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 Citation Map Services E-mail this article to a friend 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 Barbosa, J. Articles by Pina-Vaz, C. Search for Related Content PubMed PubMed Citation Articles by Barbosa, J. Articles by Pina-Vaz, C.

 Previous Article  |  Next Article 

Clinical and Vaccine Immunology, July 2009, p. 1021-1024, Vol. 16, No. 7
1071-412X/09/$08.00+0     doi:10.1128/CVI.00031-09
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Cytometric Approach for Detection of Encephalitozoon intestinalis, an Emergent Agent

Joana Barbosa,^1,2* Acácio Gonçalves Rodrigues,^1,3,4 and Cidália Pina-Vaz^1,4,5

Department of Microbiology, Faculty of Medicine, University of Porto, Porto,¹ Escola Superior de Saúde Jean Piaget, Vila Nova Gaia,² Burn Unit, Department of Plastic and Reconstructive Surgery, Hospital S. João, Porto,³ Cardiovascular Research and Development Unit, Faculty of Medicine, University of Porto, Porto,⁴ Department of Microbiology, Hospital S. João, Porto, Portugal⁵

Received 9 January 2009/ Returned for modification 20 March 2009/ Accepted 6 May 2009

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Encephalitozoon intestinalis is responsible for intestinal disease in patients with AIDS and immunocompetent patients. The infectious form is a small spore that is resistant to water treatment procedures. Its detection is very important, but detection is very cumbersome and time-consuming. Our main objective was to develop and optimize a specific flow cytometric (FC) protocol for the detection of E. intestinalis in hospital tap water and human feces. To determine the optimal specific antibody (Microspor-FA) concentration, a known concentration of E. intestinalis spores (Waterborne, Inc.) was suspended in hospital tap water and stool specimens with different concentrations of Microspor-FA, and the tap water and stool specimens were incubated under different conditions. The sensitivity limit and specificity were also evaluated. To study spore infectivity, double staining with propidium iodide (PI) and Microspor-FA was undertaken. Distinct approaches for filtration and centrifugation of the stool specimens were used. E. intestinalis spores stained with 10 µg/ml of Microspor-FA at 25°C overnight provided the best results. The detection limit was 5 x 10⁴ spores/ml, and good specificity was demonstrated. Simultaneous staining with Microspor-FA and PI ensured that the E. intestinalis spores were dead and therefore noninfectious. With the stool specimens, better spore recovery was observed with a saturated solution of NaCl and centrifugation at 1,500 x g for 15 min. A new approach for the detection of E. intestinalis from tap water or human feces that ensures that the spores are not viable is now available and represents an important step for the prevention of this threat to public health.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Microsporidia comprise a diverse group of eukaryotic obligate intracellular parasites (12), including over 140 genera corresponding to more than 1,200 species. Only eight genera have clinical importance and are considered human pathogens (9, 25). Enterocytozoon bieneusi and Encephalitozoon intestinalis, both of which are responsible for gastrointestinal disease, are currently the most prevalent microsporidia identified in humans (6, 16) and among other mammals (12).

The true prevalence of microsporidiosis is not yet defined due to underdiagnosis, especially in immunocompetent individuals (13, 14). Only a small number of cases of Encephalitozoon intestinalis infection have been documented (25); E. intestinalis (reclassified from Septata intestinalis) has been detected in AIDS patients in developed countries and in Africa, as well as in other immunocompromised individuals, including patients who have undergone transplantation (8, 9, 24).

The infectious form is a small spore (1.8 to 5.0 µm), which is the only recognized viable stage of microsporidia outside a host cell. It has two rigid extracellular walls made of chitin, thus suggesting a potential link to a fungal cell (9, 12). The clinical manifestations of E. intestinalis infections in immunocompetent patients range from asymptomatic infections to self-limited diarrhea. However, in immunocompromised patients, it causes chronic diarrhea, which tends to disseminate, with the kidney being the major organ affected (8, 9). Spores may spread to the environment from infected patients via feces, urine, and/or other body fluids and tissues (9, 24). Consequently, the routes of transmission may involve person-to-person contact as well as waterborne or food-borne contamination, especially in developing countries with poor sanitation (11, 25). The microsporidial spores are usually very resistant to environmental conditions (24) and to the usual water treatment procedures (10), and they remain infective for long periods of time, especially when they are protected from desiccation (9). As a result of these characteristics, the U.S. Environmental Protection Agency placed microsporidia in first place as a candidate contaminant for drinking water (10). However, the infectious load needed to cause disease is not yet known.

The detection of spores in human feces or other human body fluids is very cumbersome and difficult. During the last 20 years, several methods for the recovery and detection of microsporidial spores have been developed and improved; these include electron microscopy and histologic examination of tissues samples (25). Light and/or immunofluorescence microscopy with polyclonal or monoclonal antibodies directed against microsporidial spores is nevertheless the most commonly used procedure, especially for the diagnosis of infection in immunocompromised patients (8, 9, 16, 25). Such methods are very time-consuming and are subject to human error (8, 16). Although molecular studies are highly sensitive and specific, they are too complex, too difficult for use for routine analysis, and expensive and are unable to assess spore viability (24). Flow cytometry (FC) allows both morphofunctional evaluation and quantification of individual microorganisms; it additionally provides significant advantages and a high degree of specificity, especially when it is combined with specific monoclonal or polyclonal antibodies against the E. intestinalis spore wall (8, 9, 16, 25). Our research team has been exploring distinct applications of FC in microbiology in order to increase its diagnostic sensitivity with clinical samples (17, 19) and to evaluate its use for determination of the susceptibility profiles of microorganisms (18, 20, 21, 22). Our main objective was to develop and optimize a specific FC protocol for the detection of E. intestinalis in hospital tap water and human feces after the simulation of the conditions in environmental and clinical settings.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Optimization of fluorescent staining. Encephalitozoon intestinalis Didier from culture in rabbit RK-13 cells that had been stored in 5% formalin at a concentration of 1 x 10⁶/ml (Waterborne, Inc., New Orleans, LA) was used. A total of 1 x 10⁵ spores/ml of E. intestinalis were suspended in 800 µl of sterile water and stained with serial concentrations (0.5, 1.0, 2.0, 5.0, and 10.0 µg/ml) of an Alexa Fluor 488 mouse monoclonal antibody (Microspor-FA; 20x concentrate; A700 AF488; Waterborne, Inc.), followed by incubation in the dark at different temperatures (37°C and 25°C) and for different times (45, 60, 90, and 180 min and overnight). Stool samples were centrifuged at distinct velocities (1,000 x g and 1,500 x g) and times (5 to 15 min). The supernatant was discarded and the pellet was resuspended in 1 ml of sterile H₂O, stained with the monoclonal antibody, and vortexed for 30 s. It was then transferred to a propylene tube and analyzed by FC at FL1 (green fluorescence, 535 nm).

FC analysis. The optical characteristics of the spore suspensions were evaluated on a FACSCalibur cytometer (standard model; BD Biosciences, Sydney, Australia) equipped with three photomultipliers with standard filters (FL1, band pass of 530/30 nm; FL2, band pass of 585/42 nm, FL3, long pass of 650 nm) and a 15-mW 488-nm argon laser. Cell Quest Pro software (version 4.0.2; BD Biosciences) was used to evaluate the results. Acquisition settings were defined with an unstained sample (autofluorescence), and the photomultiplier voltage was adjusted to the first logarithmic decade. Instrument controls followed the standard described procedures.

Assessment of sensitivity and specificity. To assess the sensitivity, serial concentrations of E. intestinalis spores (1 x 10³ to 5 x 10⁵/ml) were stained with the previously optimized antibody concentration and were analyzed by FC.

To assess the specificity, 1 x 10⁵ spores/ml of E. intestinalis were mixed with (i) bacterial suspensions consisting of 1.5 x 10⁸ cells/ml of Escherichia coli ATCC 35218 or 1.5 x 10⁸ cells/ml of Staphylococcus aureus ATCC 25923; (ii) fungal suspensions consisting of 5 x 10⁶ blastoconidia/ml of Candida albicans ATCC 10231; or (iii) parasite suspensions consisting of 2 x 10⁵ cysts or oocysts/ml of Giardia lamblia and Cryptosporidium parvum purchased from Waterborne, Inc.

The mixture was stained with the microsporidial monoclonal antibody (Microspor-FA; 20x concentrate; A700 AF488; Waterborne, Inc.) conjugated with Alexa Fluor 488 at the previously optimized concentration and incubated overnight at 25°C in the dark. After incubation, the cell suspensions were vortexed for 30 s, transferred to a propylene vial, and analyzed by FC.

Staining for live and dead spores. Propidium iodide (PI; Sigma) at 5.0 µg/ml was used to stain E. intestinalis spores with and without the specific fluorescent monoclonal antibody in the dark at different temperatures (37°C and 25°C) and times (45, 60, 90, and 180 min and overnight). Following the staining of the spores, the parasite suspensions were vortexed for 30 s, transferred to a propylene tube, and analyzed by FC at FL3 (red fluorescence, 670 nm).

Evaluation of human stool samples. Stool samples from healthy human volunteers were collected and stored at 4°C until use. One gram of stool sample was diluted with 5 ml of deionized sterile water and filtered through double gauze, and a known concentration of E. intestinalis spores (10⁵ spores/ml) was deposited on the filter. Deionized sterile water was then added to obtain a final 5-ml volume of filtrate. The material retained on the filter was collected and centrifuged in sterilized conical propylene tubes. After the first centrifugation, the supernatant was decanted into another sterilized tube. For further studies; sterile water was added to the tube, as performed with the original filtrate, up to a 5-ml volume, followed by a new centrifugation cycle. After the supernatant was decanted into conical tubes, sterile water was again added, and the process was repeated until a clean supernatant was obtained in both vials.

Several distinct procedures were compared with the aim of reducing the loss of spores in samples and to improve spore detection: centrifugation at different times (10 and 15 min) and velocities (1,000 x g and 1,500 x g), the use of distinct NaCl flotation solutions (solution A, NaCl at 360 g/liter and specific gravity of 1.21; solution B, saturated NaCl) and ZnSO₄·7H₂O (33%; 703 g/liter; specific gravity, 1.118), and the use of different incubation times before recovery (90 min and 24 h). After flotation, the upper 1 ml of supernatant and the upper 1 ml of sediment were transferred to Eppendorf tubes and stained with the specific monoclonal antibody (Waterborne, Inc.), followed by incubation (at 37°C for 90 min or at 25°C overnight) in the dark. FC analysis was performed by using the previously optimized conditions.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Optimization of FC protocol. With increasing antibody concentrations, an increase in the mean intensity of fluorescence (MIF) was evident when 10⁵ E. intestinalis spores/ml were stained and incubated overnight at room temperature and in the dark (Fig. 1). The use of the specific antibody at 10 µg/ml resulted in the highest MIF for the stained spores (10⁵ spores/ml). Overnight incubation at room temperature (25°C) in the dark resulted in a higher level of fluorescence than incubation at 37°C for 90 min.

View larger version (14K): [in this window] [in a new window]

FIG. 1. MIF of Encephalitozoon intestinalis spores labeled with serial concentrations of an Alexa Fluor 488 monoclonal antibody (Microspor-FA; 20x concentrate; A700 A488; Waterborne, Inc.). All experiments were performed twice.

Assessment of sensitivity and specificity. A decrease in the MIF was registered with a reduction in the spore concentration. A detection limit of 5 x 10⁴ spores/ml was established, since below that value the fluorescence intensity was not enough to allow discrimination of the spores. No interference with fungi or parasites occurred. Because of the similar sizes of the spores and bacteria, more events on the scatter were found when bacterial suspensions were used, although they were not stained, thus demonstrating the specificity of the staining.

Staining for live and dead spores. To confirm that the E. intestinalis spores studied were dead, PI staining was used simultaneously with the specific fluorescent antibody. When both fluorescent probes were used, dead E. intestinalis spores could be distinguished from other organisms (Fig. 2).

View larger version (15K): [in this window] [in a new window]

FIG. 2. Two-dimensional dot plot correlating FL1 (green fluorescence, 535 nm) with FL3 (red fluorescence, 620 nm) of Encephalitozoon intestinalis spores without staining (autofluorescence) (A), E. intestinalis spores stained with 10.0 µg/ml of specific antibody (Microspor-FA; A488; Waterborne, Inc.) (B), and E. intestinalis spores stained with 10.0 µg/ml of specific antibody and 5.0 µg/ml of PI (C).

Evaluation of stool samples. A better separation of the stained spores from debris was obtained with the use of a saturated NaCl solution rather than a ZnSO₄·7H₂O solution. Regarding the incubation times for recovery, similar results were obtained after 90 min and 24 h. The optimal speed of centrifugation that allowed the recovery of a good percentage of spores was 1,500 x g for 15 min.

Similar results were obtained when the results obtained with the upper 1 ml of the supernatant and the results obtained with the upper 1 ml of the sediment after staining with 10.0 µg/ml of specific antibody at 25°C overnight in the dark were compared.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
In recent decades, parasitic protozoa have been found to have a great potential to cause waterborne and food-borne diseases (7). Outbreaks with such organisms have enormous economic and social consequences, particularly in terms of health care costs and a lack of confidence in the quality of water systems and sanitation standards (11). Although Cryptosporidium sp. and Giardia lamblia are the major protozoan pathogens responsible for waterborne outbreaks in industrialized countries (11), microsporidia are considered emerging opportunistic parasites (16). The ubiquitous presence of such parasites in aquatic ecosystems and their invariable resistance to most of the conventional water treatment procedures make the development of preventive strategies mandatory in order to ensure the safety of the public water supply. One waterborne outbreak due to microsporidia reported during the summer of 1985 in France (24) most likely resulted from contamination and deficiencies in surface water treatment, which was done by flocculation, ozoflotation, and filtration instead of chlorination (4).

The identification of the microsporidial species responsible for an infection is very important, namely, to define the appropriate treatment, as well as to ensure its eradication from the environment. While E. intestinalis infections are treated with albendazole (1), no treatment is yet available for other species (23) except Enterocytozoon bieuneusi (1), the treatment of which requires very specific drugs, like fumagillin.

The diagnosis of microsporidiosis in a routine laboratory can theoretically be performed by direct visualization of spores in clinical specimens by light or fluorescence microscopy (26, 27); nevertheless, it is tricky to differentiate the parasite from the other elements, like debris, usually present in biological samples (25). Diagnosis based upon PCR assays is emerging. Apart from being expensive, PCR presents several additional disadvantages, particularly because it does not allow the easy quantification or determination of the viability of microorganisms. It also requires a considerable amount of time and equipment, as well as expertise (15, 25). The development of immunofluorescent reagents (specific monoclonal or polyclonal antibodies directed against microsporidial spores) has contributed to improvements in the diagnosis of microsporidial infections. FC, a method based upon the evaluation of cell fluorescence, has repeatedly been shown to be specific; to have a high level of sensitivity, which is 10-fold greater than that of microscopy (5); and to not be dependent on technician expertise for the evaluation of samples, as is the case in immunofluorescence microscopy. In addition, it provides an objective means of analysis that allows the rapid assessment of clinical specimens for the presence of opportunistic microsporidial organisms. Accompanying the increasing complexity of diagnostic laboratory techniques, high costs, and the need for human expertise, the use of more sensitive and automated methods, especially for biological samples like stool samples, has been recommended (9, 25).

Clinical samples usually contain many different microorganisms, like parasites, fungi, and bacteria. Staining could differentiate microsporidial spores from the debris normally present in biological samples, especially if a specific monoclonal or polyclonal antibody is used in combination with a fluorochrome reagent. However, in fecal samples, cross-reactions with other distinct microorganisms could occur, preventing the use of polyclonal antibodies in this setting (25). In order to confirm the specificity of the monoclonal antibody used in this study, the possibility of cross-reactions with other organisms was investigated by using both prokaryotic microorganisms (Escherichia coli, Staphylococcus aureus) and eukaryotic microorganisms (Candida albicans, Cryptosporidium parvum, Giardia lamblia) mixed with an E. intestinalis spore solution. The bacteria, fungi, and other parasites present in the mixed suspensions did not interfere with the detection of the microsporidia. We verified that bacteria showed up in the autofluorescence zone but did not stain with the microsporidium-specific antibody, which was certainly because their size is similar to that of microsporidial spores. It would be very interesting to evaluate whether cross-reactions with Enterocytozoon bieneusi, a human microsporidial species prevalent worldwide, would occur, but E. bieneusi microorganisms are not commercially available. Also, and according to the manufacturer, the antibody used in this study does not stain E. bieneusi because it is specific to the genus Encephalitozoon. Specificity is a very relevant topic for any diagnostic procedure. Our results support the use of the protocol described here with clinical samples, as it provides a reliable means of detection of E. intestinalis spores and clearly separates E. intestinalis spores from debris and other microorganisms. The conventional microscopic diagnostic procedure is highly dependent on the experience of the microscopist and the spore concentration in the sample. The protocol described here provides a detection limit (10⁴ spores/ml) well bellow the concentration usually detected by conventional procedures (5). Additionally, the simultaneous staining with the specific antibody and PI allowed the clear distinction of dead parasites among the debris and the viable microorganisms that could be present in clinical samples. This is of extreme relevance since it allows the future study of the infectious potential of samples, as this parasite is able to resist common water treatments (9, 10).

The use of saturated NaCl solution improved the detection of the parasite comparing with the use of other solutions. This might result from the fact that spores can easily float, in contrast to other protozoan parasites like Giardia lamblia and Cryptosporidium parvum cysts and oocysts. An improvement in parasite recovery over that in our previous studies (2, 3) was also found by the use of a higher centrifugation velocity.

A new cytometric detection method is now available for the detection of E. intestinalis spores in water and stool samples, as well as for assessment of their viability based upon dye (PI) exclusion. The use of a specific antibody allows the clear discrimination between E. intestinalis spores and the debris or other microorganisms often present in water and human stool samples. We now intend to apply this protocol to the routine analysis of clinical samples, especially those from immunocompromised patients, as well as to the evaluation of environmental water samples.

 
* Corresponding author. Mailing address: Department of Microbiology, Porto Faculty of Medicine, University of Porto, Al. Prof. Hernani Monteiro, Porto 4200-319, Portugal. Phone and fax: 351 22 551 3662. E-mail: gui75{at}sapo.pt

Published ahead of print on 13 May 2009.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Alfa Cisse, O., A. Ouattara, M. Thellier, I. Accoceberry, S. Biligui, D. Minta, O. Doumbo, I. Desportes-Livage, M. A. Thera, M. Danis, and A. Datry. 2002. Evaluation of an immunofluorescent-antibody test using monoclonal antibodies directed against Enterocytozoon bieneusi and Encephalitozoon intestinalis for diagnosis of intestinal microsporidiosis in Bamako (Mali). J. Clin. Microbiol. 40:1715-1718.[Abstract/Free Full Text]
2
2. Barbosa, J., S. Costa-de-Oliveira, A. G. Rodrigues, and C. Pina-Vaz. 2008. Optimization of a flow cytometry protocol for detection and viability assessment of Giardia lamblia. Travel Med. Infect. Dis. 6:234-239.[CrossRef][Medline]
3
3. Barbosa, J. M., S. Costa-de-Oliveira, A. G. Rodrigues, T. Hanscheid, H. Shapiro, and C. Pina-Vaz. 2008. A flow cytometric protocol for detection of Cryptosporidium spp. Cytometry A 73:44-47.[Medline]
4
4. Cotte, L., M. Rabodonirina, F. Chapuis, F. Bailly, F. Bissuel, C. Raynal, P. Gelas, F. Persat, M. A. Piens, and C. Trepo. 1999. Waterborne outbreak of intestinal microsporidiosis in persons with and without immunodeficiency virus infection. J. Infect. Dis. 180:2003-2008.[CrossRef][Medline]
5
5. Delaunay, A., G. Gargala, X. Li, L. Favennec, and J. J. Ballet. 2000. Quantitative flow cytometric evaluation of maximal Cryptosporidium parvum oocyst infectivity in a neonate mouse model. Appl. Environ. Microbiol. 66:4315-4317.[Abstract/Free Full Text]
6
6. Didier, E. S., and L. M. Weiss. 2006. Microsporidiosis: current status. Curr. Opin. Infect. Dis. 19:485-492.[Medline]
7
7. Dowd, S. E., D. John, J. Eliopolus, C. P. Gerba, J. Naranjo, R. Klein, B. López, M. de Mejía, C. E. Mendoza, and I. L. Pepper. 2003. Confirmed detection of Cyclospora cayetanesis, Encephalitozoon intestinalis and Cryptosporidium parvum in water used for drinking. J. Water Health 1:117-123.[Medline]
8
8. Franzen, C., A. Muller, P. Hartmann, and B. Salzberger. 2004. Quantitation of microsporidia in cultured cells by flow cytometry. Cytometry A 60:107-114.[Medline]
9
9. Garcia, L. S. 2002. Laboratory identification of the microsporidia. J. Clin. Microbiol. 40:1892-1901.[Free Full Text]
10
10. Hoffman, R. M., M. M. Marshall, D. M. Polchert, and B. H. Jost. 2003. Identification and characterization of two subpopulations of Encephalitozoon intestinalis. Appl. Environ. Microbiol. 69:4966-4970.[Abstract/Free Full Text]
11
11. Karanis, P., C. Kourenti, and H. Smith. 2007. Waterborne transmission of protozoan parasites: a worldwide review of outbreaks and lessons learnt. J. Water Health 5:1-38.[CrossRef][Medline]
12
12. Keeling, P., and N. M. Fast. 2002. Microsporidia: biology and evolution of highly reduced intracellular parasites. Annu. Rev. Microbiol. 56:93-116.[CrossRef][Medline]
13
13. Lobo, M. L., A. Teles, M. Barão da Cunha, J. Henriques, A. M. Lourenço, F. Antunes, and O. Matos. 2003. Microsporidia detection in stools from pets and animals from the zoo in Portugal: a preliminary study. J. Eukaryot. Microbiol. 50:581-582.[CrossRef][Medline]
14
14. Matos, O., M. L. Lobo, L. Gonçalves, and F. Antunes. 2002. Diagnostic use of 3 techniques for identification of microsporidian spores among AIDS patients in Portugal. Scand. J. Infect. Dis. 34:591-593.[CrossRef][Medline]
15
15. Menotti, J., B. Cassinat, C. Sarfati, O. Liguory, F. Derouin, and J. M. Molina. 2003. Development of a real-time PCR assay for quantitative detection of Encephalitozoon intestinalis DNA. J. Clin. Microbiol. 41:1410-1413.[Abstract/Free Full Text]
16
16. Moss, D. M., G. P. Croppo, S. Wallace, and G. S. Visvesvara. 1999. Flow cytometric analysis of microsporidia belonging to the genus Encephalitozoon. J. Clin. Microbiol. 37:371-375.[Abstract/Free Full Text]
17
17. Pina-Vaz, C., S. Costa-Oliveira, and A. G. Rodrigues. 2004. Safe susceptibility testing of Mycobacterium tuberculosis by flow cytometry with the fluorescent nucleic acid stain SYTO 16. J. Med. Microbiol. 53:77-81.
18
18. Pina-Vaz, C., S. Costa-Oliveira, A. G. Rodrigues, and A. Espinel-Ingroff. 2005. Comparison of two probes for testing susceptibilities of pathogenic yeasts to voriconazole, itraconazole, and caspofungin by flow cytometry. J. Clin. Microbiol. 43:4674-4679.[Abstract/Free Full Text]
19
19. Pina-Vaz, C., S. Costa-Oliveira, A. G. Rodrigues, and A. Salvador. 2004. Novel method using a laser scanning cytometer for detection of Mycobacteria in clinical samples. J. Clin. Microbiol. 42:906-908.[Abstract/Free Full Text]
20
20. Pina-Vaz, C., A. Rodrigues, F. Sansonetty, J. Martinez-de-Oliveira, A. P. Fonseca, and P. A. Mardh. 2000. Antifungal activity of local anesthetics against Candida species. Infect. Dis. Obstet. Gynecol. 8:124-137.[CrossRef][Medline]
21
21. Pina-Vaz, C., F. Sansonetty, A. Rodrigues, J. Martinez-de-Oliveira, A. P. Fonseca, and P. A. Mardh. 2000. Antifungal activity of ibuprofen alone and in combination with fluconazole against Candida species. J. Med. Microbiol. 49:831-840.[Abstract/Free Full Text]
22
22. Pina-Vaz, C., F. Sansonetty, A. G. Rodrigues, S. Costa-de-Oliveira, C. Tavares, and J. Martinez-de-Oliveira. 2001. Cytometric approach for a rapid evaluation of susceptibility of Candida strains to antifungals. Clin. Microbiol. Infect. 7:609-618.[CrossRef][Medline]
23
23. Santillana-Hayat, M., C. Sarfati, S. Fournier, F. Chau, R. Porcher, J. M. Molina, and F. Derouin. 2002. Effects of chemical and physical agents on viability and infectivity of Encephalitozoon intestinalis determined by cell culture and flow cytometry. Antimicrob. Agents Chemother. 46:2049-2051.[Abstract/Free Full Text]
24
24. Stine, S. W., F. D. Vladich, I. L. Pepper, and C. Gerba. 2005. Development of a method for the concentration and recovery of Microsporidia from tap water. J. Environ. Sci. Health Part A 40:913-925.
25
25. Thellier, M., and J. Breton. 2008. Enterocytozoon bieneusi in human and animals focus on laboratory identification and molecular epidemiology. Parasite 15:349-358.[Medline]
26
26. Weber, R., R. T. Bryan, R. L. Owen, C. M. Wilcox, L. Gorelkin, and G. S. Visvesvara. 1992. Improved light-microscopical detection of microsporidia spores in stool and duodenal aspirates. N. Engl. J. Med. 326:161-166.[Abstract]
27
27. Weber, R., R. T. Bryan, D. A. Schwartz, and R. Owen. 1994. Human microsporidial infections. Clin. Microbiol. Rev. 7:426-461.[Abstract/Free Full Text]

---

Clinical and Vaccine Immunology, July 2009, p. 1021-1024, Vol. 16, No. 7
1071-412X/09/$08.00+0     doi:10.1128/CVI.00031-09
Copyright © 2009, 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 Citation Map Services E-mail this article to a friend 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 Barbosa, J. Articles by Pina-Vaz, C. Search for Related Content PubMed PubMed Citation Articles by Barbosa, J. Articles by Pina-Vaz, C.