Previous Article | Next Article 
Journal of Clinical Microbiology, November 1999, p. 3465-3468, Vol. 37, No. 11
0095-1137/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Value of PCR for Detection of Toxoplasma
gondii in Aqueous Humor and Blood Samples from Immunocompetent
Patients with Ocular Toxoplasmosis
Germán
Bou,1,*
Marta S.
Figueroa,2
Paloma
Martí-Belda,1
Enrique
Navas,1 and
Antonio
Guerrero3
Servicio de
Microbiología1 and Servicio de
Oftalmología,2 Hospital Ramón y
Cajal, 28034 Madrid, and Servicio de Microbiología,
Complejo Hospitalario Juan Canalejo, 15006 La
Coruña,3 Spain
Received 4 March 1999/Returned for modification 4 June
1999/Accepted 13 July 1999
 |
ABSTRACT |
Toxoplasma gondii infection is an important cause of
chorioretinitis in the United States and Europe. Most cases of
Toxoplasma chorioretinitis result from congenital
infection. Patients are often asymptomatic during life, with a peak
incidence of symptomatic illness in the second and third decades of
life. Diagnosis is mainly supported by ophthalmological examination and
a good response to installed therapy. However, establishment of a
diagnosis by ophthalmological examination alone can be difficult in
some cases. To determine the diagnostic value of PCR for the detection
of T. gondii, 56 blood and 56 aqueous humor samples from 56 immunocompetent patients were examined. Fifteen patients with a
diagnosis of ocular toxoplasmosis had increased serum anti-T.
gondii immunoglobulin G levels but were negative for
anti-T. gondii immunoglobulin M (group 1), and 41 patients
were used as controls (group 2). Samples were taken before
antiparasitic therapy was initiated, and only one blood sample and one
aqueous humor sample were obtained for each patient. Single nested PCRs
and Southern blot hybridization were performed with DNA extracted from
these samples. The results obtained showed sensitivity and specificity
values of 53.3 and 83%, respectively. Interestingly, among all
patients with ocular toxoplasmosis, a positive PCR result with the
aqueous humor sample was accompanied by a positive PCR result with the
blood sample. This result suggests that ocular toxoplasmosis should not
be considered a local event, as PCR testing of blood samples from
patients with ocular toxoplasmosis yielded the same result as PCR
testing of aqueous humor samples. PCR testing may be useful for
discriminating between ocular toxoplasmosis and other ocular diseases,
and also can avoid the problems associated with ocular puncture.
| |
INTRODUCTION |
Toxoplasma gondii
infection is an important cause of chorioretinitis in the United States
and Europe. Most cases of Toxoplasma chorioretinitis result
from congenital infection (16). Patients are often
asymptomatic, with a peak incidence of symptomatic disease in the
second and third decades of life. The characteristic lesion is a focal
necrotizing retinitis that initially appears in the fundus as a
yellowish white, elevated cotton patch with indistinct margins, usually
on the posterior pole (15). The clinical ophthalmological findings together with positive anti-T. gondii serology
provide sufficient information on which to base a diagnosis; the latter would be confirmed by a good response to installed therapy. However, in
immunocompromised patients the clinical findings are sometimes not
clear enough to make a definitive diagnosis, raising the question of
which therapeutic strategy should be used. A wrong decision not only
could cause a delay in adequate treatment and the preventable loss of a
functioning retina for the infected patient but also could expose the
uninfected patient to the toxic side effects of unnecessary medication.
Regarding the laboratory diagnosis of ocular toxoplasmosis, cell
culture of intraocular fluids is particularly insensitive when only a
small amount of material is available; furthermore, it may take days to
weeks to obtain a result. Detection of locally produced antibodies may
be useful for immunocompetent patients but is probably not useful for
immunocompromised patients. PCR detects the DNAs of microorganisms and
is a rapid method which has been used to detect T. gondii
DNA in different biological samples (4, 9, 12, 14, 19). Most
of these samples can be obtained only by invasive procedures, and only blood can easily be obtained from the patients. This technique has also
been used with immunocompetent and immunosuppressed patients with
sight-threatening retinitis; so far, the results that have been
obtained are inconsistent, and there are discrepancies in the
sensitivity values (1, 2, 10, 18, 21). Therefore, we
examined the diagnostic value of PCR for the detection of T. gondii in blood and aqueous humor samples from a large group of patients with and without ocular toxoplasmosis.
| |
MATERIALS AND METHODS |
Patients.
Fifty-six blood and 56 aqueous humor samples from
56 patients were examined. Serum anti-T. gondii
immunoglobulin G (IgG) and IgM were detected by enzyme-linked
immunosorbent assay. Patients were divided into two groups. (i) Group 1 consisted of 15 patients with a clinical diagnosis of ocular
toxoplasmosis reactivation. The mean age of the patients was 35.5 years
(age range, 18 to 59 years). Anterior chamber taps were taken after
topical application of oxybuprocaine chloride (0.4%) and tetracaine
chloride (0.1%). The eye was gently grasped with a fixating forceps at
the nasal side, and the anterior chamber was entered just anterior of
the limbus from the temporal side with a 30-gauge needle on a 1-ml syringe. After the collection of 0.2 to 0.3 ml of aqueous humor, the
needle was removed. Patients were examined after 1 h. No
complications were seen. Samples were taken from these patients before
specific antiparasitic therapy was started. No problems were associated with the puncture. All these patients had ocular symptoms, and indirect
ophthalmoscopy of a dilated eye showed the typical active lesions of
T. gondii retinitis adjacent to pigmented scars (the typical
appearance of congenital ocular toxoplasmosis reactivation). Antitoxoplasmic drugs (oral pyrimethamine, 50 mg daily for 2 days and
then 25 mg daily, plus oral folinic acid, 10 to 20 mg daily, and either
1 or 1.5 g of sulfadiazine four times daily) were administered for 6 to
8 weeks to patients with clinically diagnosed or suspected T. gondii retinitis. Ophthalmoscopic examination and the response to
antitoxoplasmic therapy (resolution of the retinal lesion) suggested
ocular toxoplasmosis. After antitoxoplasmic therapy, symptoms improved
and ocular lesions resolved, leaving pigmented scars. (ii) Group 2 consisted of 41 patients with no clinical evidence of ocular
toxoplasmosis. The mean age of the patients was 58.1 years (age range,
23 to 80 years). These patients had other ocular diseases, and aqueous
humor and blood samples were obtained before vitreoretinal surgery. For
all patients, only one blood sample and one aqueous humor sample were examined.
Serology.
Serologic determinations were performed with the
kit VIDAS Toxo IgG and IgM (Bio Mérieux, Paris, France), and the
manufacturer's directions were followed.
DNA preparation.
DNA was prepared by using a kit (DNA III
Extraction; Real, Valencia, Spain) in accordance with the
manufacturer's directions. For blood and aqueous humor samples, 5 ml
of anticoagulated blood and a 200- to 300-µl portion of aqueous
humor, respectively, were used for DNA extraction.
Amplification procedures.
The amplification reactions were
performed with 50-µl reaction mixtures containing the following: 100 pmol of each primer, 20 mM Tris-HCl (pH 8.3), 50 mM KCl, 2.5 mM
MgCl2, 200 µM concentrations of each deoxynucleoside
triphosphate, and 2.5 U of Taq DNA polymerase (Perkin-Elmer,
Norwalk, Conn.). A total of 2.0 µg of genomic DNA was used for blood
samples, and an undetectable amount of DNA was used when PCR was done
with aqueous humor samples. Samples were overlaid with 60 µl of
paraffin oil, and the reactions were run in a Perkin-Elmer
thermocycler. After initial denaturation of the DNA at 94°C for 4 min, 55 cycles were run, as follows: 94°C for 1 min, 42°C for
30 s, and 72°C for 2 min. The final extension step continued for
an additional 10 min at 72°C. For the nested PCR, the amplification
reactions were performed in 50-µl reaction mixtures containing the
following: 30 pmol of each primer, 100 µM concentrations of each
deoxynucleoside triphosphate, 20 mM Tris-HCl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.02% gelatin, and 2.5 U of Taq DNA
polymerase (Perkin-Elmer). A total of 20 µl of a 1:200 dilution of
the first PCR product was used for the second amplification. Samples
were overlaid with 60 µl of paraffin oil, and reactions were run in a
Perkin-Elmer thermocycler. After initial denaturation of the DNA at
94°C for 4 min, 17 cycles were run, as follows: 94°C for 1 min
20 s, 53°C for 2 min, and 72°C for 2 min and 30 s. The
final extension step was continued for an additional 5 min at 72°C.
Primer pair P1 (5'-TGCATAGGTTGCAGTCACTG-3') and P3
(5'-TCTTTAAAGCGTTCGTGGTC-3') (Cruachen, Progenetic, S.L.,
Madrid, Spain), which amplifies a 133-bp DNA fragment of the repetitive
B1 gene of T. gondii (3), was used for the first
PCR. Primer pair P1 and P2 (5'-GGCGACCAATCTGCGAATACAC-3'),
which amplifies an internal 97-bp fragment of the first amplicon,
was used for the nested PCR. After amplification, an aliquot of 25 µl
from each reaction mixture was run on a 3.0% electrophoresis-grade
agarose gel in 1× TBE buffer (0.09 M Tris-borate, 0.002 M EDTA), and
DNA was stained with ethidium bromide (50 µg/ml). Bands were
visualized under UV illumination.
A Southern blot hybridization (20) was performed with all
the samples after the first PCR. After gel electrophoresis the amplified DNA samples were denatured (0.5 N NaOH, 1.5 M NaCl) for 30 min, renatured (1.0 M Tris-HCl [pH 7.5], 1.5 M NaCl) for 30 min, and
then transferred to a positively charged nylon membrane (Boehringer
Mannheim, Mannheim, Germany) by capillary transfer with 20× SSC buffer
(3 M NaCl, 0.3 M sodium citrate [pH 7.0]). DNA was coupled to the
membrane by using a UV chamber (Bio-Rad, Madrid, Spain) at 150 mJ. The
membrane was washed with 2× SSC and prehybridized at 68°C for 1 h with hybridization solution (Boehringer Mannheim), and afterward, 5 pmol of the 133-bp amplicon that was 3' labeled with digoxigenin-dUTP
with terminal transferase was added to the mixture solution and the
mixture was hybridized at 68°C overnight. After a washing step (with
2× SSC-0.1% sodium dodecyl sulfate and 0.1× SSC-0.1% sodium
dodecyl sulfate for 1 h each) at 68°C, the probe was detected by
a chemiluminescence method (Boehringer Mannheim).
A clinical sample was considered to be
T. gondii DNA
positive (PCR positive) if the PCR test (single and/or nested PCR) was
positive and the 133-bp amplified fragment of the first PCR hybridized
with the probe in the Southern blotting experiment. A patient
was
considered to be
T. gondii DNA positive (PCR positive) if
a
positive amplification of one of the samples (blood or aqueous
humor)
was
obtained.
The DNA extraction, amplification, and detection steps were physically
separated and were performed in three different rooms.
Positive
controls and negative controls (reaction mixtures without
DNA) were
used in all experiments. The amplification experiment
was considered to
be invalid when a failure in any of the controls
occurred.
Virulent strain
T. gondii RH was grown in monolayers of
MRC-5 cells and was used as a positive control. When a cytopathic
effect was observed in the culture (flask of 7.5 by 3.5 cm), cells
and
tachyzoites were collected and DNA was obtained and resuspended
in 100 µl of distilled water. After the amplification reaction
with primers
P1 and P3 and further nested PCR under the conditions
described above,
the fragment of the correct size was detected
up to a dilution of
1:10
6 with water. DNA extracted at a dilution of
1:10
5 was used as a positive control in all amplification
experiments.
A correct DNA amplification yielded a band of 133 bp in
the first
amplification and a band of 97 bp in the nested PCR (Fig.
1).
View larger version (50K):
[in this window]
[in a new window]
|
FIG. 1.
T. gondii PCR products resolved on a 3%
agarose gel with 1× TBE buffer. Lane 1, DNA molecular size marker
( V; Boehringer Mannheim); lane 2, product of the first
amplification; lane 3, product of the nested PCR.
|
|
| |
RESULTS |
The PCR was positive for 8 of the 15 blood samples and 7 of the 15 aqueous humor samples from patients in group 1. No amplification was
observed with blood and aqueous humor samples from the other seven
patients in group 1. Thus, a correspondence of positive results by PCR
between blood and aqueous humor samples was observed for seven
patients; for one patient with ocular toxoplasmosis, only the blood
sample was positive. In addition, all these patients had higher serum
anti-T. gondii IgG levels but were negative for anti-T. gondii IgM. With DNA from blood and aqueous humor
samples from patients in group 2, no amplification of DNA fragments
from blood was observed for 35 patients, whereas a positive PCR result was obtained for 7 patients. For these seven patients, we also observed
a correspondence of positive PCR results between blood and aqueous
humor samples for all except one patient, for whom only the aqueous
humor sample was positive by DNA amplification. Twenty-five and 16 of
the 41 patients in group 2 were positive and negative for anti-T.
gondii IgG, respectively. Among the seven PCR-positive patients in
group 2, five were positive for anti-T. gondii IgG, while
the other two were negative for anti-T. gondii IgG. A
summary of the results is shown in Table
1.
View this table:
[in this window]
[in a new window]
|
TABLE 1.
Summary of results of diagnosis of toxoplasmosis by PCR
with 56 blood and 56 aqueous humor samples from 56 patients with and
without ocular toxoplasmosis
|
|
DNAs from all PCR-positive samples hybridized with the B1 probe. None
of the DNA samples that were PCR negative hybridized with the B1 probe.
With these results we conclude that a very good correlation exists
between the results of the nested PCR and those of the Southern blot assay.
| |
DISCUSSION |
We report here the results of our experience with the use of PCR
for the identification of T. gondii in blood and aqueous humor samples from patients with and without ocular toxoplasmosis. Currently, the clinical diagnosis of ocular toxoplasmosis is based on
the observation of a necrotizing lesion on the fundus, response to
treatment, and serologic determination. However, in cases of atypical
retinitis or when the fundus is hidden by vitreal inflammation, establishment of a diagnosis by ophthalmological examination alone can
be difficult. In such cases the aqueous humor analysis may be used as a
diagnostic tool for confirmation of ocular toxoplasmosis. Determination
of intraocular production of antitoxoplasmic antibodies has been
performed previously, but the results obtained had erratic values
(2, 21). PCR has mostly been used to detect T. gondii in different biological samples (4, 9, 12, 14,
19). Aqueous humor samples from patients with ocular
toxoplasmosis have also been used with the PCR technique (1, 2,
10, 18, 21). All these studies were carried out by examining
aqueous humor samples exclusively. Sensitivities ranged from 2.3 to
75%, but specificities ranged from 75 to 100% (2, 10, 18,
21). In our opinion, these studies had several important
limitations; first, most of them were retrospective studies; second,
the final diagnosis was made on the basis of treatment response but was not microbiologically confirmed; and third, very small numbers of
samples were tested in the studies. We report here sensitivity and
specificity values of 53 and 83%, respectively, when we consider the
results for both blood and aqueous humor samples and a sensitivity value of 46.6% when we consider only the results for aqueous humor samples among a collection of 112 samples from 56 patients. The low
sensitivity values in this work are similar to those reported previously (2, 10, 18, 21). The distance between the anterior chamber and the initial chorioretinal site of inflammation must be considered in the interpretation of sensitivity values when a
PCR is performed with an ocular sample (2). On the other hand, the likelihood that the necrotizing lesions are caused by an
immunological mechanism and not by the presence of the parasite cannot
be ruled out (17). Other investigators have reported that
sample storage conditions before the amplification may have an
important influence on the PCR sensitivity values (13). In our study, some of the DNA samples were kept at 
20°C before the amplification was performed because immediate processing was
impossible. This can account for some reduction in the sensitivity
values. On the other hand, despite the very good DNA extraction method, the presence of inhibitors in the aqueous humor samples cannot be ruled
out. The specificity values obtained in our assay are similar to those
obtained in other reports (1, 18, 21). When the sequences of
the P1 and P3 primers are compared with sequences in the GenBank
database, only matches with the T. gondii B1 gene were
observed, thus revealing a theoretical high degree of specificity of
the amplification reactions.
An interesting point to be considered is that blood from patients with
ocular toxoplasmosis yielded positive amplifications with the B1
primers with even higher band intensities than those with the DNAs from
the aqueous humors from the same patients. This result strongly
suggests that ocular toxoplasmosis should not be considered a local
event. Thus, in our study a positive PCR result was obtained with the
blood from 8 of 15 (53.3%) patients with a diagnosis of ocular
toxoplasmosis. Similar findings have been reported by Dupouy-Camet et
al. (6) in the case of cerebral toxoplasmosis in human
immunodeficiency virus-positive patients. Those investigators detected
T. gondii DNA by PCR in the blood of 9 of 13 patients (69%)
with confirmed cerebral toxoplasmosis (6). Moreover, other
studies (5, 7, 8, 11) showed sensitivity values of tests
with blood of between 10 and 35%. An important question is this: Why
was the T. gondii PCR positive with blood from 53% of the
patients with reactivated congenital corioretinitis in our study? There
are two hypotheses: (i) the source of parasitemia or T. gondii DNAemia was the ocular lesion or (ii) ocular toxoplasmosis
reactivation occurs during immunosuppressive states (such as pregnancy)
and is associated with reactivations in other body tissues (muscle,
lung, brain, etc.). Tachyzoites or T. gondii DNA released
from the cysts in these tissues may be the source of parasitemia or
T. gondii DNA in blood. In any case, preventive measures
should be taken for pregnant women after they have had ocular toxoplasmosis.
We have also detected T. gondii DNA in the blood and aqueous
humor of patients without ocular manifestations of toxoplasmosis (6 of
41 blood samples and 7 of 41 aqueous humor samples from control
patients in group 2, and among these patients five were positive for
anti-T. gondii IgG). In order to explain this result, we
consider it possible that a small number of parasites might be released
from tissues into the blood at a subclinical level, and their presence
can be detected by PCR in IgG-positive patients and under some
circumstances (such as low-level immunosuppressive states). We cannot
explain why the aqueous humors from these seven patients had positive
PCR results because the patients had ocular diseases with no
relationship to ocular toxoplasmosis. Moreover, by the PCR technique,
results must be interpreted with caution; although no failures were
observed among the controls, the appearance of a false-positive result
cannot be ruled out.
In summary, although the diagnosis of the ocular reactivation of
toxoplasmosis requires the presence of typical lesions, a positive
result for anti-T. gondii IgG, a negative result for anti-T. gondii IgM, and a response to installed therapy, the
results reported here suggest that the T. gondii PCR is a
useful technique that can be used as a laboratory tool for the
diagnosis of ocular toxoplasmosis. In addition, PCR testing of blood
samples from patients with ocular toxoplasmosis yielded the same
results as PCR testing of aqueous humor samples, thus avoiding all the
problems associated with the ocular puncture. Considering the
sensitivity and specificity values, PCR may be useful for
discriminating between ocular toxoplasmosis and other ocular diseases.
| |
ACKNOWLEDGMENTS |
We thank the National Institute of Health (INSALUD) for economic support.
We thank L. de Rafael for critical reading of the manuscript.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Servicio de
Microbiología, Hospital Ramón y Cajal, Crta. Colmenar
Viejo Km 9,1, 28034 Madrid, Spain. Phone: 34-1-3368082. Fax:
34-1-3368809. E-mail: germanbou{at}mailcity.com.
| |
REFERENCES |
| 1.
|
Aouizerate, F.,
J. Cazenave,
L. Poirier,
P. Verin,
A. Cheyrou,
J. Begueret, and F. Lagoutte.
1993.
Detection of Toxoplasma gondii in aqueous humor by the polymerase chain reaction.
Br. J. Ophthalmol.
77:107-109[Abstract/Free Full Text].
|
| 2.
|
Brezin, A. P.,
C. E. Eqwuagu,
C. Silveria,
P. Thulliez,
M. Martins,
R. Mahdi,
R. Belfort, and R. B. Nusseblatt.
1991.
Analysis of aqueous humor in ocular toxoplasmosis.
N. Engl. J. Med.
10:699.
|
| 3.
|
Burg, J. L.,
C. M. Grover,
P. Pouletty, and J. C. Boothroyd.
1989.
Direct and sensitive detection of a pathogenic protozoan, Toxoplasma gondii, by polymerase chain reaction.
J. Clin. Microbiol.
27:1787-1792[Abstract/Free Full Text].
|
| 4.
|
Cingolani, A.,
A. De Luca,
A. Ammassari,
R. Murri,
A. Linzalone,
R. Grillo, and A. Antinori.
1996.
PCR detection of Toxoplasma gondii DNA in CSF for the differential diagnosis of AIDS-related focal brain lesions.
J. Med. Microbiol.
45:472-476[Abstract/Free Full Text].
|
| 5.
|
Dupon, M.,
J. Cazenave,
J. L. Pellegrin,
J. M. Ragnaud,
A. Cheyrou,
I. Fischer,
B. Leng, and J. Y. Lacut.
1995.
Detection of Toxoplasma gondii by PCR and tissue culture in cerebrospinal fluid and blood of human immunodeficiency virus-seropositive patients.
J. Clin. Microbiol.
33:2421-2426[Abstract].
|
| 6.
|
Dupouy-Camet, J.,
S. L. de Souza,
C. Maslo,
A. Paugam,
A. Saimot,
R. Benarous,
C. Tourte-Schaefer, and F. Derouin.
1993.
Detection of Toxoplasma gondii in venous blood from AIDS patients by polymerase chain reaction.
J. Clin. Microbiol.
31:1866-1869[Abstract/Free Full Text].
|
| 7.
|
Filice, G. A.,
J. A. Hitt,
C. D. Mitchell,
M. Blackstad, and S. W. Sorensen.
1993.
Diagnosis of toxoplasma parasitemia in patients with AIDS by gene detection after amplification with polymerase chain reaction.
J. Clin. Microbiol.
31:2327-2331[Abstract/Free Full Text].
|
| 8.
|
Franzen, C.,
M. Altfeld,
P. Hegener,
P. Hartmann,
G. Arendt,
H. Jablonowski,
J. Rockstroh,
V. Diehl,
B. Salzberger, and G. Fatkenheuer.
1997.
Limited value of PCR for detection of Toxoplasma gondii in blood from human immunodeficiency virus-infected patients.
J. Clin. Microbiol.
35:2639-2641[Abstract].
|
| 9.
|
Fuentes, I.,
M. Rodríguez,
C. J. Domingo,
F. Del Castillo,
T. Juncosa, and J. Alvar.
1996.
Urine sample used for congenital toxoplasmosis diagnosis by PCR.
J. Clin. Microbiol.
34:2368-2371[Abstract].
|
| 10.
|
Garweg, J.,
M. Boehnke, and F. Koerner.
1996.
Restricted applicability of the polymerase chain reaction for the diagnosis of ocular toxoplasmosis.
Ger. J. Ophthalmol.
5:104-108[Medline].
|
| 11.
|
Guy, E. C., and D. H. Joynson.
1995.
Potential of the polymerase chain reaction in the diagnosis of active Toxoplasma infection by detection of parasite in blood.
J. Infect. Dis.
172:319-322[Medline].
|
| 12.
|
Ho-Yen, D. O.,
A. W. L. Joss,
A. H. Balfour,
E. T. Smyth,
D. Baird, and J. M. Chatterton.
1992.
Use of the polymerase chain reaction to detect Toxoplasma gondii in human blood samples.
J. Clin. Pathol.
45:910-913[Abstract/Free Full Text].
|
| 13.
|
James, G. S.,
V. G. Sintchenko,
D. J. Dickenson, and G. L. Gilbert.
1996.
Comparison of cell culture, mouse inoculation, and PCR for detection of Toxoplasma gondii: effects of storage conditions on sensitivity.
J. Clin. Microbiol.
34:1572-1575[Abstract].
|
| 14.
|
Joss, A. W. L.,
J. Chatterton,
R. Evans, and D. O. Yen.
1993.
Toxoplasma polymerase chain reaction on experimental blood samples.
J. Med. Microbiol.
38:38-43[Abstract/Free Full Text].
|
| 15.
|
Mets, M. B.,
E. Holfels,
K. M. Boyer,
C. N. Swisher,
N. Roizen,
L. Stein,
M. Stein,
J. Hopkins,
S. Withers,
D. Mack,
R. Luciano,
D. Patel,
J. S. Remington,
P. Meier, and R. McLeod.
1996.
Eye manifestations of congenital toxoplasmosis.
Am. J. Ophthalmol.
122:309-324[Medline].
|
| 16.
|
Montoya, J. G., and J. S. Remington.
1996.
Toxoplasmic chorioretinitis in the setting of acute acquired toxoplasmosis.
Clin. Infect. Dis.
23:277-282[Medline].
|
| 17.
|
Nusseblatt, R. B.,
K. K. Mittal,
S. Furhman,
S. D. Sharma, and A. G. Palestine.
1989.
Lymphocyte proliferative responses of patients with ocular toxoplasmosis to parasite and retinal antigens.
Am. J. Ophthalmol.
107:632-641[Medline].
|
| 18.
|
Robert, F.,
T. Ouatas,
P. Blanche,
C. Tourte-Schaefer,
D. Sicard, and J. Dupouy-Camet.
1996.
Evaluation rétrospective de la détection de Toxoplasma gondii par réaction de polymérisation en chaîne chez des patients sidéens.
Presse Med.
25:541-545.
|
| 19.
|
Rodrîguez, J. C.,
M. M. Martinez,
A. R. Martínez, and G. Royo.
1997.
Evaluation of different techniques in the diagnosis of Toxoplasma encephalitis.
J. Med. Microbiol.
46:597-601[Abstract/Free Full Text].
|
| 20.
|
Sambrook, J.,
E. F. Fritsch, and T. Maniatis.
1989.
Molecular cloning: a laboratory manual, 2nd ed.
Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y
|
| 21.
|
Verbraak, F. D.,
M. Galema,
G. H. Van den Horn,
M. Bruinenberg,
L. Luyendijk,
S. A. Danner, and A. Kijlstra.
1996.
Serological and polymerase chain reaction-based analysis of aqueous humor samples in patients with AIDS and necrotizing retinitis.
AIDS
10:1091-1099[Medline].
|
Journal of Clinical Microbiology, November 1999, p. 3465-3468, Vol. 37, No. 11
0095-1137/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
This article has been cited by other articles:
-
Switaj, K., Master, A., Borkowski, P. K., Skrzypczak, M., Wojciechowicz, J., Zaborowski, P.
(2006). Association of Ocular Toxoplasmosis with Type I Toxoplasma gondii Strains: Direct Genotyping from Peripheral Blood Samples. J. Clin. Microbiol.
44: 4262-4264
[Abstract]
[Full Text]
-
Balansard, B, Bodaghi, B, Cassoux, N, Fardeau, C, Romand, S, Rozenberg, F, Rao, N A, LeHoang, P
(2005). Necrotising retinopathies simulating acute retinal necrosis syndrome. Br. J. Ophthalmol.
89: 96-101
[Abstract]
[Full Text]
-
Garweg, J. G., Garweg, S.-D. L., Flueckiger, F., Jacquier, P., Boehnke, M.
(2004). Aqueous Humor and Serum Immunoblotting for Immunoglobulin Types G, A, M, and E in Cases of Human Ocular Toxoplasmosis. J. Clin. Microbiol.
42: 4593-4598
[Abstract]
[Full Text]
-
Simon, A., Labalette, P., Ordinaire, I., Frealle, E., Dei-Cas, E., Camus, D., Delhaes, L.
(2004). Use of Fluorescence Resonance Energy Transfer Hybridization Probes To Evaluate Quantitative Real-Time PCR for Diagnosis of Ocular Toxoplasmosis. J. Clin. Microbiol.
42: 3681-3685
[Abstract]
[Full Text]
-
Villard, O., Filisetti, D., Roch-Deries, F., Garweg, J., Flament, J., Candolfi, E.
(2003). Comparison of Enzyme-Linked Immunosorbent Assay, Immunoblotting, and PCR for Diagnosis of Toxoplasmic Chorioretinitis. J. Clin. Microbiol.
41: 3537-3541
[Abstract]
[Full Text]
-
Ramchandani, M, Weaver, J B, Joynson, D H M, Murray, P I
(2002). Acquired ocular toxoplasmosis in pregnancy. Br. J. Ophthalmol.
86: 938-939
[Full Text]
-
Gaudio, P A, Gopinathan, U, Sangwan, V, Hughes, T E
(2002). Polymerase chain reaction based detection of fungi in infected corneas. Br. J. Ophthalmol.
86: 755-760
[Abstract]
[Full Text]
-
Grigg, M. E., Boothroyd, J. C.
(2001). Rapid Identification of Virulent Type I Strains of the Protozoan Pathogen Toxoplasma gondii by PCR-Restriction Fragment Length Polymorphism Analysis at the B1 Gene. J. Clin. Microbiol.
39: 398-400
[Abstract]
[Full Text]
Top
Abstract
Introduction
