MATERIALS AND METHODS

Ethical approval

The present study involved the non-invasive collection of fecal samples and rectal swabs from companion cats and dogs presented to veterinary clinics, rescue organizations, and animal shelters. Sample collection was performed by licensed veterinarians following standard veterinary clinical procedures to minimize animal stress and discomfort. Written informed consent was obtained from all animal owners or authorized caretakers before sample collection. No animals were subjected to experimental infection, invasive procedures, or harmful interventions specifically for the purpose of this study. All sampling procedures complied with institutional guidelines for animal welfare and ethical research involving animals. The research does not require approval from the Institutional Animal Ethics Committee of the College of Veterinary Science and Animal Husbandry, Kamdhenu University, Anand, Gujarat, India.

Study period and location

This cross-sectional study was conducted between September 2024 and September 2025 in and around Anand District, Gujarat, India. Laboratory investigations, including molecular detection, virus isolation, sequencing, and phylogenetic analyses, were performed at the Department of Veterinary Biotechnology, College of Veterinary Science and Animal Husbandry, Kamdhenu University, Anand, Gujarat, India.

Study design

A cross-sectional epidemiological study was designed to determine the prevalence, molecular characteristics, and cross-species circulation of FPV and CPV among companion animals. The study further compared the diagnostic performance of RAT, conventional PCR, and qPCR and evaluated the molecular diversity of circulating viruses through VP2 gene sequencing and phylogenetic analysis.

Sample collection

A total of 200 fecal samples were collected from 100 cats and 100 dogs presented to the Veterinary Clinical Complex, College of Veterinary Science and Animal Husbandry, Anand, Gujarat, India, as well as from private veterinary clinics, rescue organizations, and animal shelters in the surrounding region. Samples were obtained from animals exhibiting clinical signs suggestive of gastroenteritis, including diarrhea, vomiting, dehydration, anorexia, and abdominal discomfort, as well as from apparently healthy animals for comparative analysis.

Freshly voided feces or rectal swabs were collected aseptically using sterile screw-cap swabs (HiMedia Laboratories Pvt. Ltd., Mumbai, India). Swabs were either collected dry or pre-moistened with Hanks' Balanced Salt Solution to maintain viral stability. Each sample was assigned a unique identification code according to the host species. Samples were transported to the laboratory on ice in insulated containers and stored at 4°C for short-term processing or at −20°C for long-term storage until further analysis. Apparently healthy animals were defined as those not exhibiting clinical signs of gastrointestinal disease at the time of sampling.

DNA extraction

Viral DNA was extracted using the QIAamp DNA Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer's instructions. Briefly, 200 µL of a 10% (w/v) fecal suspension prepared in phosphate-buffered saline (pH 7.4) was used for extraction. Samples were lysed using proteinase K and Buffer AL, followed by column-based purification. DNA was eluted in 50 µL of elution buffer and stored at −20°C until analysis.

DNA concentration and purity were determined using a NanoDrop spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Extraction controls were included throughout the procedure to monitor potential contamination and ensure extraction quality. Samples with A260/A280 ratios between 1.8 and 2.0 were considered acceptable for downstream molecular analyses.

PCR detection of FPV and CPV

Conventional PCR targeting a conserved region of the VP2 gene was performed using primers described by Carreno et al. [12]. Owing to the high sequence similarity between FPV and CPV, these primers amplified parvoviral DNA from both feline and canine samples; however, differentiation between the two viruses was not possible using conventional PCR alone.

Each PCR reaction was prepared in a total volume of 25 µL containing 12.5 µL of 2× PCR Master Mix (Takara Bio Inc., Kusatsu, Japan), 10 pmol of each primer, 2 µL of template DNA, and nuclease-free water to the final reaction volume.

Thermal cycling conditions consisted of an initial denaturation at 95°C for 5 min, followed by 35 cycles of denaturation at 95°C for 30 s, annealing at 55°C for 30 s, and extension at 72°C for 1 min, with a final extension at 72°C for 10 min.

PCR products were resolved on a 1% agarose gel prepared in Tris-borate-EDTA buffer at 90 V for 50 min, stained with ethidium bromide, and visualized under ultraviolet illumination using a gel documentation system (Bio-Rad Laboratories, Hercules, CA, USA).

Positive controls, negative controls, and no-template controls (NTC) were included in every PCR run to ensure assay reliability. Previously validated primers targeting conserved regions of the VP2 gene were used to ensure assay specificity. The primer and probe sequences used in this study are presented in Table 1.

Quantitative PCR for virus differentiation

The duplex probe-based qPCR assay was performed as described by Decaro et al. [13] for the simultaneous detection and differentiation of FPV and CPV. The assay targeted the VP2 gene using virus-specific hydrolysis probes labeled with FAM for CPV and VIC for FPV.

Each reaction was prepared in a final volume of 20 µL containing 10 µL of 2× TaqMan™ Universal PCR Master Mix (Thermo Fisher Scientific), 0.4 µM of each primer, 0.2 µM of each probe, and 2 µL of template DNA.

Amplification was carried out using a QuantStudio Real-Time PCR System (Thermo Fisher Scientific) under the following cycling conditions: initial activation at 95°C for 10 min, followed by 40 cycles of denaturation at 95°C for 15 s and annealing/extension at 50°C for 60 s.

Fluorescence signals were recorded during each amplification cycle, and cycle threshold (Ct) values ≤35 were considered positive. All reactions were performed in duplicate and included positive controls, negative controls, and NTCs. Assay performance was evaluated based on consistent amplification of positive controls and the absence of amplification in negative and NTC. Primer and probe details are provided in Table 1.

Target virus | Primer/probe | Sequence (5′–3′) | Assay type | Amplicon size (bp) | Reference
FPV (VP2) | FPV F | CAGGAAGATATCCAGAAGGA | PCR | 681 | [12]
 | FPV R | GGTGCTAGTTGATATGTAATAAACA |  |  | 
FPV/CPV | FPV/CPV-F | ACAAGATAAAAGACGTGGTGTAACTCAA | Probe-based qPCR | 83 | [13]
 | FPV/CPV-R | CAACCTCAGCTGGTCTCATAATAGT |  |  | 
 | FPV probe | VIC-ATGGGAAATACAGACTATAT-BHQ |  |  | 
 | CPV probe | FAM-ATGGGAAATACAAACTATAT-BHQ |  |  | 

Virus isolation in cell culture

Virus isolation was attempted using the Crandell-Rees feline kidney (CRFK) cell line obtained from the Indian Veterinary Research Institute, Bengaluru, India. Cells were maintained in Dulbecco's Modified Eagle Medium supplemented with 10% fetal bovine serum and 1% antibiotic mixture containing penicillin (100 U/mL), streptomycin (100 µg/mL), and amphotericin B (0.25 µg/mL). Cell cultures were incubated at 37°C and routinely monitored to confirm the absence of mycoplasma contamination.

Before inoculation, fecal supernatants were filtered through 0.22-µm syringe filters to remove bacterial contaminants. Approximately 200 µL of clarified inoculum was added to each well. Blind passages were performed every 3–4 days by transferring 200 µL of culture supernatant onto fresh CRFK monolayers. All procedures were conducted under biosafety level 2 conditions.

Following adsorption at 37°C for 1 h with intermittent rocking, maintenance medium containing 2% fetal bovine serum was added. Cell cultures were examined daily for cytopathic effects (CPE) for up to 7 days. Samples that failed to produce CPE were subjected to up to 3 blind passages to improve viral recovery.

Sequencing and molecular characterization

PCR products were purified using a commercial PCR purification kit and subjected to Sanger sequencing using an ABI 3730 Genetic Analyzer (Applied Biosystems, Foster City, CA, USA). Sequencing was performed with forward primers, and sequence quality was assessed prior to downstream analyses.

Sequence editing and assembly were performed using BioEdit software. Similarity searches were conducted using the Basic Local Alignment Search Tool available through the National Center for Biotechnology Information (NCBI).

Phylogenetic analysis was performed using Molecular Evolutionary Genetics Analysis version 12 software. The evolutionary history was inferred using the Maximum Likelihood method based on the Tamura three-parameter model with 1,000 bootstrap replicates. Because only partial VP2 gene sequences were obtained, complete genetic characterization and definitive subtype differentiation of the circulating strains were limited.

Statistical analysis

Data generated in the present study were analyzed using descriptive statistics. The prevalence of FPV and CPV infections was calculated as the percentage of positive samples among all samples examined. Because of the observational nature of the study and the absence of complete epidemiological data for all investigated variables, advanced statistical analyses to determine associations or risk factors were not performed.