. Molecular characterization of Chlamydophila pneumoniae isolates from Western barred bandicoots. J. Med. Microbiol. 56:407-417.[Abstract/Free Full Text]
38
38. Lieberman, D., S. Kahane, and M. G. Friedman. 1997. Pneumonia with serological evidence of acute infection with the Chlamydia-like microorganism "Z." Am. J. Respir. Crit. Care Med. 156:578-582.[Abstract/Free Full Text]
39
39. Maghsood, A. H., J. Sissons, M. Rezaian, D. Nolder, D. Warhurst, and N. A. Khan. 2005. Acanthamoeba genotype T4 from the UK and Iran and isolation of the T2 genotype from clinical isolates. J. Med. Microbiol. 54:755-759.[Abstract/Free Full Text]
40
40. Marciano-Cabral, F., R. Puffenbauer, and G. A. Cabral. 2000. The increasing importance of Acanthamoeba infections. J. Eukaryot. Microbiol. 47:29-36.[CrossRef][Medline]
41
41. Marolda, C. L., B. Hauröder, M. A. John, R. Michel, and M. A. Valvano. 1999. Intracellular survival and saprophytic growth of isolates from the Burkholderia cepacia complex in free-living amoebae. Microbiology 145:1509-1517.[Abstract/Free Full Text]
42
42. Marrie, T. J., D. Raoult, B. La Scola, R. J. Birtles, and E. de Carolis. 2001. Legionella-like and other amoebal pathogens as agents of community-acquired pneumonia. Emerg. Infect. Dis. 7:1026-1029.[Medline]
43
43. Matsumoto, A., N. Higashi, and A. Tamura. 1973. Electron microscope observation on the effects of polymyxin B sulfate on cell walls of Chlamydia psittaci. J. Bacteriol. 113:357-364.[Abstract/Free Full Text]
44
44. Matsumoto, A., and G. P. Manire. 1970. Electron microscopic observations on the fine structure of cell walls of Chlamydia psittaci. J. Bacteriol. 104:1332-1337.[Abstract/Free Full Text]
45
45. Maurin, M., D. Bryskier, and D. Raoult. 2002. Antibiotic susceptibilities of Parachlamydia acanthamoeba in amoebae. Antimicrob. Agents Chemother. 46:3065-3067.[Abstract/Free Full Text]
46
46. Mergeryan, H. 1991. The prevalence of Acanthamoeba in the human environment. Rev. Infect. Dis. 5:390-391.
47
47. Michel, R., K. D. Müller, R. Amann, and E. N. Schmid. 1998. Legionella-like slender rods multiplying within a strain of Acanthamoeba sp. isolated from drinking water. Parasitol. Res. 84:84-88.[CrossRef][Medline]
48
48. Michel, R., K. D. Müller, and R. Hoffmann. 2001. Enlarged Chlamydia-like organisms as spontaneous infection of Acanthamoeba castellanii. Parasitol. Res. 87:248-251.[CrossRef][Medline]
49
49. Ossewaarde, J. M., and A. Meijer. 1999. Molecular evidence for the existence of additional members of the order Chlamydiales. Microbiology 145:411-417.[Abstract/Free Full Text]
50
50. Peeling, R. W., and R. C. Brunham. 1996. Chlamydiae as pathogens: new species and new issues. Emerg. Infect. Dis. 2:307-319.[Medline]
51
51. Ripa, K. T., and P. Mårdh. 1977. Cultivation of Chlamydia trachomatis in cycloheximide-treated McCoy cells. J. Clin. Microbiol. 6:328-331.[Abstract/Free Full Text]
52
52. Reed, L. J., and H. Muench. 1938. A simple method of estimating fifty percent endpoints. Am. J. Hyg. 27:493-497.
53
53. Roblin, P. M., W. Dumornay, and M. R. Hammerschlag. 1992. Use of HEp-2 cells for improved isolation and passage of Chlamydia pneumoniae. J. Clin. Microbiol. 30:1968-1971.[Abstract/Free Full Text]
54
54. Rockey, D. D., and A. Matsumoto. 2000. The chlamydial developmental cycles, p. 403-425. In Y. V. Brun and L. L. Shimkets (ed.), Prokaryotic development. ASM Press, Washington, DC.
55
55. Rodríguez-Zaragoza, S. 1994. Ecology of free-living amebae. Crit. Rev. Microbiol. 20:225-241.[Medline]
56
56. Rurangirwa, F. R., P. M. Dilbeck, T. B. Crawford, T. C. McGuire, and T. F. McElwain. 1999. Analysis of the 16S rRNA gene of micro-organism WSU 86-1044 from an aborted bovine foetus reveals that it is a member of the order Chlamydiales: proposal of Waddliaceae fam. nov., Waddlia chondrophila gen. nov., sp. nov. Int. J. Syst. Bacteriol. 49:577-581.[Abstract/Free Full Text]
57
57. Schachter, J. 1988. The intracellular life of Chlamydia. Curr. Top. Microbiol. Immunol. 138:109-139.[Medline]
58
58. Schramm, H., C. R. Bagnell, and P. B. Wyrick. 1996. Vesicles containing Chlamydia trachomatis serovar L2 remain above pH6 within HEC-1B cells. Infect. Immun. 64:1208-1214.[Abstract]
59
59. Schuster, F. L. 2002. Cultivation of pathogenic and opportunistic free-living amebas. Clin. Microbiol. Rev. 15:342-354.[Abstract/Free Full Text]
60
60. Stephens, R. S., K. Koshiyama, E. Lewis, and A. Kubo. 2001. Heparin-binding outer membrane protein of chlamydiae. Mol. Microbiol. 40:691-699.[CrossRef][Medline]
61
61. Storey, C., M. Lusher, P. Yates, and S. Richmond. 1993. Evidence for Chlamydia pneumoniae of non-human origin. J. Gen. Microbiol. 139:2621-2626.[Abstract/Free Full Text]
62
62. Stothard, D. R., J. M. Schroeder-Diedrich, M. H. Awwad, R. J. Gast, D. R. Ledee, S. Rodríguez-Zaragoza, C. L. Dean, P. A. Fuerst, and T. J. Byers. 1998. The evolutionary history of the genus Acanthamoeba and the identification of eight new 18S rRNA gene sequence types. J. Eukaryot. Microbiol. 45:45-54.[Medline]
63
63. Taraska, T., D. M. Ward, R. S. Ajioka, P. B. Wyrick, S. R. Davis-Kaplan, C. H. Davis, and J. Kaplan. 1996. The late chlamydial inclusion membrane is not derived from the endocytic pathway and is relatively deficient in host proteins. Infect. Immun. 64:3713-3727.[Abstract]
64
64. Uruma, T., H. Yamaguchi, M. Fukuda, H. Kawakami, H. Goto, T. Kishimoto, Y. Yamamoto, A. Tomoda, and S. Kamiya. 2005. Chlamydia pneumoniae growth inhibition in human monocytic THP-1 cells and human epithelial HEp-2 cells by a novel phenoxazine derivative. J. Med. Microbiol. 54:1143-1149.[Abstract/Free Full Text]

---

Applied and Environmental Microbiology, October 2008, p. 6397-6404, Vol. 74, No. 20
0099-2240/08/$08.00+0     doi:10.1128/AEM.00841-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Matsuo, J. Articles by Yamaguchi, H. Search for Related Content PubMed PubMed Citation Articles by Matsuo, J. Articles by Yamaguchi, H. Agricola Articles by Matsuo, J. Articles by Yamaguchi, H.