JOURNAL OF CLINICAL MICROBIOLOGY, Jan. 2011, p. 42–47
Copyright © 2011, American Society for Microbiology. All Rights Reserved.
Vol. 49, No. 1
Toward Molecular Parasitologic Diagnosis: Enhanced Diagnostic
Sensitivity for Filarial Infections in Mobile Populations?
Doran L. Fink,1† Gary A. Fahle,2Steven Fischer,2Daniel F. Fedorko,2and Thomas B. Nutman1*
Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases,1and Microbiology Service, Department of
Laboratory Medicine, Warren Grant Magnuson Clinical Center,2National Institutes of Health, Bethesda, Maryland 20892
Received 21 August 2010/Returned for modification 28 September 2010/Accepted 21 October 2010
The diagnosis of filarial infections among individuals residing in areas where the disease is not endemic
requires both strong clinical suspicion and expert training in infrequently practiced parasitological methods.
Recently developed filarial molecular diagnostic assays are highly sensitive and specific but have limited
availability and have not been closely evaluated for clinical use outside populations residing in areas of
endemicity. In this study, we assessed the performance of a panel of real-time PCR assays for the four most
common human filarial pathogens among blood and tissue samples collected from a cohort of patients
undergoing evaluation for suspected filarial infections. Compared to blood filtration, real-time PCR was
equally sensitive for the detection of microfilaremia due to Wuchereria bancrofti (2 of 46 samples positive by
both blood filtration and PCR with no discordant results) and Loa loa (24 of 208 samples positive by both blood
filtration and PCR, 4 samples positive by PCR only, and 3 samples positive by blood filtration only). Real-time
PCR of skin snip samples was significantly more sensitive than microscopic examination for the detection of
Onchocerca volvulus microfiladermia (2 of 218 samples positive by both microscopy and PCR and 12 samples
positive by PCR only). The molecular assays required smaller amounts of blood and tissue than conventional
methods and could be performed by laboratory personnel without specialized parasitology training. Taken
together, these data demonstrate the utility of the molecular diagnosis of filarial infections in mobile
Infections due to filarial nematodes are among the most
prevalent parasitic diseases throughout the world. Although
the transmission of these organisms is geographically restricted
to areas in developing countries where the disease is endemic,
modern human travel patterns have resulted in the migration
of infected individuals to regions where filarial infections have
been eradicated or have never been present, including re-
source-rich countries such as the United States. Despite being
relatively infrequent, filarial infections are sporadically diag-
nosed in refugees and other immigrants from areas of ende-
micity, in long-term residents of regions where filarial regions
are endemic (members of the armed services, students, mis-
sionaries, aid workers, and volunteers), and, rarely, among
Four filarial pathogens account for the vast majority of hu-
man disease. Wuchereria bancrofti (transmitted in sub-Saharan
Africa, Southeast Asia, the Caribbean, South America, and the
Western Pacific) and Brugia malayi (transmitted in Southern
and Southeast Asia, Indonesia, and the Philippines) are both
agents of lymphatic filariasis and together infect upwards of
120 million people. Onchocerca volvulus causes onchocerciasis,
or river blindness, in 20 to 40 million people, mainly in sub-
Saharan Africa but also to a lesser extent in Latin America and
the Arabian Peninsula. Loa loa, the African eyeworm, infects
between 3 and 13 million people in Central and Western
The diagnosis of filarial infections in mobile populations can
be challenging for several reasons. These infections are often
subclinical, manifest with nonspecific signs and symptoms
(such as itching or subcutaneous edema), or present clinically
after a prepatent period of months to years following exposure.
Because of the relative inexperience of many physicians in
countries where these pathogens are not endemic, the recog-
nition of these infections requires a high degree of clinical
suspicion. Patients often come to attention due to incidental
laboratory findings such as unexplained eosinophilia. Clinical
diagnosis is sometimes possible based on compatible exposure
history. More commonly, laboratory confirmation requires ex-
pert training in methods that may not be practiced routinely at
most medical centers.
Conventional parasitologic assays involve the isolation of
microfilariae from patient blood or tissue samples followed by
staining and identification of organisms by microscopic exam-
ination for key morphological features. Serological assays (a
limited number of which are available commercially) are an
alternative but suffer from poor specificity and an inability to
distinguish between active and prior infection (1, 20, 27). An
immunochromatographic card-type assay (Filariasis Now) de-
tects circulating antigen of W. bancrofti but is not useful for any
of the other pathogens (28) and is not available commercially
in Europe or North America. In recent years both conven-
tional and real-time PCR assays have been developed for all
four of the major filarial pathogens (7, 8, 11, 14, 21, 22, 24–26).
While these assays have shown great promise with regard to
* Corresponding author. Mailing address: LPD, NIAID, NIH, 4
Center Drive, Room B1-03, Bethesda, MD 20892. Phone: (301) 496-
5398. Fax: (301) 480-3757. E-mail: email@example.com.
† Present address: Division of Vaccines and Related Products Ap-
plications, Office of Vaccine Research and Regulation, Center for
Biologics Evaluation and Research, Food and Drug Administration,
Rockville, MD 20852.
?Published ahead of print on 27 October 2010.
high-level sensitivity and specificity, none is currently commer-
cially available, and none, to our knowledge, has been in use by
clinical pathology laboratories.
Selected filarial molecular diagnostic tools have been stud-
ied with patient populations in areas where filarial infections
are endemic (2, 4, 6, 18, 19), in the context of specific research
projects. However, the performance of these assays has not
been well described among internationally mobile populations
residing in resource-rich countries where the disease is not
endemic. The Clinical Parasitology Unit at the National Insti-
tute of Allergy and Infectious Diseases serves to evaluate and
treat patients with suspected parasitic diseases on a referral
basis. The patients are primarily immigrants, returned expatri-
ates, or travelers referred from throughout the United States
and, on occasion, internationally. Since 1999, we have incor-
porated a panel of real-time PCR assays adapted from previ-
ously described conventional PCR targets (10, 13, 16, 31, 32) as
part of routine clinical care. In this study, we have assessed the
performance of this molecular diagnostic panel in comparison
to conventional parasitology methods and report its utility over
the past decade among patients referred to the NIH for an
evaluation of suspected filarial infection.
MATERIALS AND METHODS
Patients and specimen collection. Assay data were collected prospectively
from all patients referred to the Clinical Parasitology Unit of the Laboratory of
Parasitic Diseases, National Institute of Allergy and Infectious Diseases, Na-
tional Institutes of Health, between April 1999 and December 2009. All patients
were evaluated under protocols approved by the NIAID Institutional Review
Board and registered (protocols NCT00001230 and NCT00001645). Written
informed consent was obtained from all subjects. Patients were either immigrants
from regions of the world where filarial disease is endemic or travelers to these
Diagnostic assays were selected based on compatible clinical symptoms as well
as geographic exposure history in relation to the known distributions of the major
human filarial pathogens. Laboratory evaluation for all patients included basic
screening studies (complete blood count, complete metabolic profile, urinalysis,
and stool examination for ova and parasites) as well as serology for antifilarial
IgG and IgG4 (12). For patients with suspected infection with blood-borne
filariae, whole-blood samples were obtained by venipuncture either at midday (L.
loa) or at midnight (W. bancrofti and B. malayi). For patients with suspected
onchocerciasis, six skin snip samples were typically obtained (1 mm3each from
shoulder, iliac crest, and thigh) by using a Walzer-type corneoscleral biopsy
instrument. Two of the six snips were submitted for DNA extraction and PCR
analysis, while the remaining four snips were examined by conventional methods.
Certain patients were evaluated longitudinally, with multiple blood or tissue
samples collected over time following treatment, and some patients had dupli-
cate blood specimens submitted on the same day or on consecutive days.
For the purposes of sensitivity and specificity calculations only, the “gold
standard” for loiasis required either the demonstration of microfilariae in the
blood, having an adult worm extracted, or having positive antifilarial antibodies
plus Calabar swellings and response to definitive diethylcarbamazine therapy.
For lymphatic filariasis the gold standard required a sample being circulating
filarial antigen positive and having positive antifilarial antibodies. For onchocer-
ciasis, the gold standard required either the demonstration of microfilariae in
skin snip samples or the presence of antifilarial antibodies and onchocerca-
specific antibodies (3), a compatible clinical picture, and a definitive response to
Parasitologic assays. Blood filtration and microscopic evaluation for micro-
filariae were performed by using 1 ml of anticoagulated blood as described
previously (15). Skin snip samples were immersed individually into normal sa-
line-filled wells of a plastic flat-bottom 96-well plate and examined after 24 h of
incubation at 37°C for the presence of O. volvulus microfilariae.
DNA extraction. Skin snip samples were digested in 200 ?l 0.1 M EDTA
disodium salt solution (Na2EDTA) (Sigma-Aldrich), 2 ?l proteinase K (20 mg/
ml) (Invitrogen), and 2 ?l 10% sodium dodecyl sulfate (SDS) solution (Sigma-
Aldrich) and incubated at 56°C in a heat block for 1 h. Following incubation, 4
?l 1.0 M dithiothreitol (DTT) solution (Sigma-Aldrich) was added, and the
samples were vortexed briefly and then incubated at 95°C in a heat block for 1 h.
The digested suspension was added to 0.9 ml NucliSens lysis buffer, and DNA
was extracted by using the NucliSens nucleic acid isolation kit as recommended
by the manufacturer (bioMe ´rieux).
Between 1999 and November 2001, 200 ?l of whole blood (EDTA) was
aliquoted into 0.9-ml NucliSens lysis buffer, and DNA was extracted by using the
NucliSens nucleic acid isolation kit. In an attempt to potentially increase the
PCR sensitivity, the volume of blood extracted by this method was increased to
1.0 ml (added to 9.0 ml lysis buffer) beginning in December 2001. To verify the
successful recovery of DNA and removal of PCR inhibitors, the lysis buffer
containing the digested tissue or whole-blood specimens were spiked with an
internal control (pBR322 plasmid DNA) before nucleic acid isolation. All sam-
ples were eluted with 50 ?l of elution buffer.
PCR assays. Two different types of assays were performed. Prior to May 2005,
all assays were performed as previously described (16), except that probes were
labeled with europium instead of fluorescein and detection of the amplification
products was performed by using the Delfia plate hybridization assay (Perkin-
Elmer Wallac, Inc.), with the resulting time-resolved fluorescence signals being
measured on a time-resolved fluorometer. The primers and probes were then
modified or redesigned (Table 1) in order to convert each assay to a real-time
Since May 2005, the B. malayi, L. loa, and W. bancrofti assays were performed
by using a LightCycler (LC) 1.2 instrument (Roche Diagnostics) with a 20-?l
reaction mixture consisting of 1? LC FastStart DNA Master HybProbe mixture
containing FastStart Taq polymerase, reaction buffer deoxynucleoside triphos-
phate (dNTP) mix (with dUTP substituted for dTTP), 1.0 mM MgCl2(Roche),
an additional 3.0 mM MgCl2, 1.0 ?M each primer, 0.2 ?M each fluorescence
resonance energy transfer (FRET) probe, 1 U uracil-DNA-glycosylase (UNG),
and 10 ?l of extracted DNA. The reaction mixture was preincubated for 10 min
at 30°C to activate UNG, and DNA was denatured and UNG was inactivated at
95°C for 10 min. The template amplification consisted of 45 cycles of 5 s at 95°C,
10 s at 55°C, and 20 s at 72°C. The O. volvulus real-time PCR was performed in
a 25-?l reaction mixture consisting of 1? QuantiTect SYBR green PCR master
mix (Qiagen) containing HotStarTaq DNA polymerase, reaction buffer, 2.5 mM
MgCl2, dNTP mix (containing a dTTP-dUTP mixture), SYBR green I and ROX
(6-carboxy-X-rhodamine) fluorescent dyes, 0.5 ?M each primer, 0.5 U UNG,
and 5 ?l of extracted DNA. Amplification was performed with a Rotorgene-3000
instrument (Qiagen) with cycling parameters of 10 min at 30°C, 10 min at 95°C,
and 45 cycles of 15 s at 95°C, 30 s at 57°C, and 30 s at 72°C. A melt curve analysis
was then performed by reducing the temperature to 60°C for 45 s and then
raising the temperature 1°C every 5 s up to 99°C. To be considered positive, the
melt peak temperature from the patient specimen must match the positive
control within the specified range (72°C ? 2°C). The internal control in
extracted samples was detected by amplification in a separate qualitative LC
real-time PCR as described previously (5).
The analytical sensitivity analysis of the assays used in this study demonstrated
lower limits of detection equal to 10 fg/?l for B. malayi, 2.5 fg/?l for L. loa, 100
fg/?l for W. bancrofti, and 400 fg/?l for O. volvulus.
Two hundred patients were evaluated at our clinic for sus-
pected filarial infections between April 1999 and December
2009. Parasitologic and molecular diagnostic testing for these
patients is summarized in Table 2. In total, 392 specimens were
collected (256 blood samples and 136 skin snip sets), and 887
diagnostic assays were performed on these specimens (356
conventional parasitologic studies and 531 PCR assays). All
skin snip microscopic studies were paired with at least one (and
usually two) PCR assay performed on the same tissue, while
the pairing of blood filtration studies with PCR assays per-
formed on the same blood samples was variable. In total, 19
patients were diagnosed with L. loa infection, 11 were diag-
nosed with onchocerciasis, 1 was diagnosed with lymphatic
filariasis due to W. bancrofti, and none was diagnosed with
infection due to B. malayi (Fig. 1).
The performance of PCR assays on blood samples in com-
parison to blood filtration is summarized in Table 3. Of 208
VOL. 49, 2011CLINICAL EVALUATION OF FILARIAL MOLECULAR DIAGNOSTICS43
specimens tested for L. loa microfilaremia, 24 were positive by
both PCR and blood filtration, while 177 were negative by both
methods. L. loa microfilaremia levels among positive blood
filtration assays ranged from 1 organism/ml to 7,400 organisms/
ml. Discordant results occurred for 7 specimens, with 4 being
positive by PCR only and 3 being positive by blood filtration
only (1 to 2 organisms/ml). Removing posttreatment samples,
there were 17 diagnoses of loiasis made concurrently by PCR
and blood filtration, 1 diagnosis by blood filtration alone, and
2 diagnoses by PCR alone.
Comparative data were available for fewer samples when the
performance of the W. bancrofti and B. malayi PCR assays
were evaluated. Forty-six paired assays for W. bancrofti micro-
filaremia resulted in 2 specimens that were positive by both
PCR and blood filtration (100 to 800 organisms/ml), 44 spec-
imens that were negative by both methods, and no discordant
results. B. malayi was not detected in any sample either by PCR
or by blood filtration.
Mansonella perstans was detected by blood filtration in 3
samples (7 to 64 organisms/ml), including one patient who had
a coinfection with L. loa detected by both PCR and blood
filtration (data not shown). No other filarial coinfections
were diagnosed. There were no paired blood samples for
which discordant pathogens were identified by blood filtra-
tion and PCR.
Laboratory evaluation of patients with suspected onchocer-
ciasis, summarized in Table 4, included 218 skin snip assays
with paired microscopic and PCR assay results. There were 14
positive PCR assays from 11 patients, only 2 of which (both
from a single patient’s skin snip set) were also positive by
conventional parasitology. No skin snip samples were positive
by microscopic evaluation but negative by PCR.
The clinical performance statistics for L. loa and O. volvulus
PCR assays that were performed during initial patient evalu-
FIG. 1. Diagnosis of filarial infections by PCR compared to blood
filtration and skin snip microscopy.
TABLE 1. Primer and probe sequences, product size, and target DNA for each PCR assay
Assay target and
primer or probe
324 HhaI repeat
503 Loa interspersed repeat
134 SspI repeat
aFluor., fluorescein label; Phos., phosphate cap.
TABLE 2. Summary data for parasitologic and molecular testing
for suspected filarial infections
Evaluation for suspected
PCR for L. loa
PCR for W. bancrofti
PCR for B. malayi
PCR for O. volvulus
aNA, not applicable.
44FINK ET AL.J. CLIN. MICROBIOL.
ations showed that the sensitivity, specificity, positive predic-
tive value (PPV), and negative predictive value (NPV) were all
100% for the L. loa and O. volvulus assays based on the out-
lined gold standard criteria. The inclusion of follow-up (post-
treatment) assays in the statistical analysis decreased the sen-
sitivity and NPV of the L. loa PCR assay to 90.3% (28/31
samples) and 98.6% (206/209), respectively, with no changes in
the other L. loa or O. volvulus statistics. Performance statistics
were not calculated for the B. malayi and W. bancrofti assays
due to an insufficient number of infections.
An accurate diagnosis of filarial infections in mobile popu-
lations can be challenging. Infected individuals may present
with nonspecific symptoms or laboratory findings, and proper
evaluation requires both a strong degree of clinical suspicion as
well as specialized knowledge of filarial epidemiology and
pathogenesis and expertise in the morphological classification
of filarial parasites by microscopic examination. Further diffi-
culties arise from the extremely limited commercial availability
of filarial diagnostic assays that can distinguish not only be-
tween specific pathogens but also between active current in-
fections and those that occurred in the past.
In this study, we assessed the performance and feasibility in
a clinical diagnostic setting of a panel of real-time PCR assays
designed to detect species-specific genomic DNA target se-
quences of the four most prevalent pathogenic filariae of hu-
mans: W. bancrofti, B. malayi, O. volvulus, and L. loa. Com-
pared to conventional parasitology methods, our PCR assays
were overall equal to or significantly more sensitive among
blood and skin snip samples collected from a cohort of 200
patients undergoing evaluations for suspected filarial infec-
Differences in assay performance were most striking for the
detection of O. volvulus in skin snip samples, with diagnosis by
a positive PCR assay for 11 patients compared to diagnosis by
microscopy for only one patient. One possible interpretation of
this discordance is that only one patient was truly infected with
O. volvulus, with the remaining diagnoses representing false-
positive PCR assays. This situation is unlikely, however, since
DNA extraction and assay setup were conducted under rigor-
ous conditions to protect against cross-sample contamination,
and all PCR runs included internal negative controls with ver-
ified negative assay results. Moreover, each of the positive
patients was treated definitively with ivermectin, and each pa-
tient had a clinical response. Some patients also had a Maz-
zotti-type reaction following ivermectin treatment, which indi-
cates a high likelihood of O. volvulus infection. Each of the
PCR-positive patients was also found retrospectively to be
positive by highly O. volvulus-specific serological assays (3, 28;
data not shown).
More likely, discordances represent situations in which very
low numbers of O. volvulus microfilariae are present in skin
snip samples such that microscopy is truly insensitive com-
pared to PCR (23). It is possible that PCR allows the detection
of O. volvulus DNA found within the entirety of the tissue,
while microscopy detects only organisms that are capable of
extruding themselves from submerged tissue samples. The in-
creased sensitivity of our O. volvulus PCR assay is notable in
that for each patient evaluation, 4 to 6 skin snip samples were
typically examined microscopically, while only 2 samples were
processed for DNA extraction and PCR. Our PCR assay there-
fore achieved a higher rate of detection despite having to
overcome a potential loss of sensitivity due to sampling error.
Furthermore, there were no patients diagnosed with O. volvu-
lus infection by conventional microscopy but negative by PCR
The performance of our PCR assay for L. loa infection was
similar to that of blood filtration. Three paired assays were
positive by blood filtration but negative by PCR, all of which
were posttreatment samples collected from patients whose
pretreatment evaluation included positive blood filtration and
PCR assays on paired samples. Quantification of microfilare-
mia by blood filtration in each of these instances was only 1 or
2 organisms/ml, and there were several positive PCR assays for
which the paired blood filtration identified microfilaremia at
only 1 to 2 organisms/ml. Therefore, the false-negative PCR
assays were most likely due to sampling error in the context of
very low-level microfilaremia rather than any inherent inability
of the assay to detect small quantities of parasite DNA. In
further support of this interpretation, there were four paired
blood samples (from four patients) for which PCR was positive
but blood filtration was negative. Two of the patients had
recently undergone medical therapy for loiasis, with pretreat-
ment blood samples being positive by both PCR and blood
filtration, while the other two patients likely had low-level
microfilaremia upon initial evaluation. These four cases illus-
trate the utility of our L. loa PCR assay for the detection of
low-level infection both at the time of diagnosis and during the
course of monitoring the response to therapy.
An important limitation of our study is that our clinic eval-
uated on average only 20 new patients each year, reflecting the
scarcity of opportunities to diagnose filarial infections in the
United States, even at a national referral center. In particular,
there were relatively few evaluations for suspected lymphatic
filariasis. Due to the limited number of positive assays for W.
bancrofti and B. malayi (either PCR or blood filtration), it is
difficult to draw firm conclusions regarding the performance of
TABLE 3. Comparison of results from paired blood filtration and
PCR assays on blood samples from patients with
suspected filarial infections
No. of samples with result
L. loa W. bancrofti B. malayi
Blood filtration positive
Blood filtration negative
TABLE 4. Results of PCR assays on skin snip samples from
patients with suspected onchocerciasis
No. of samples with result
Skin snip positive
Skin snip negative
VOL. 49, 2011CLINICAL EVALUATION OF FILARIAL MOLECULAR DIAGNOSTICS 45
our PCR diagnostics for these organisms, although these types
of assays have been used successfully in research laboratories
in countries where filarial disease is endemic.
In addition to observed gains in sensitivity, there are some
distinct advantages of PCR in comparison to other available
diagnostic methodologies. First, PCR directly detects filarial
DNA, in contrast to serological and circulating-antigen assays,
which measure indirect indicators of infection (e.g., antibod-
ies) and which may be persistently positive long after all or-
ganisms have died (for both antibody and circulating-antigen
assays). Second, our PCR assays achieve similar increased sen-
sitivities compared to that of conventional microscopy while
requiring smaller amounts of patient material as a starting
point (200 ?l of blood for PCR versus 1 ml or greater for
filtration and 1 to 2 skin snip samples for PCR versus 6 skin
snip samples for microscopy). Finally, PCR assays can be run
by laboratory personnel with generalized training and do not
require a specialized knowledge of parasite morphology and
There are several disadvantages of PCR that must also be
recognized. At this time, PCR assays require costly reagents
such as kits for DNA extraction from blood or tissue, enzymes
and primers that must be stored frozen, and thermal cycler
machines with the ability to detect fluorescence emission (for
real-time PCR assays). On a per-assay cost (between $10 and
$12 ), once equipment is in place, PCR is likely to be of
equal or lower cost than antibody-based or parasitologic meth-
ods because of significantly lower labor costs than those of
PCR may soon become more suitable for point-of-care use
in resource-poor countries with ongoing advances in the de-
velopment of hand-held, battery-operated devices using mi-
crofluidic methods for all-in-one DNA extraction, amplifica-
tion, and detection (9, 17, 29, 30). Unlike the situation with
blood filtration, our PCR assay results are currently not re-
ported quantitatively. However, the generation of a standard
curve using defined numbers of organisms would be a relatively
easy adjustment to the real-time PCR format to allow a quan-
titative assessment of microfilaremia (methods which we have
recently developed for L. loa). Another avenue for improve-
ment would be to multiplex the PCR assays using a different
fluorescent reporter for each organism. Such an adjustment
would allow the detection of microfilarial coinfections by a
single assay, negating the advantage that blood filtration pro-
vides in this regard. The development and validation of an M.
perstans-specific PCR assay would also be necessary for this
In summary, we have demonstrated the utility and feasibility
of a panel of real-time PCR assays for the diagnosis of filarial
infections among immigrants and travelers which have been
used clinically for more than a decade. Our data demonstrate
that the PCR panel is exquisitely species specific and slightly
more sensitive than blood filtration for the detection of micro-
filaremia. Additionally, our real-time PCR assay for O. volvulus
is significantly more sensitive than conventional microscopy for
the detection of skin microfilariae. Although not quite ready
for widespread use in areas of endemicity, the successful per-
formance of these molecular assays is an important step for-
ward in making accurate filarial diagnostic tools more accessi-
ble to clinical parasitology programs that serve internationally
This research was supported by the Intramural Research Program of
the NIAID, NIH.
1. Akue, J. P., T. G. Egwang, and E. Devaney. 1994. High levels of parasite-
specific IgG4 in the absence of microfilaremia in Loa loa infection. Trop.
Med. Parasitol. 45:246–248.
2. Boatin, B. A., L. Toe, E. S. Alley, N. J. Nagelkerke, G. Borsboom, and J. D.
Habbema. 2002. Detection of Onchocerca volvulus infection in low preva-
lence areas: a comparison of three diagnostic methods. Parasitology 125:
3. Burbelo, P. D., H. P. Leahy, M. J. Iadarola, and T. B. Nutman. 2009. A
four-antigen mixture for rapid assessment of Onchocerca volvulus infection.
PLoS Negl Trop. Dis. 3:e438.
4. Chansiri, K., and S. Phantana. 2002. A polymerase chain reaction assay for
the survey of bancroftian filariasis. Southeast Asian J. Trop. Med. Public
5. Cohen, J. I., G. A. Fahle, M. A. Kemp, K. Apakupakul, and T. P. Margolis.
2010. Human herpesvirus 6-A, 6-B, and 7 in vitreous fluid samples. J. Med.
6. Dissanayake, S., A. Rocha, J. Noroes, Z. Medeiros, G. Dreyer, and W. F.
Piessens. 2000. Evaluation of PCR-based methods for the diagnosis of in-
fection in bancroftian filariasis. Trans. R. Soc. Trop. Med. Hyg. 94:526–530.
7. Fischer, P., T. Supali, H. Wibowo, I. Bonow, and S. A. Williams. 2000.
Detection of DNA of nocturnally periodic Brugia malayi in night and day
blood samples by a polymerase chain reaction-ELISA-based method using
an internal control DNA. Am. J. Trop. Med. Hyg. 62:291–296.
8. Hassan, M., M. M. Sanad, I. el-Karamany, M. Abdel-Tawab, M. Shalaby, A.
el-Dairouty, K. Assal, M. K. Gamal-Edin, and M. Adel el-Kadi. 2005. De-
tection of DNA of W. bancrofti in blood samples by QC-PCR-ELISA-based.
J. Egypt. Soc. Parasitol. 35:963–970.
9. Hua, Z., J. L. Rouse, A. E. Eckhardt, V. Srinivasan, V. K. Pamula, W. A.
Schell, J. L. Benton, T. G. Mitchell, and M. G. Pollack. 2010. Multiplexed
real-time polymerase chain reaction on a digital microfluidic platform. Anal.
10. Klion, A. D., N. Raghavan, P. J. Brindley, and T. B. Nutman. 1991. Cloning
and characterization of a species-specific repetitive DNA sequence from Loa
loa. Mol. Biochem. Parasitol. 45:297–305.
11. Kluber, S., T. Supali, S. A. Williams, E. Liebau, and P. Fischer. 2001. Rapid
PCR-based detection of Brugia malayi DNA from blood spots by DNA
detection test strips. Trans. R. Soc. Trop. Med. Hyg. 95:169–170.
12. Lal, R. B., and E. A. Ottesen. 1988. Enhanced diagnostic specificity in human
filariasis by IgG4 antibody assessment. J. Infect. Dis. 158:1034–1037.
13. Lizotte, M. R., T. Supali, F. Partono, and S. A. Williams. 1994. A polymerase
chain reaction assay for the detection of Brugia malayi in blood. Am. J. Trop.
Med. Hyg. 51:314–321.
14. McCarthy, J. S., M. Zhong, R. Gopinath, E. A. Ottesen, S. A. Williams, and
T. B. Nutman. 1996. Evaluation of a polymerase chain reaction-based assay
for diagnosis of Wuchereria bancrofti infection. J. Infect. Dis. 173:1510–
15. McPherson, T., and T. B. Nutman. 2007. Filarial nematodes. In P. R. Mur-
ray, et al. (ed.), Manual of clinical microbiology, 9th ed. ASM Press, Wash-
16. Nutman, T. B., P. A. Zimmerman, J. Kubofcik, and D. D. Kostyu. 1994. A
universally applicable diagnostic approach to filarial and other infections.
Parasitol. Today 10:239–243.
17. Pipper, J., M. Inoue, L. F. Ng, P. Neuzil, Y. Zhang, and L. Novak. 2007.
Catching bird flu in a droplet. Nat. Med. 13:1259–1263.
18. Rahmah, N., A. N. Ashikin, A. K. Anuar, R. H. Ariff, B. Abdullah, G. T.
Chan, and S. A. Williams. 1998. PCR-ELISA for the detection of Brugia
malayi infection using finger-prick blood. Trans. R. Soc. Trop. Med. Hyg.
19. Ramzy, R. M. 2002. Field application of PCR-based assays for monitoring
Wuchereria bancrofti infection in Africa. Ann. Trop. Med. Parasitol.
20. Rao, K. V., M. Eswaran, V. Ravi, B. Gnanasekhar, R. B. Narayanan, P.
Kaliraj, K. Jayaraman, A. Marson, N. Raghavan, and A. L. Scott. 2000. The
Wuchereria bancrofti orthologue of Brugia malayi SXP1 and the diagnosis of
bancroftian filariasis. Mol. Biochem. Parasitol. 107:71–80.
21. Rao, R. U., L. J. Atkinson, R. M. Ramzy, H. Helmy, H. A. Farid, M. J.
Bockarie, M. Susapu, S. J. Laney, S. A. Williams, and G. J. Weil. 2006. A
real-time PCR-based assay for detection of Wuchereria bancrofti DNA in
blood and mosquitoes. Am. J. Trop. Med. Hyg. 74:826–832.
22. Rao, R. U., G. J. Weil, K. Fischer, T. Supali, and P. Fischer. 2006. Detection
of Brugia parasite DNA in human blood by real-time PCR. J. Clin. Micro-
46FINK ET AL.J. CLIN. MICROBIOL.
23. Taylor, H. R., B. Munoz, E. Keyvan-Larijani, and B. M. Greene. 1989.
Reliability of detection of microfilariae in skin snips in the diagnosis of
onchocerciasis. Am. J. Trop. Med. Hyg. 41:467–471.
24. Toe, L., B. A. Boatin, A. Adjami, C. Back, A. Merriweather, and T. R.
Unnasch. 1998. Detection of Onchocerca volvulus infection by O-150 poly-
merase chain reaction analysis of skin scratches. J. Infect. Dis. 178:282–285.
25. Toure, F. S., O. Bain, E. Nerrienet, P. Millet, G. Wahl, Y. Toure, O. Doumbo,
L. Nicolas, A. J. Georges, L. A. McReynolds, and T. G. Egwang. 1997.
Detection of Loa loa-specific DNA in blood from occult-infected individuals.
Exp. Parasitol. 86:163–170.
26. Toure, F. S., L. Kassambara, T. Williams, P. Millet, O. Bain, A. J. Georges,
and T. G. Egwang. 1998. Human occult loiasis: improvement in diagnostic
sensitivity by the use of a nested polymerase chain reaction. Am. J. Trop.
Med. Hyg. 59:144–149.
27. Vincent, J. A., S. Lustigman, S. Zhang, and G. J. Weil. 2000. A comparison
of newer tests for the diagnosis of onchocerciasis. Ann. Trop. Med. Parasitol.
28. Weil, G. J., C. Steel, F. Liftis, B. W. Li, G. Mearns, E. Lobos, and T. B.
Nutman. 2000. A rapid-format antibody card test for diagnosis of onchocer-
ciasis. J. Infect. Dis. 182:1796–1799.
29. Wulff-Burchfield, E., W. A. Schell, A. E. Eckhardt, M. G. Pollack, Z. Hua,
J. L. Rouse, V. K. Pamula, V. Srinivasan, J. L. Benton, B. D. Alexander, D. A.
Wilfret, M. Kraft, C. B. Cairns, J. R. Perfect, and T. G. Mitchell. 2010.
Microfluidic platform versus conventional real-time polymerase chain reac-
tion for the detection of Mycoplasma pneumoniae in respiratory specimens.
Diagn. Microbiol. Infect. Dis. 67:22–29.
30. Yeung, S. S., T. M. Lee, and I. M. Hsing. 2008. Electrochemistry-based
real-time PCR on a microchip. Anal. Chem. 80:363–368.
31. Zhong, M., J. McCarthy, L. Bierwert, M. Lizotte-Waniewski, S. Chanteau,
T. B. Nutman, E. A. Ottesen, and S. A. Williams. 1996. A polymerase chain
reaction assay for detection of the parasite Wuchereria bancrofti in human
blood samples. Am. J. Trop. Med. Hyg. 54:357–363.
32. Zimmerman, P. A., R. H. Guderian, E. Aruajo, L. Elson, P. Phadke, J.
Kubofcik, and T. B. Nutman. 1994. Polymerase chain reaction-based diag-
nosis of Onchocerca volvulus infection: improved detection of patients with
onchocerciasis. J. Infect. Dis. 169:686–689.
VOL. 49, 2011 CLINICAL EVALUATION OF FILARIAL MOLECULAR DIAGNOSTICS47