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: [email protected] (M. Rishniw). 0304-4017/$ – see front matter # 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2005.10.013 304 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 305 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. 306 M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314 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. 308 M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314 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. 309 M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314 Fig. 5. (Continued ). 310 M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314 311 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 312 M. Rishniw et al. / Veterinary Parasitology 135 (2006) 303–314 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.
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