Discrimination between six species of canine microfilariae Mark Rishniw ,

Veterinary Parasitology 135 (2006) 303–314
www.elsevier.com/locate/vetpar
Discrimination between six species of canine microfilariae
by a single polymerase chain reaction
Mark Rishniw a,*, Stephen C. Barr b, Kenny W. Simpson b,
Marguerite F. Frongillo c, Marc Franz d, Jose Luis Dominguez Alpizar e
a
Department of Biomedical Sciences, Box 11, Veterinary Research Tower, College of Veterinary Medicine,
Cornell University, Ithaca, NY 14853, USA
b
Department of Clinical Sciences, Cornell University, Ithaca, NY, USA
c
Department of Microbiology and Immunology, Cornell University, Ithaca, NY, USA
d
Woodbury Animal Hospital, Woodbury, NY, USA
e
Department of Parasitology, Universidad Autónoma de Yucatán, Mexico
Received 16 August 2005; received in revised form 6 October 2005; accepted 6 October 2005
Abstract
Canine dirofilariasis caused by Dirofilaria immitis is usually diagnosed by specific antigen testing and/or identification of
microfilariae. However, D. immitis and at least six other filariae can produce canine microfilaremias with negative heartworm
antigen tests. Discriminating these can be of clinical importance.
To resolve discordant diagnoses by two diagnostic laboratories in an antigen-negative, microfilaremic dog recently imported
into the US from Europe we developed a simple molecular method of identifying different microfilariae, and subsequently
validated our method against six different filariae known to infect dogs by amplifying ribosomal DNA spacer sequences by
polymerase chain reaction using common and species-specific primers, and sequencing the products to confirm the genotype of
the filariae. We identified the filaria in this dog as D. repens. This is the first case of D. repens infection in the United States.
Additionally, we examined microfilariae from five additional antigen-negative, microfilaremic dogs and successfully
identified the infecting parasite in each case. Our diagnoses differed from the initial morphological diagnosis in three of
these cases, demonstrating the inaccuracy of morphological diagnosis. In each case, microfilariae identified morphologically as
A. reconditum were identified as D. immitis by molecular methods. Finally, we demonstrated that our PCR method should
amplify DNA from at least two additional filariae (Onchocerca and Mansonella), suggesting that this method may be suitable for
genotyping all members of the family Onchocercidae.
# 2005 Elsevier B.V. All rights reserved.
Keywords: Dirofilaria; Immitis; Dipetalonema; Acanthocheilonema; Reconditum; Repens; Canine; Knott’s test; Polymerase chain reaction;
PCR
* Corresponding author. Tel.: +1 607 253 3108.
E-mail address: mr89@cornell.edu (M. Rishniw).
0304-4017/$ – see front matter # 2005 Elsevier B.V. All rights reserved.
doi:10.1016/j.vetpar.2005.10.013
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M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314
1. Introduction
Canine heartworm disease is generally diagnosed
by antigen testing for Dirofilaria immitis, and/or
identification of microfilariae in the blood of infected
dogs. However, other filariae, including Acanthocheilonema (Dipetalonema) reconditum, Dirofilaria
(Nochtiella) repens, Acanthocheilonema (Dipetalonema) dracunculoides, Cercopithifilaria grassi, Brugia malayi, Brugia pahangi, Brugia ceylonensis and
approximately 1% of Dirofilaria immitis infestations,
can produce persistent microfilaremias with negative
heartworm antigen tests (Courtney et al., 1993;
Fischer et al., 2002; Genchi, 2003). In such cases,
diagnosis relies on examination of the microfilariae,
generally using a modified Knott’s test, or alkaline
phosphatase staining (Chalifoux and Hunt, 1971;
Knott, 1935). These methods are imperfect, and rely
on specialist training to accurately differentiate the
filariae. However, in regions where one of these
filariae is not enzootic, parasitologists may exclude
that filaria from consideration, or fail to recognize it,
especially if a history of potential exposure is not
provided (Cordonnier et al., 2002).
Accurate identification of filarial species in dogs
can be clinically important, because of the zoonotic
concerns and therapeutic implications. While A.
reconditum and A. dracunculoides have few (if any)
clinical consequences, D. repens infestations have
been associated with subcutaneous granulomas and
pruritus in dogs (Baneth et al., 2002; Bredal et al.,
1998; Manuali et al., 2005; Tarello, 2003), and
subcutaneous, conjunctival and pulmonary nodules,
often confused with neoplastic tumors, in humans
(Cordonnier et al., 2002; Pampiglione et al., 1996,
2001; Romano et al., 1976; Vakalis and Himonas,
1997). To reduce the risk of zoonotic transmission,
treatment of D. repens infestations with filaricidal
agents, or at least microfilaricidal agents, is recommended (adulticidal therapy can be postponed until
clinical signs occur if risk of treatment is deemed to
outweigh risk of disease). Adult D. repens have been
successfully treated with melarsomine hydrochloride
(Immiticide, Merial; two injections of 2.5 mg/kg IM
24 h apart) (Baneth et al., 2002). Microfilariae are
susceptible to microfilaricidal doses of avermectins
(ivermectin 50 mg/kg PO or SQ; doramectin 400 mg/
kg SQ; effects of milbemycin have not been
investigated) (Baneth et al., 2002; Tarello, 2003).
On the other hand, treatment of A. dracunculoides or
A. reconditum infestations is not usually required, so
they should be positively identified to avoid inappropriate adulticide therapy.
Over the last decade, several investigators have
examined the molecular differentiation of filariae,
using Southern blotting to identify DNA repeats from
D. repens (Chandrasekharan et al., 1994) or polymerase chain reaction (PCR) to differentiate either D.
repens or A. reconditum from D. immitis (Bredal et al.,
1998; Favia et al., 1996, 1997; Mar et al., 2002;
Vakalis et al., 1999). However, no existing studies
examined the ability of PCR to differentiate between
more than two filariae. While it would be possible to
run multiple PCR reactions using published primer
sequences to identify a specific filaria, a single
differentiating reaction would provide an expeditious
means of obtaining a diagnosis. Additionally,
sequences are not available for all filariae of interest
at this time, so species-specific reactions may fail in
these cases.
This study was prompted by several clinical cases
requiring accurate identification of microfilariae in
dogs. These dogs had negative heartworm antigen
tests and, in several instances, discrepant morphological identifications of the microfilariae. Using
molecular techniques, we sought to accurately identify
the filarial species in each case, and subsequently
developed a PCR-based method that distinguishes
between the six species tested to date (D. immitis, A.
reconditum, D. repens, A. dracunculoides, B. pahangi,
B. malayi) with a single primer pair. Our technique
simplifies filarial identification, compared with previous studies.
2. Materials and methods
2.1. Case reports
2.1.1. Case 1
A 2-year-old intact male Labrador Retriever, born
in the Czech Republic, but exported as a puppy to The
Netherlands, was presented to the referring veterinarian (MF) for routine evaluation after importation from
The Netherlands in January 2004 to Canada and
subsequently New York State (United States) in April
M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314
2004. The dog was clinically normal and had never
been tested for heartworm or administered heartworm
preventative. The veterinarian identified a microfilaremia on routine heartworm screening, and performed
a heartworm antigen test using a Dirocheck ELISA
(Synbiotics Corporation, San Diego, CA), which was
negative. Repeated antigen testing at several diagnostic laboratories failed to diagnose D. immitis
infestation. Consequently, the veterinarian submitted
blood for morphological classification of microfilariae
to two diagnostic laboratories. Examination of the
microfilariae using modified Knott’s tests resulted in
discrepant diagnoses: one laboratory identified the
microfilariae as A. reconditum, while the other
identified them as D. repens.
2.1.2. Case 2
An 11-year-old male neutered mixed breed dog
was presented to a referring veterinarian in San Diego,
California, for routine evaluation. As with the first
case, a microfilaremia was detected, but antigen
testing for D. immitis was negative on multiple
occasions. The microfilariae were identified as A.
reconditum by a commercial diagnostic laboratory.
2.1.3. Case 3
An 8-year-old dog (signalment unknown) was
presented to the referring veterinarian in Utica, New
York, for evaluation of hematuria secondary to
prostatitis. Microfilariae were detected on urinalysis,
and subsequently on examination of blood. Antigen
tests for D. immitis were negative. The microfilariae
were morphologically identified as D. immitis with a
Knott’s test.
2.1.4. Case 4
An 8-year-old female speyed miniature schnauzer
was presented to the referring veterinarian after
adoption from an animal shelter in Clemmons, North
Carolina. Again, a microfilaremia was detected and
identified morphologically as A. reconditum, but
antigen testing for D. immitis was negative on
multiple occasions.
2.1.5. Case 5
A 1-year-old male intact Pug was presented for
heartworm testing in Modesto, California. Microfilaremia was detected and identified morphologically as
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A. reconditum, but antigen testing for D. immitis was
negative.
2.1.6. Case 6
An 8-year-old male neutered Malamute was
presented for heartworm testing in Vallejo, California.
Microfilaremia was detected, and identified morphologically as A. reconditum but antigen testing for D.
immitis was negative.
2.2. Sample preparation
In all six clinical cases, 1 ml of blood (anticoagulated with heparin or EDTA) was submitted.
We used blood from a microfilaremic dog with D.
immitis infestation and from a dog with A. reconditum
infestation as positive controls. In both controls
antigen testing and Knott’s testing provided the
filarial diagnosis. The sample of A. recondituminfested blood was obtained by one of the authors and
morphologically identified as A. reconditum. Fresh
adult B. malayi and B. pahangi worms were
generously provided by T. Klei of Louisiana State
University. Blood from a dog infested with A.
dracunculoides was generously provided by Francisco
Alonso de Vega of Murcia University in Spain after
morphologic diagnosis with alkaline phosphatase
staining.
Finally, we used DNA from a non-infected dog as a
negative control in all PCR reactions.
2.3. DNA extraction
In all clinical cases, the D. immitis positive control
specimen and the Brugia specimens, 500 ml anticoagulated blood (or five adult Brugia filariae) were
suspended in 500 ml lysis buffer (50 mM Tris, pH 8.0;
100 mM EDTA, 100 mM NaCl, 1% SDS) and
digested with 10 ml proteinase K (10 mg/ml) for
16 h at 55 8C, followed by a standard phenol/chloroform extraction method, as described previously
(Sambrook and Russell, 2001).
The A. reconditum and A. dracunculoides positive
control specimens were submitted on filter paper as
dried blood spots. DNA was extracted using a QIAmp
DNA Blood Mini Kit DNA extraction kit (Qiagen
Inc., Valencia, CA) according to manufacturer’s
instructions.
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2.4. Polymerase chain reaction filarial genotyping
We designed primer pairs that spanned the internal
transcribed spacer region 2 (ITS2) of the ribosomal
DNA of either D. immitis, or A. reconditum, and a
primer pair spanning the 5S ribosomal intergenic region
of D. repens based on previously published sequences
(Table 1) (Favia et al., 2000; Mar et al., 2002). These
primers were used to specifically genotype each filarial
species. Additionally, we designed a primer pair that
would amplify fragments of different fragment length
from both D. immitis and A. reconditum (DIDR-F1,
DIDR-R1) (referred to as the pan-filarial primers
throughout the remainder of the manuscript).
For additional independent genotyping validation,
we designed primers from published sequences of the
cytochrome oxidase subunit 1 (COI) gene for each of
these species (Table 1).
Each PCR reaction consisted of 1.5 mM MgCl2,
250 mM dNTPs, 20 mM Tris–HCl (pH 8.4), 50 mM
KCl, 2.5 U of Taq polymerase and 1 ml of DNA
solution in a total volume of 20 ml. The PCR
procedure consisted of a denaturing step at 94 8C
for 2 min and 32 cycles of denaturing (30 s at 94 8C),
annealing (30 s at 60 8C for ITS2-based primers and
30 s at 58 8C for COI-based primers) and extension
(30 s at 72 8C); a final extension (7 min at 72 8C) and a
soak at 4 8C in an Eppendorf Mastercycler thermal
cycler (Brinkmann Instruments Inc., Westbury, NY).
The PCR product (10 ml) was examined on a 1.5%
agarose gel, and fragments of predicted size were
extracted from both reactions and purified using a
QiaQuick PCR purification kit (Qiagen Inc., Valencia,
CA). The PCR products were ligated into pGEM-T
Easy Vector System 1 TA cloning vectors (Promega
US, Madison, WI) and cloned (Sambrook and Russell,
2001). Candidate clones were screened by restriction
digests, and clones with expected digestion fragments
were sequenced at the institutional sequencing facility
on an Applied Biosystems Automated 3730 DNA
Table 1
Primer sequences used to amplify PCR products from filarial and canine DNA
Primer pair Primer sequence
Gene target
D.imm-F1a
D.imm-R1
A.rec-F1a
A.rec-R1
D.rep-F1
D.rep-R1
DIDR-F1
DIDR-R1
CAT CAG GTG ATG ATG TGA TGA T
TTG ATT GGA TTT TAA CGT ATC ATT T
CAG GTG ATG GTT TGA TGT GC
CAC TCG CAC TGC TTC ACT TC
TGT TTC GGC CTA GTG TTT CGA CCA
ACG AGA TGT CGT GCT TTC AAC GTG
AGT GCG AAT TGC AGA CGC ATT GAG
AGC GGG TAA TCA CGA CTG AGT TGA
ITS2
22
25
ITS2
20
20
5SrRNA
24
24
5.8S-ITS2-28S 24
24
DI COI -F1
DI COI-R1
AR COI-F1
AR COI-R1
DR COI-F1
DR COI-R1
Actin F2
Actin R2
AGT GTA GAG GGT CAG CCT GAG TTA
ACA GGC ACT GAC AAT ACC AAT
AGT GTT GAG GGA CAG CCA GAA TTG
CCA AAA CTG GAA CAG ACA AAA CAA GC
AGT GTT GAT GGT CAA CCT GAA TTA
GCC AAA ACA GGA ACA GAT AAA ACT
ACC ACT GGT ATT GTC ATG GAC TCT G
GCT CTT CTC CAG GGA GGA CGA
COI
a
Primer size Product
(base-pairs) origin
24
21
COI
24
26
COI
24
24
Canine b-actin 25
21
Product size Genbank
(base-pairs) accession #
D. immitis
302
AF217800
A. reconditum
348
AF217801
D. repens
247 and 153 AJ242966
AJ242967
D. immitis
542
AF217800
A. reconditum
578
AF217801
D. repens
484
AY693808
A. dracunculoides 584
DQ018785
B. pahangi
664
AY988600
B. malayi
615
AY988599
B. timori
625b
AF499130
M. ozzardi
430b
AF228554
O. volvulus
470b
AF228575
D. immitis
203
AB192887
A. reconditum
208
AJ544876
D. repens
209
AJ271614
Canis familiaris
276
AF021873
D.imm-F1, D.imm-R1, A.rec-F1 and A.rec-R1 are identical to ITS2F-Di, ITS2R-Di, ITS2F-Dr and ITSR-Dr primers published by Mar et al.
(2002).
b
Predicted product size based on Genbank sequence.
M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314
307
Analyzer (Applied Biosystems, Foster City, CA) using
universal M13 forward and reverse primers. Sequence
results were compared with those of other species
within the Genbank data bank at the National Center
for Biomedical Information.
Additionally, we examined DNA quality by
performing PCR for canine actin on each sample,
to ensure sufficient template for the microfilarial
PCR.
2.5. Microfilarial phenotyping
In addition to the PCR genotyping, we performed a
modified Knott’s test and wet smear on the blood
submitted from Cases 1–3 and the positive control
samples. The morphological diagnosis for the six
clinical cases is the preliminary morphological
diagnosis provided with initial sample submission
and not the diagnosis obtained by one of the authors
(MFF).
3. Results
All blood-derived DNA samples were positive for
canine actin (not shown), indicating adequate DNA
extraction.
Primers specific for D. immitis (D.imm-F1 and
D.imm-R1), A. reconditum (A.rec-F1 and A.rec-R1)
and D. repens (D.rep-F1 and D.rep-R1) amplified the
expected products only from D. immitis (302 bp), A.
reconditum (348 bp) and D. repens (247 bp, 153 bp)
DNA, respectively (Fig. 1). The doublet amplicon
obtained with primers specific for D. repens has been
previously reported (Favia et al., 2000). There was
marginal amplification of a product from A. reconditum DNA (247 bp) with D. repens-specific primers.
No filariae-specific primer pairs amplified any
products from DNA of the normal dog (negative
control), or from DNA extracted from B. pahangi,
B. malayi or A. dracunculoides (data not shown).
These results confirmed the genotyping of D. immitis,
A. reconditum and D. repens for subsequent analysis
and the specificity of the primer sets.
The pan-filarial primer pair (DIDR-F1 and DIDRR1) amplified the expected product from D. immitis
DNA (542 bp), and A. reconditum DNA (578 bp)
(Fig. 2). Unexpectedly, this primer pair also amplified
Fig. 1. Gel electrophoresis of filarial PCR products in a 1.5%
agarose gel using filarial-specific ITS2-region primers. Lanes 1,
6, 11, 16: MW marker (KB Plus); lanes 2, 7, 12: D. immitis DNA;
lanes 3, 8, 13: A. reconditum DNA; lanes 4, 9, 14: D. repens (Case 1)
DNA; lanes 5, 10, 15: C. familiaris DNA (negative control). Lanes
2–5: D.immF1/D.immR1 primer pair; lanes 7–10: A.recF1/A.recR1
primer pair; lanes 12–15: D.repF1/D.repR1 primer pair.
a 484 base-pair product from the DNA extracted from
Case 1 (D. repens-infested dog). The different sizes of
the amplicons for all three filarial species allowed us to
use this pan-filarial primer set to genotype the filarial
species in Cases 2–6.
Filaria-specific primers for COI demonstrated
specificity for each of the filarial species (Fig. 3),
further confirming the genotyping of microfilariae
from Case 1 as D. repens and validating the panfilarial primer set results.
Results of PCR with pan-filarial primers from
Cases 2–4 and 6 identified the microfilariae as D.
immitis (Fig. 4) (confirmed with species-specific COI
PCR); PCR from Case 5 identified the microfilariae as
A. reconditum (also confirmed with species-specific
COI PCR).
Fig. 2. Gel electrophoresis of filarial PCR products in a 1.5%
agarose gel using pan-filarial ITS2-region primers. Lanes 1, 4:
MW marker (KB Plus); lane 2: A. reconditum DNA; lane 3: D.
immitis DNA; lane 3: D. repens (Case 1) DNA.
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Fig. 3. Gel electrophoresis of filarial PCR products in a 1.5%
agarose gel using filarial-specific COI primers. Lanes 1, 5, 9, 13:
MW marker (KB Plus); lanes 2, 6, 10: D. immitis DNA; lanes 3, 7,
11: A. reconditum DNA; lanes 4, 8, 12: D. repens (Case 1) DNA.
Sequencing of the two amplicons obtained with the
D. repens-specific ITS2-region primers (D.rep-F1,
D.rep-R1) unequivocally confirmed the presumptive
diagnosis of D. repens microfilaremia in Case 1.
Sequencing of the 484 bp amplicon obtained with the
pan-filarial primers identified a novel sequence that
Fig. 4. Gel electrophoresis of filarial PCR products in a 1.5%
agarose gel using pan-filarial ITS2-region primers. Lanes 1, 6:
MW marker (KB Plus); lane 2: Case 4 DNA; lane 3: D. immitis
DNA; lane 4: A. reconditum DNA; lane 5: D. repens (Case 1) DNA.
had high homology with the 5.8S and 28S ribosomal
RNA sequences from D. immitis (present at the 50 and
30 termini of the amplicon), but only moderate
homology with the central D. immitis ITS2 region
(Fig. 5), and no appreciable homology with A.
reconditum ITS2 region. This sequence was consequently labeled as the ITS2 region of D. repens,
submitted to the Genbank data bank at the National
Center for Biomedical Information and assigned the
accession number AY693808.
Morphologic examination of the microfilariae from
Case 1 using a modified Knott’s technique, showed a
lack of a cephalic hook, a slightly tapering anterior
end, and no consistent tail bend (Fig. 6). The
concentration of microfilariae (abundant) and the
motility of live microfilariae (not propulsive) on a wet
smear, were most consistent with a diagnosis of D.
repens. Examination of the microfilariae from Cases 2
and 3 by one of the authors (MFF) supported our
molecular diagnosis of D. immitis.
Results of PCR from B. malayi, B. pahangi and A.
dracunculoides DNA, using the pan-filarial primers,
produced bands at 615 bp (B. malayi), 664 bp (B.
pahangi) and 584 bp (A. dracunculoides) (data not
shown). Sequencing of these amplicons revealed
unique sequences, which were subsequently submitted
to Genbank and assigned accession numbers
AY988599 (B. malayi), AY988600 (B. pahangi) and
DQ018785 (A. dracunculoides).
While PCR with pan-filarial primers failed to
discriminate between A. reconditum and A. dracunculoides, restriction fragment length polymorphism
PCR (RFLP-PCR) using MseII resulted in two
fragments (251 bp and 333 bp) only in A. dracunculoides (data not shown) allowing discrimination
between these two Acanthocheilonema species.
Sequence comparison of the ITS2 region amplified
by our pan-filarial primers (DIDR-F1 and DIDR-R1)
and previously published filarial sequences of this
region demonstrated 77–86% similarity between
species of the same genus (e.g. D. immitis and D.
repens showed 77% similarity, all three Brugia species
showed 86% similarity), but only 30–60% similarity
between species of different genera (e.g. B. pahangi
and A. reconditum showed only 30% similarity).
Phylogenetic tree analysis also grouped sequences
most closely by genus, i.e., species of the same genus
grouped together.
M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314
Fig. 5. Nucleotide sequence alignment of 5.8S-ITS2-28S ribosomal DNA region of seven filariae. The 5.8S and 28S ribosomal DNA are identified above the 50 and 30 ends of the
sequences; shaded sequences within the majority sequence are primer binding sequences for the pan-filarial primers.
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Fig. 5. (Continued ).
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Fig. 6. Modified Knott’s stain of microfilaria from Case 1 (D. repens). The head is slightly tapered, there is no cephalic hook, and the tail is
straight, consistent with Dirofilaria repens. Magnification 100.
4. Discussion
We initiated this study to resolve a discordant
diagnosis of filariasis in a dog, and successfully
identified a D. repens infestation in a dog in the United
States by PCR genotyping of microfilarial DNA
extracted from the patient’s blood. We also resolved
discordant diagnoses provided by standard morphological examination of the microfilariae (Knott’s test).
Unexpectedly, the pan-filarial primers we designed
(DIDR-F1 and DIDR-R1), based on sequence homology of D. immitis and A. reconditum, amplified not only
the expected 542 and 578 bp products from these
filariae, but also a 484 bp product from D. repens, that
was present in the Case 1 of this study, a 584 bp product
from A. dracunculoides, a 615 bp product from B.
malayi, and a 664 bp product from B. pahangi. This
permitted a single PCR reaction to differentiate
between D. immitis and five other filariae found in
dogs, removing the need to perform multiple speciesspecific genotyping reactions. We subsequently identified microfilariae from an additional four dogs as D.
immitis and as A. reconditum in one dog with this panfilarial primer set. Our molecular identification was
concordant with independent morphological phenotyping of the microfilariae by an experienced parasitologist
(MFF), but discordant with the initial morphological
diagnosis in 3/5 cases, illustrating the inaccuracy of
morphological diagnosis by minimally trained technicians. Finally, we were able to provide additional
genotyping information for several filariae that usually
require specialized diagnostic interpretation.
Previous investigators have utilized PCR technology to diagnose filarial infestations in dogs and
humans (Baneth et al., 2002; Favia et al., 1996, 1997;
Mar et al., 2002); however, in each of these studies,
only two filarial species were compared. We designed
our pan-filarial primers from previously submitted
sequences, and optimized them for efficient amplification.
The need to correctly identify the filarial species is
becoming relevant throughout the world, because of
the increased frequency of transportation of dogs
between continents and countries (Genchi, 2003;
Zahler et al., 1997). While D. repens has been
considered an exotic parasite in some parts of the
world, such as the United States and Australia, and
consequently not considered in the differential
diagnosis of canine filariasis in these continents, this
case illustrates the potential diagnostic complications
that can arise, and provides a simple molecular
strategy for correctly identifying the filaroid. The
potential mosquito vectors of D. repens—Aedes
aegypti (Cancrini et al., 2003), Ae. albopictus (Anyanwu et al., 2000) and Ae. vexans—are enzootic in the
United States and Ae. vexans is enzootic in the region
of New York State where this dog currently lives
(Nassau County Department of Health, 2003).
Similarly, Ae. aegypti is enzootic in Australia; Ae.
albopictus is not found in Australia. Thus, the
potential exists for spread of D. repens in these
regions. Whether the strains in the United States or
Australia can incubate larvae and transmit the
infestations is unknown. Interestingly, Ae. aegypti is
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relatively resistant to infestation with D. immitis, and
is considered an inefficient intermediate host for
transmission of heartworm disease (Apperson et al.,
1989).
The geographic region in which Case 1 acquired
the infection is unknown. Dirofilaria repens has not
been reported in The Netherlands, but has been
identified in the Czech Republic (where this dog was
born), Hungary, Italy, Greece and France (Cordonnier
et al., 2002; Elek et al., 2000; Pampiglione et al., 2001;
Vakalis and Himonas, 1997; Vasilkova et al., 1992).
The microfilariae from Case 2 were initially
incorrectly identified as A. reconditum, partly based
on a negative antigen test, while the microfilariae in
Case 3 had been identified correctly as D. immitis. In
Cases 2 and 3, examination by an experienced
parasitologist (MFF) correctly identified the microfilariae as D. immitis. Causes for antigen-negative
D. immitis microfilaremias include prior adulticide
therapy without subsequent microfilaricidal therapy or
natural death of adult heartworm with persistence of
microfilariae (which have a lifespan of approximately
2 years) (Otto, 1977). Additionally, at least one
product insert (Dirocheck Canine Heartworm Antigen
Test Kit Product Insert, Synbiotics, San Diego, CA)
also lists subthreshold antigenemia, immune clearance
of antigen–antibody complexes, false positive microfilarial results (due to contamination of reagents used
for sample preparation), transfusions from microfilaremic dogs, or destruction of antigen due to
improper storage as potential causes of discordant
results. While a Dirofilaria microfilaremia does not
necessarily endanger the health of the infested patient,
the patient can act as a reservoir for further infestation
of other dogs if the intermediate mosquito host is
present. Microfilaricidal therapy can resolve this
situation.
The microfilaremia in Case 3 was initially
identified during a routine urinalysis for hematuria
(microfilaruria). We believe this is the first reported
case of microfilaruria in a dog. A prior report exists of
microfilaruria in a cat (Beaufils et al., 1991).
We did not test the discriminatory ability of our
pan-filarial primer set against all filariae that are
known to infest dogs or cats (Genchi, 2003). At least
one additional ‘‘Dipetalonema-like’’ filaria exists in
Europe and Africa—Cercopithifilaria (Acanthocheilonema) grassi—but has not been reported in the
United States. It is possible that our pan-filarial primer
set will fail to discriminate this filaria from D. immitis,
however, D. immitis-specific primer sets should
correctly genotype the filarial species in question if
this becomes necessary. Further studies are required to
validate the pan-filarial primers against C. grassi.
Additionally, reports exist of novel filariae in dogs in
Ireland and the United States (Jackson, 1975; Pacheco
and Tulloch, 1970). Techniques such as those
described in this report might be useful in genotyping
such filariae.
The B. malayi sequence we obtained from adult
filariae showed only 86% similarity with a published
B. malayi sequence (Genbank # AF499130), comparable to the degree of similarity between our B. pahangi
sequence and the published B. malayi sequence
(AF499130), or between our B pahangi and B. malayi
sequences. This differed substantially from the
similarity between the published sequences for B.
malayi (AF499130) and B. timori (AF499132), which
the authors claimed were 99.5% similar, with only 4
base-pair substitutions along the 600 base-pair region
(Fischer et al., 2002). Several reasons exist for these
discrepant results. Fischer et al claimed that this
degree of similarity was unique to these two species,
however, they did not examine DNA from adult
Brugia species, but only from microfilaria, and by
supposition based on epidemiological data. Thus, their
initial phenotypic classification, which they used as a
gold standard for that study, could be erroneous. We
could find no study by these authors that confirmed
their results using adult B. malayi and B. timori. Given
the much lower degree of similarity between species
of a single genus detailed in this report (e.g. D. immitis
and D. repens, B. malayi and B. pahangi, or A.
reconditum and A. dracunculoides all showed intragenus, interspecies variation of approximately 20%,
similar to the difference we observed between B.
malayi and the previously published B. timori), we
suspect that the sequence for B. malayi (AF499130)
published by Fischer et al. (2002) is incorrect and is in
fact the sequence for B timori. Examination of the
data presented by these authors indicates a broad
genetic variation of the Hha1 tandem repeat in these
species, which also suggests a misclassification of
microfilariae.
A less plausible alternative is that inter-species
variation of the ITS2 region between B. malayi and B.
M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314
timori (<0.5%) is less than the intra-species variation
of B. malayi from different regions (>20%). Several
studies of ITS2 regions in various helminths suggest
that while intra-species variation can occur, it is less
than inter-species variation, and that in most cases
intra-species variation is limited to single nucleotide
polymorphisms (Conole et al., 1999; Gasser et al.,
1999a,b; Huby-Chilton et al., 2001; Newton et al.,
1998; Woods et al., 2000) and previous studies of other
filariae show similar results (Gasser et al., 1996).
These findings agree with our observations of ITS2
variability in filariae and support our genotypic
identification of B. malayi. Finally, it is possible that
the sample of adult B. malayi provided for our study
was incorrectly identified; however, phenotypic
classification of filariae is generally more robust
using adult filariae than microfilariae. The reason for
the discrepancy between our data and that of Fischer
et al. (2002) remains to be elucidated.
Finally, we examined the ability of our pan-filarial
primer sequences to amplify the 5.8S-ITS2-28S
region from two additional filariae in silico. Analysis
of published sequences for Onchocerca volvulus
(Genbank # AF228575), and Mansonella ozzardi
(Genbank # AF228554) demonstrated that our panfilarial primers should amplify 470 and 430 bp
amplicons, respectively. This suggests that our primer
set might be uniquely capable of amplifying DNA that
can be used to genotype all filariae of the family
Onchocercidae.
We recommend using the pan-filarial ITS2-region
DIDR-F1 and DIDR-R1 primers with DNA from
D. immitis (as a positive control and amplicon size
standard) against which to compare amplicons from
clinical cases of microfilaremia that test negative for
D. immitis antigen. Amplicons approximately 40 basepairs larger than the D. immitis amplicon (542 basepairs) are diagnosed as A. reconditum (578 base-pairs)
or A. dracunculoides (584 base-pairs), while those
approximately 60 base-pairs smaller than D. immitis
are diagnosed as D. repens (484 base-pairs). Discrimination of Brugia species by comparing amplicons is limited because B. malayi (615 base-pairs) is
similar to B. timori (625 base-pairs), but these species
should be easily differentiated from B. pahangi (664
base-pairs). However, PCR-RFLP will discriminate B.
malayi and B. timori. If further confirmation is required, either species-specific PCR can be performed,
313
using the published primer pairs from this and other
studies, or the amplicons can be sequenced, however,
we feel that simple gel electrophoresis should be
sufficient for diagnosis in most cases. Additionally, we
feel that clinicians and parasitologists should be
alerted to the possibility that D. repens infestation
might be an emerging condition with zoonotic
potential in the United States.
Acknowledgements
The authors would like to thank Dwight Bowman,
Susan Wade and Stephanie Schaaf for their technical
insights and providing D. immitis positive control
samples; Andrew Fei and Fabiano Montiani Ferreira
for assistance in obtaining A. reconditum samples;
Thom Klei for generously providing Brugia filaria;
Francisco Alonso de Vega for generously providing
A. dracunculoides samples and Veterinary Information Network for providing access to the case that
initiated this study.
References
Anyanwu, I.N., Agbede, R.I., Ajanusi, O.J., Umoh, J.U., Ibrahim,
N.D., 2000. The incrimination of Aedes (stegomyia) aegypti as
the vector of Dirofilaria repens in Nigeria. Vet. Parasitol. 92,
319–327.
Apperson, C.S., Engber, B., Levine, J.F., 1989. Relative suitability
of Aedes albopictus and Aedes aegypti in North Carolina to
support development of Dirofilaria immitis. J. Am. Mosq.
Control Assoc. 5, 377–382.
Baneth, G., Volansky, Z., Anug, Y., Favia, G., Bain, O., Goldstein,
R.E., Harrus, S., 2002. Dirofilaria repens infection in a dog:
diagnosis and treatment with melarsomine and doramectin. Vet.
Parasitol. 105, 173–178.
Beaufils, J.P., Martin-Granel, J., Bertrand, F., 1991. Présance de
microfilaires de Dirofilaria immitis dans les urines d’un chat
occlus. Prat. Med. Chir. An. Com. 26, 467–472 in French.
Bredal, W.P., Gjerde, B., Eberhard, M.L., Aleksandersen, M.,
Wilhelmsen, D.K., Mansfield, L.S., 1998. Adult Dirofilaria
repens in a subcutaneous granuloma on the chest of a dog. J.
Small Anim. Pract. 39, 595–597.
Cancrini, G., Romi, R., Gabrielli, S., Toma, L., Di Paolo, M.,
Scaramozzino, P., 2003. First finding of Dirofilaria repens in
a natural population of Aedes albopictus. Med. Vet. Entomol. 17,
448–451.
Chalifoux, L., Hunt, R.D., 1971. Histochemical differentiation of
Dirofilaria immitis and Dipetalonema reconditum. J. Am. Vet.
Med. Assoc. 158, 601–605.
314
M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314
Chandrasekharan, N.V., Karunanayake, E.H., Franzen, L., Abeyewickreme, W., Pettersson, U., 1994. Dirofilaria repens: cloning
and characterization of a repeated DNA sequence for the diagnosis of dirofilariasis in dogs, Canis familiaris. Exp. Parasitol.
78, 279–286.
Conole, J.C., Chilton, N.B., Jarvis, T., Gasser, R.B., 1999. Intraspecific and interspecific variation in the second internal transcribed spacer (ITS-2) sequence for Metastrongylus (Nematoda:
Metastrongyloidea) detected by high resolution PCR-RFLP. Int.
J. Parasitol. 29, 1935–1940.
Cordonnier, C., Chatelain, D., Nevez, G., Sevestre, H., Gontier,
M.F., Raccurt, C.P., 2002. Difficulties in diagnosis of human
dirofilariasis outside known enzootic areas. Rev. Med. Interne
23, 71–76.
Courtney, C.H., Zeng, Q.Y., MacKinnon, B.R., 1993. Comparison of
two antigen tests and the modified Knott’s test for detection of
canine heartworm at different worm burdens. Canine Pract. 18,
5–7.
Elek, G., Minik, K., Pajor, L., Parlagi, G., Varga, I., Vetesi, F.,
Zombori, J., 2000. New human Dirofilarioses in Hungary.
Pathol. Oncol. Res. 6, 141–145.
Favia, G., Lanfrancotti, A., Della Torre, A., Cancrini, G., Coluzzi,
M., 1996. Polymerase chain reaction-identification of Dirofilaria repens and Dirofilaria immitis. Parasitology 113 (Pt. 6),
567–571.
Favia, G., Lanfrancotti, A., della Torre, A., Cancrini, G., Coluzzi,
M., 1997. Advances in the identification of Dirofilaria repens
and Dirofilaria immitis by a PCR-based approach. Parassitologia
39, 401–402.
Favia, G., Bazzocchi, C., Cancrini, G., Genchi, C., Bandi, C., 2000.
Unusual organization of the 5S ribosomal spacer in Dirofilaria
repens: absence of a canonical spliced leader 1 sequence.
Parasitol. Res. 86, 497–499.
Fischer, P., Wibowo, H., Pischke, S., Ruckert, P., Liebau, E., Ismid,
I.S., Supali, T., 2002. PCR-based detection and identification of
the filarial parasite Brugia timori from Alor Island, Indonesia.
Ann. Trop. Med. Parasitol. 96, 809–821.
Gasser, R.B., LeGoff, L., Petit, G., Bain, O., 1996. Rapid delineation
of closely-related filarial parasites using genetic markers in
spacer rDNA. Acta Trop. 62, 143–150.
Gasser, R.B., Rossi, L., Zhu, X., 1999a. Identification of Nematodirus
species (Nematoda: Molineidae) from wild ruminants in Italy
using ribosomal DNA markers. Int. J. Parasitol. 29, 1809–1817.
Gasser, R.B., Woods, W.G., Huffman, M.A., Blotkamp, J., Polderman, A.M., 1999b. Molecular separation of Oesophagostomum
stephanostomum and Oesophagostomum bifurcum (Nematoda:
Strongyloidea) from non-human primates. Int. J. Parasitol. 29,
1087–1091.
Genchi, C., 2003. Epidemiology and distribution of Dirofilaria
and dirofilariosis in Europe: state of the art. In: Proceedings
of the 2003 Helminthological Colloquium, Vienna, Austria, pp.
6–11., http://www.vu-wien.ac.at/i116/oegtp/downloads_oegtp/
Abstractband.pdf.
Huby-Chilton, F., Beveridge, I., Gasser, R.B., Chilton, N.B., 2001.
Single-strand conformation polymorphism analysis of genetic
variation in Labiostrongylus longispicularis from kangaroos.
Electrophoresis 22, 1925–1929.
Jackson, R.F., 1975. A filarial parasite in greyhounds in Southern
Ireland. Vet. Rec. 97, 476–477.
Knott, J., 1935. The periodicity of the mf of Wuchereia bancrofti,
preliminary report of some injection experiments. Trans. Roy.
Soc. Trop. Med. Hyg. 29, 59–64.
Manuali, E., Eleni, C., Giovannini, P., Costarelli, S., Ciorba, A.,
2005. Unusual finding in a nipple discharge of a female dog:
dirofilariasis of the breast. Diagn. Cytopathol. 32, 108–
109.
Mar, P.H., Yang, I.C., Chang, G.N., Fei, A.C., 2002. Specific
polymerase chain reaction for differential diagnosis of Dirofilaria immitis and Dipetalonema reconditum using primers
derived from internal transcribed spacer region 2 (ITS2). Vet.
Parasitol. 106, 243–252.
Nassau County Department of Health, 2003. Nassau County Mosquito Surveillance and Control Report.
Newton, L.A., Chilton, N.B., Beveridge, I., Gasser, R.B., 1998.
Systematic relationships of some members of the genera Oesophagostomum and Chabertia (Nematoda: Chabertiidae) based
on ribosomal DNA sequence data. Int. J. Parasitol. 28, 1781–
1789.
Otto, G.F., 1977. The significance of microfilaremia in the diagnosis
of heartworm infection. In: Proceedings of the American
Heartworm Society Heartworm Symposium. pp. 22–30.
Pacheco, G., Tulloch, G.S., 1970. Microfilariae of Dirofilaria striata
in a dog. J. Parasitol. 56, 248.
Pampiglione, S., Rivasi, F., Angeli, G., Boldorini, R., Incensati,
R.M., Pastormerlo, M., Pavesi, M., Ramponi, A., 2001. Dirofilariasis due to Dirofilaria repens in Italy, an emergent zoonosis:
report of 60 new cases. Histopathology 38, 344–354.
Pampiglione, S., Rivasi, F., Paolino, S., 1996. Human pulmonary
dirofilariasis. Histopathology 29, 69–72.
Romano, A., Sachs, R., Lengy, J., 1976. Human ocular dirofilariasis
in Israel. Isr. J. Med. Sci. 12, 208–214.
Sambrook, J., Russell, D.W. (Eds.), 2001. Molecular Cloning: A
Laboratory Manual, 3rd ed., vol. 1. Cold Spring Harbor Laboratory Press, Cold Spring Harbor.
Tarello, W., 2003. Dermatitis associated with Dirofilaria repens
microfilariae in three dogs in Saudi Arabia. J. Small Anim. Pract.
44, 132–134.
Vakalis, N., Spanakos, G., Patsoula, E., Vamvakopoulos, N.C., 1999.
Improved detection of Dirofilaria repens DNA by direct polymerase chain reaction. Parasitol. Int. 48, 145–150.
Vakalis, N.C., Himonas, C.A., 1997. Human and canine dirofilariasis in Greece. Parassitologia 39, 389–391.
Vasilkova, D., Klisenbauer, D., Juhas, T., Moravec, F., Uhlikova, M.,
Hubner, J., Konakova, G., 1992. Isolation of Dirofilaria repens
in vitreoretinal findings. Cesk. Oftalmol. 48, 274–277.
Woods, W.G., Whithear, K.G., Richards, D.G., Anderson, G.R.,
Jorgensen, W.K., Gasser, R.B., 2000. Single-strand restriction
fragment length polymorphism analysis of the second internal
transcribed spacer (ribosomal DNA) for six species of Eimeria
from chickens in Australia. Int. J. Parasitol. 30, 1019–
1023.
Zahler, M., Glaser, B., Gothe, R., 1997. Imported parasites in dogs:
Dirofilaria repens and Dipetalonema reconditum. Tierarztl.
Prax. 25, 388–392.