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Journal of Clinical Microbiology, April 1998, p. 1117-1121, Vol. 36, No. 4
Rabies Unit,
Received 3 September 1997/Returned for modification 18 December
1997/Accepted 15 January 1998
An optimized reverse transcription (RT)-PCR protocol for the
intravitam detection of rabies virus genomic RNA was tested with clinical samples obtained from 28 patients suspected of having rabies,
9 of whom were confirmed to have had rabies by postmortem examination.
RT-PCR using saliva combined with an immunofluorescence assay performed
with skin biopsy samples allowed detection of rabies in the nine
patients.
According to results of global
surveillance by the World Health Organization, about 50,000 cases of
human rabies occur each year (45), the majority of them in
developing countries (63). Human rabies can be prevented
through the combination of stringent animal vaccination, quarantine
programs, and the availability of expensive vaccines and specific
immunoglobulins (32). In spite of the fact that most
developed countries have good public health measures in place to
prevent human rabies, these countries are dealing with an increasing
number of human rabies cases. This is clearly the case in the United
States, where bat rabies virus variants as well as some imported cases
of human rabies have recently achieved an increased public health
significance (41, 51, 53, 56). This is also the case in
France, where seven human cases were recorded from 1988 to 1997; only
eight cases had been recorded in the preceding 20 years. All of the
human rabies cases in France were imported. A similar situation has
also been observed throughout the whole European Union. Of the 15 cases
reported in the last 10 years, 12 were imported from Asia, Africa, and Latin America (46; unpublished data from France).
These cases underscore the importance of alerting travellers of the
risk of rabies contamination and of the prophylactic methods to prevent the disease (42). They also indicate the importance of
maintaining a good system of surveillance, of keeping medical staffs
well informed on clinical presentation of a disease which rarely occurs in developed countries, and of the necessity to develop diagnostic tools for the identification of rabies in these patients.
The clinical diagnosis of rabies is sometimes suggested by
epidemiological (history of exposure) and clinical (e.g., paresthesia, hydrophobia) findings (36). However, the disease is often
mistaken for other disorders (30). Differentiation from
other neurologic diseases may require extensive investigations.
Therefore, diagnosis is often confirmed late in the course of the
disease or postmortem (31). Delays in diagnosis greatly
increase the number of contacts that require postexposure prophylaxis.
The average number of contacts (hospital personnel, family) receiving
postexposure treatments (PET) is approximately 50 per case
(n = 19) in France and between 41 and 55 per case in
the United States (29, 35). In the United States, one case
resulted in 209 PET (50), and 290 PET for one case were
reported recently in France. The early diagnosis of rabies is also
essential to eliminate the expense and discomfort of unnecessary
diagnostic tests and inappropriate therapy.
A wide variety of viruses, bacteria, and parasites, all of which are
capable of causing aseptic meningitis and encephalitis, have been
detected by PCR (39, 49, 52, 58). The objective of the
present study was to establish a reverse transcription (RT)-PCR
protocol for use in evaluating diagnostic specimens, including saliva
and cerebrospinal fluid (CSF). CSF samples were centrifuged at
11,000 × g for 20 min at 4°C. Total RNA was
extracted from specimens of saliva and pellets of CSF by four different techniques, including the following: (i) proteinase K
(34), (ii) guanidinium thiocyanate together with silica
particles (5, 6), (iii) cationic surfactant (Catrimox-14;
Iowa Biotechnology Corporation) (43), and (iv) chelating
resin (Chelex 100; Bio-Rad) (60). The proteinase K method
was performed as follows. Briefly, 200 µl of biological fluid
(saliva or CSF including the pellet) was incubated for 2 h at
37°C with 400 µl of proteinase K buffer containing 40 µg of
proteinase K (Gibco BRL). The RNA was then purified by a
phenol-chloroform extraction and precipitated in absolute ethanol. The
pellets were resuspended in 100 µl of pyrolyzed water. Considering
that the N gene is the most conserved in the lyssa viruses (except some
domains of the L protein gene) and that the sequence data concerning
this gene are the most exhaustive, we used primers in the N gene that
were shown to allow amplification of a wide range of genetically
diverse lyssa viruses (7, 40). One microliter of primer N12
(5' GTAACACCTCTACAATGG 3', positions 57 to 74; all the
positions of the primers are given based on the PV strain sequence
[59]) (100 ng/µl) was incubated with 2 µl of RNA
(1 µg) at 65°C for 3 min and chilled on ice. Each tube was then
incubated at 37°C for 90 min with 4 µl of a solution containing
each nucleotide triphosphate (10 mM), 0.6 µl of RNasin (15 U), 1 µl
of dithiothreitol, 2 µl of Super Script reverse transcriptase buffer
(Gibco BRL), and 0.5 µl of Super Script reverse transcriptase (100 U;
Gibco BRL). The RNA-cDNA hybrids were diluted 10 times in Tris-EDTA
buffer. Five microliters of diluted cDNA was mixed with 50 µl
containing a 1 µM concentration of N12 and N40 (5' GCTTGATGATTGGAACTG 3', positions 1368 to 1349), 20 mM each
nucleotide triphosphate, 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 3 mM
MgCl2, and 2 U of Taq polymerase (Gibco BRL).
PCR was performed with a GeneAmp PCR System 9600 (Perkin-Elmer) by
using the following program: 1 cycle of denaturation at 94°C for
60 s, annealing at 50°C for 90 s, and elongation at 72°C
for 90 s; 30 cycles of denaturation for 50 s; and a final
cycle of elongation for 5 min. The optimum temperature for annealing
and magnesium ion concentration for the PCR was determined by standard
titration experiments of a cDNA of the N gene of strain 9147FRA. Five
microliters of PCR products was diluted into 95 µl of pyrolyzed
water, denatured for 3 min at 95°C, and chilled on ice. The samples
were then distributed onto a Hybond nylon transfer membrane (Amersham)
with a Bio-Dot microfiltration unit (Bio-Rad) and fixed by irradiation
on a UV transilluminator. Primer set N3 (5' GTCTCTTTGAAGCCTGAG 3',
positions 113 to 130)-N23 (5' GGTCTCTCGTCAGTTCCAT 3',
positions 464 to 446) and primer set N17 (5'
TTCTTCCACAAGAACTTTG 3', positions 848 to 866)-N2 (5'
CCCATATAGCATCCTAC 3', positions 1030 to 1013) were used to
generate two digoxigenin-labelled probes by PCR. Hybridization with the
two complementary digoxigenin-labelled probes and detection by
chemiluminescence were performed by using a nonradioactive DNA
labelling and detection kit (Boehringer).
The four different methods of RNA extraction were tested in parallel,
each on two independent series of experimentally infected samples, to
determine the threshold of detection of the RT-PCR. Briefly, negative
specimens of saliva and CSF were pooled and infected with serial
dilutions of a fox rabies isolate (isolate 9147FRA) (40).
This virus was produced on BHK-21 cells infected at a multiplicity of
infection of 0.1 and harvested 48 h after infection to limit the
number of defective viral particles. The Catrimox-14 and Chelex 100 techniques produced negative results. In our hands, poor results were
achieved when guanidinium thiocyanate was used together with silica
particles (threshold of detection, around 104 focus-forming
units [FFU]/ml). However, the results obtained with the proteinase K
method were reproducible with a threshold of detection of
102 FFU/ml in the saliva and 10 FFU/ml in the CSF.
These results are similar to the theoretical viral infectivity obtained
by rapid tissue culture infection test (RTCIT) (20 FFU/ml)
(9). The difference in the results for saliva and CSF can be
explained by the presence of RNase in the saliva.
Forty-one specimens of CSF and 44 specimens of saliva collected daily
by aspiration or by use of cotton swabs were obtained from 28 patients
with manifestations of aseptic meningitis, encephalitis, and suspected
rabies infection (Table 1). Rabies was
confirmed in the brain tissue of nine of these patients by postmortem
examination (8, 28). One patient each was infected in the
Middle East, Latin America (38), and India, and six patients
were infected in Africa (18). One of these patients (patient
P13) was known to be infected with human immunodeficiency virus
(1). The day of onset of illness was defined as the day when
the first symptom attributed to rabies could be noted. These data are
missing for two patients. The average duration of illness was 12.4 days
for the seven remaining rabies-infected patients. The specificity of
RT-PCR was one for the samples of saliva and CSF. Eleven of the 37 putative positive specimens of saliva (sensitivity, 0.30) and only 2 of
22 specimens of CSF (sensitivity, 0.09) were confirmed positive (Table
2). The bronchoalveolar washing was
negative. Intravitam diagnosis performed by RT-PCR identified the
presence of rabies virus in the saliva of five patients (sensitivity
for the patient, 0.56) and in the CSF of only two patients (sensitivity for the patient, 0.22). RT-PCR performed with saliva samples is of particular help in early diagnosis of the disease (Table 2).
Patient P7, who experienced a long clinical period, was confirmed
positive for rabies at the time of death. Specimens of saliva from
patient P7 were confirmed positive consecutively on days 21 and 22 of
the disease, became negative on day 23, and became positive again on
day 24. To investigate the relationship between the intermittence of
rabies virus excretion and the time of day of sample collection, 11 specimens of saliva were collected every 2 or 4 h from one patient
(patient P28) during a 5-day period midway through the course of the
disease. This period starts 2 days after the last collection of saliva
which was found positive, 9 days after the onset of the symptoms, and
12 days before death. None of these specimens were confirmed positive.
The question of titer and duration of virus shedding is of particular
importance for rabies diagnosis because saliva remains the major source
of exposure in contacts for whom rabies prophylaxis is recommended (35). The signal obtained after RT-PCR using some saliva
samples indicated that the level of rabies virus in the saliva could
reach more than 103 FFU/ml (data not shown). This evidence
confirms that any contact with saliva from a patient suspected of
having rabies should be taken seriously.
Most of the conventional techniques used for postmortem analysis of the
brain are of limited value to support the intravitam diagnosis of
rabies (9, 37, 61). We reviewed records of 39 cases of
intravitam diagnosis of rabies in the United States during the period
1960 to 1996 (3, 10-27) and of 16 cases of intravitam
diagnosis performed in France during the period 1970 to 1997 (Table
3). This confirms that the corneal smear
first developed with mice by Schneider (54) was too
insensitive for accurate clinical diagnosis (3, 4, 44, 62).
The only test that has demonstrated reliable results is the
immunofluorescence (IF) test on skin biopsy samples (62). In
our study, the IF test (33), performed on frozen sections of
the skin biopsy samples, exhibited the highest sensitivity
(sensitivity, 0.86; n = 7). It detected the presence of
rabies virus very early in the course of the disease and could be
considered one of the most important tests for intravitam diagnosis. In
routine laboratory testing, we noticed that the examination of a
minimum of 20 sections was needed to ensure the observation of several
hair follicules. Typical rabies nucleocapsid inclusions can be observed
in the nerve endings around the base of some of these hair follicules
(data not shown). RT-PCR assay of saliva and IF assay of skin sections
together allowed a positive intravitam diagnosis in all of the nine
laboratory-confirmed cases of human rabies presented in this study. The
average delay between the onset of clinical symptoms and the collection
of specimens that confirmed the presence of rabies virus was 6.7 days
(n = 7). The sensitivities of detection of rabies
antibodies by enzyme-linked immunosorbent assay (ELISA) (48)
and by seroneutralization on cell culture (57) were very low
in serum as well as in CSF (results obtained from the sera and CSF of
patients P13 and P28, who had received a PET protocol different from
that recommended by the WHO Expert Committee on Rabies
[64], were excluded) (Table 2). This confirms that
serological testing is of limited value because seroconversion occurs
late in the course of the disease (55) (Table 3). The
detection of rabies antigen in the saliva by ELISA (47) also
gave poor results.
0095-1137/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Intravitam Diagnosis of Human Rabies by PCR Using
Saliva and Cerebrospinal Fluid
ABSTRACT
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TABLE 1.
Patients and specimens analyzed
TABLE 2.
Frequency of intravitam diagnosis of rabies in nine
patients in France (1986-1997)
TABLE 3.
Results of laboratory confirmation of rabies attempted in
55 patients during lifea
On the basis of these results, we propose a simple testing protocol for intravitam diagnosis of rabies. Two different specimens should be sent to the laboratory in the early phase of the disease, a skin biopsy specimen and a saliva specimen. Skin biopsy samples should be analyzed by direct IF and saliva samples should be analyzed by the RT-PCR method described above. Then, after 1 week of illness, serum samples should also be examined for the presence of rabies-specific antibodies. RT-PCR has also been proven to be a valuable adjunct to an epidemiological investigation because sequences can be obtained rapidly from PCR products, thus permitting rapid identification of the virus (2, 40).
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ACKNOWLEDGMENTS |
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We express our gratitude to those who sent us the specimens included in this evaluation, namely, P. Hubert and M. Cloud, Pediatric Intensive Care Unit, Hôpital des Enfants Malades, Paris, France; C. Laffont, Laboratory of Virology, Regional and University Hospital, Nice, France; C. Ricard, Antirabic Centre, Hôpital Bellepierre, Saint Denis, La Réunion, France; P. Hofman, Department of Pathological Anatomy and Cytology, Hôpital Pasteur, Nice, France; P. Dellamonica, Unit of Infectious Diseases, Hôpital de l'Archet, Nice, France; H. Adle-Biassette, B. Godeau, and P. Marcowicz, Hôpital Henri Mondor, Creteil, France; and H. Gallais, Unit of Infectious Diseases, Hôpital de la Conception, Marseille, France. We are grateful to Pascale Cozette and Nicole Vachet for expert technical assistance.
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FOOTNOTES |
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* Corresponding author. Mailing address: National Reference Center for Rabies, Institut Pasteur, 75724 Paris Cedex 15, France. Phone: 01 45 68 87 85. Fax: 01 40 61 30 20. E-mail: hbourhy{at}pasteur.fr.
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Journal of Clinical Microbiology, April 1998, p. 1117-1121, Vol. 36, No. 4
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Copyright © 1998, American Society for Microbiology. All rights reserved.
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[Abstract] [Full Text] -
Hughes, G. J., Smith, J. S., Hanlon, C. A., Rupprecht, C. E.
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