Abstract
Epidemiological surveillance of porcine group A rotavirus (RVA) strains was conducted in five swine herds in Ohio using historical (2004) and recent (2011 to 2012) fecal samples. Of the 371 samples examined, 9.4% (35/371) were positive for RVA. The RVA detection rates increased from 5.9% in 2004 and 8.5% in 2011 to 13.8% in 2012. A total of 23 positive samples were analyzed for RVA G and P genotypes. The dominant G-P combination was G9P[13] found in 60.9% of positive samples. The other combinations were G9P[7] (8.7%), G4P[13] (8.7%), G11P[13] (4.3%), and G11P[7] (4.3%). Sequence analysis of partial VP7 genes of selected strains revealed that the G4 strains were closely related to one another (95%) and, to a lesser extent, to human (82 to 84%) and porcine (84 to 86%) G4 strains. The G11 strains detected shared identical VP7 gene sequences (100%) and were closely related to human (85 to 86%) and other porcine (83%) G11 strains. The G9 strains identified were closely related to one another and to human and other porcine strains (96 to 97%, 89 to 91%, and 89 to 91% nucleotide identities, respectively). The VP4 gene analysis revealed that P[7] strains were closely related to each other and to P[7] strains isolated from porcine, bovine, and panda samples (91 to 99%, 92 to 99% and 92 to 99%, respectively). The P[13] strains showed a higher diversity among themselves and with other porcine P[13] strains, ranging from 83% to 99% and from 82 to 97%, respectively. Our results demonstrate broad genetic heterogeneity of the RVA strains and suggest the possibility of genetic reassortment between different RVA genotypes within these farms.
INTRODUCTION
Gastroenteritis in pigs is a multifactorial and multietiological disease. Management differences may play an important role in development of the disease, in addition to coinfection by microorganisms such as bacteria, protozoa, and viruses (1). Viral diseases of swine, such as coronavirus (transmissible gastroenteritis virus [TGEV] and porcine epidemic diarrhea virus [PEDV])], circovirus type 2 (PCV2), rotavirus (RV), swine influenza virus, and porcine respiratory and reproductive syndrome virus (PRRSV) infections, represent continuing challenges for the swine industry, increasing costs of treatment and prevention.
Rotaviruses are a major cause of viral gastroenteritis in children and young animals worldwide. RV infections cannot be diagnosed solely based on clinical signs because they do not differ from those caused by other intestinal pathogens. Laboratory tests are necessary for reliable RV diagnosis and surveillance and can be useful in clinical settings to avoid inappropriate use of antibiotics that cannot prevent viral diarrheas. Large numbers of excreted viral particles during RV infection, a wide range of susceptible species, resistance to environmental conditions and disinfectants, and the possibility of concurrent infections with different RV strains leading to new virus strains, all contribute to RV prevalence and genetic diversity (2–4).
Rotavirus is a genus of the Reoviridae family of double-stranded RNA (dsRNA) viruses, having a genome of 11 segments of dsRNA encoding six structural (VP1 to VP4, VP6, and VP7) and six nonstructural viral proteins (NSP1 to NSP5/6). The RVs are classified into seven groups (Gp), A to G, based on antigenic relationships of their VP6 protein (5). Groups A, B, and C RVs (RVAs, RVBs, and RVCs, respectively) infect humans and animals, with RVA usually in highest prevalence (2). Based on differences in the outer capsid VP7 and VP4 proteins that contain epitopes that independently induce neutralizing antibodies, RVs are classified into G and P types, respectively. Within RVAs there are 27 G genotypes and 35 P genotypes identified to date (6). Twelve G genotypes (G1 to G6, G8 to G12, and G26) and 13 P genotypes (P[1], P[5] to P[8], P[11], P[13], P[19], P[23], P[26], P[27], P[32], and P[34]) have been associated with pigs (3, 6–8). However, G3, G4, G5, and G11 are the most common G genotypes identified in swine and are usually associated with P[5], P[6], and P[28]; the others are considered less common genotypes in swine (9–11).
Porcine RVs are important enteric pathogens not only because of their economic impact on swine production in terms of mortality and morbidity but also because they are a potential source of heterologous RV infections in humans and cattle (12, 13). Knowledge of the molecular epidemiology of RVAs is crucial for preventing or reducing the incidence of porcine RVA diarrhea as well as transmission of porcine RVA to humans. Since protection against heterologous strains is lower, understanding the genetic diversity of RVAs is critical for the design of effective prophylactic tools, including vaccine development, optimization, and improvement. In this paper we report the prevalence as well as genetic diversity of RVAs in historic (2004) and recent (2011 and 2012) fecal samples from five swine farms located in the state of Ohio.
MATERIALS AND METHODS
Samples.
A total of 371 fecal samples were collected in 2004 (n = 118), 2011 (n = 130), and 2012 (n = 123) from five selected swine farms in Ohio for detection of RVAs (Table 1). All samples collected in 2004 were from nursing piglets with no information about diarrheal status (14), whereas for the 2011, age information was missing for 39 samples from farm 2, and diarrhea status was not available for 96 samples. However, samples collected in 2012 were from both nursing (n = 76) and weaned (n = 47) piglets, and they were collected from piglets with diarrhea (n = 62) and without diarrhea (n = 61). Fecal samples (feces or swabs) were collected from individual piglets and placed in labeled sterile plastic tubes and transported to the laboratory on ice. Ten percent (wt/vol) fecal suspensions were prepared in minimum essential medium (Life Technologies, NY), vortexed, and centrifuged at 1,800 × g for 30 min at 4°C. The clarified supernatants were used for RNA extraction. None of the farms surveyed used the RVA vaccine (ProSystem Rota; Intervet, Inc./Merck Animal Health) licensed for swine in the United States.
Table 1.
The year and number of samples collected, pig ages, and diarrhea status
| Collection yr (n)a | Farm no. (location)b | Sample collection period (mo) | No. of samples | Diarrhea status | Age (no. of pigs)c |
|---|---|---|---|---|---|
| 2004 (118) | 1 (central) | April | 32 | NAd | <3 wks |
| 2 (NW) | July | 31 | NA | <3 wks | |
| December | 40 | NA | <3 wks | ||
| 3 (central) | May | 15 | NA | <3 wks | |
| 2011 (130) | 2 (NW) | July | 39 | NA | NA |
| December | 20 | NA | <3 wks | ||
| 4 (NE) | June | 37 | NA | <3 wks | |
| 5 (NE) | April | 34 | 5 cases | <3 wks | |
| 2012 (123) | 2 (NW) | January | 8 | 8 cases | <3 wks (4), gilt (4) |
| March | 60 | 6 cases | <3 wks (17), >3 wks (43) | ||
| May | 55 | 50 cases | <3 wks |
n, total number of samples.
Location refers to parts of Ohio. NW, northwest; NE, northeast.
<3 weeks, nursing piglets, >3 weeks, weaned piglets.
NA, not available.
RNA extraction.
RNA was extracted from a 250-μl starting volume of centrifuged 10% fecal suspensions using an RNeasy minikit from Qiagen (Valencia, CA) according to the manufacturer's instructions. The total RNA recovered was suspended in 40 μl of nuclease-free water and stored at −70°C until used.
Detection of RVA.
Conventional reverse transcription-PCR (RT-PCR) was used for detection of the RVAs with validated primer sets, NSP3F (nucleotides [nt] 963 to 982) and NSP3R (nt 1049 to 1034) (Table 2) using Promega reagents according to the manufacturer's instructions (Promega, CA). Prior to reverse transcription, 2 μl of RNA sample was mixed with 0.5 μl of dimethyl sulfoxide (Sigma-Aldrich, MO), heated at 97°C for 5 min, and then snap-chilled on ice to denature the double-stranded segments of the rotavirus genomic RNA. For the RT-PCR the following conditions were applied: incubation for 30 min at 50°C for the RT reaction, a preheating step at 94°C for 5 min for initial denaturation, followed by 40 PCR cycles at 94°C for 15 s, 56°C for 1 min, and 72°C for 30 s, and a final extension for 7 min at 72°C, and then at 4°C for storage until the sample was removed from the machine. The PCR products from the above reaction were analyzed in a 3% agarose gel after staining with EZ-Vision dye (Amresco) and UV light transillumination. The amount of PCR inhibitors in the samples was determined to be negligible based upon results obtained by making 10-fold and 100-fold dilutions of each RNA sample.
Table 2.
Primer sets used for RVA detection and genotyping
| Target or function | Name | Sequence | Region (nt) | Amplicon size (bp) | Reference or source |
|---|---|---|---|---|---|
| Detection primer set | NSP3F | ACCATCTACACATGACCCTC | 963–982 | 87 | 15 |
| NSP3R | GGTCACATAACGCCCC | ||||
| Common VP7 gene | 5′comVP7-F | GGCTTTAAAAGAGAGAATTTC | 1–1062 | 1062 | 16 |
| 3′comVP7-R | GGTCACATCATACAATTCTAA | ||||
| G3 | G3Fdeg | GAYAATTCRTGGAARGATACACTT | 331–782 | 452 | This study |
| G3Rdeg | GTACAAGTRTTTGTYGTCACRYTCA | ||||
| G4a | JA-AV-G4-Fdeg | ACDTTAGGARTYGGATGTCA | 655–1062 | 407 | This study |
| G5a | JA-AV-G5-Rdeg | YTTWKYRTCYGCRATTTC | 1–339 | 339 | This study |
| G9 | G9-VP7-4F | ACAGCATATGCAAATTCGTCACAG | 241–388 | 148 | This study |
| G9-VP7-4R | CAGTTGGCCACCCCTTAGTCA | ||||
| G11 | JA-G11Fdeg | GAYGAYAAGTGGARAGATACTCTC | 331–589 | 259 | This study |
| JA-G11Rdeg | CYTCATYWGTYTGTTGATAATARTAAAGA | ||||
| Universal VP4 gene | 793F | TGGAAAGAAATGCARTAYAA | 793–1583 | 812 | This study |
| 1583R | CCHGARAACATNGARAACATATC | ||||
| Universal VP7 gene | VP7-39F | GCTCYTTTTRATGTATGGTATTGAATATACCAC | 39–406 | 368 | 17 |
| VP7-406R | CTTTAAAATANADGADCCWRTYGGCCA |
For G4 genotyping, common primer 3′comVP7-R was used as the reverse primer, while for G5, common primer 5′comVP7-F was used as the forward primer.
A chi-square test was used to assess the relationship between several factors (farm, year, season, age group, and diarrheal status) and RVA detection using procedure frequency in the SAS computer program (18).
Genotyping and genetic analysis of RVA strains.
Using the known human, porcine, and bovine RVA VP7 gene sequences available in GenBank, we designed genotype-specific primers (Table 2) for common G genotypes found in swine (G3, G4, G5, G9, and G11) to genotype the RVA-positive samples directly without sequencing. The specificity of these G-typing primers was evaluated using a panel of well-characterized laboratory reference strains consisting of human RVA strains Wa (G1), DS-1 (G2), M (G3), and 69 M (G8) and a characterized human G9P[6] strain obtained from the CDC and the porcine RVA strains Gottfried (G4) and OSU (G5). All genotype-specific one-step RT-PCRs were conducted individually with each primer pair, similarly to the above RVA detection RT-PCR protocol but with various annealing temperatures and extension times. Selected genotyped samples and the samples that could not be genotyped using the above primers were subjected to RT-PCR and partial sequencing using a universal primer set designed previously for the VP7 gene (17) to determine the G types for all strains and perform phylogenetic analysis. P types were determined using a universal primer pair (793F/1583R) for RT-PCR and partial sequencing of the RVA VP4 gene (nt 793 to 1604) (this study).
The nucleotide sequences obtained from the selected strains were compared with the same segments of similar sequences of known strains available in GenBank using BLAST software (http://www.ncbi.nlm.nih.gov/BLAST/). The DNA sequences were aligned using the ClustalW method. The dendrograms were constructed using the neighbor-joining method supported with a bootstrap test of 1,000 replicates for each gene in MEGA5 software (19). The tree was drawn to scale, with branch lengths in the same units as those of the evolutionary distances used to infer the phylogenetic tree. The evolutionary distances were computed using the maximum composite likelihood method (19) and are in the units of the number of base substitutions per site. The P genotypes were assigned to each sample after sequence analysis using either a BLAST (blastn) search (http://www.ncbi.nlm.nih.gov/) and/or RotaC2.0 automated genotyping tool for group A rotaviruses (20).
RESULTS
Incidence of RVA.
A total of 371 samples from five swine herds were tested, and 9.4% (35/371) were positive for RVAs (Table 3). In comparison with screening results for groups B (RVB) and C (RVC) rotaviruses and porcine sapoviruses (SaV), we found that 25.7% (9/35) of RVA-positive samples also contained either RVB or RVC or SaV. There were significant differences in RVA detection among the farms (P < 0.01) and the years (P < 0.05) studied. The prevalence of RVAs varied among farms (0 to 18.9%) and between different years within the same farm, such as seen in farm 2, ranging from 1.4% to 13.8% (Table 3). The RVAs were detected in four out of the five farms sampled. An increase in RVA detection was observed from 5.9% in 2004 to 8.5% in 2011 and then to 13.8% in 2012. There were no significant differences (P > 0.05) in RVA detection rates between the seasons during the years surveyed. Overall, the prevalence of RVAs was highest in spring (11.1%), followed by summer (8%) and then winter (6.9%). However, RVA detection rates in seasons differed within each year: in 2004, the highest prevalence occurred in spring (13%), while in 2011, summer (11.6%) had the highest prevalence, and in 2012, the detection rate was highest in winter (18.2%), as shown in Table 3.
Table 3.
Summary of RVA prevalence in the Ohio swine herds during the study period
| Parametera | No. of samples | Prevalence (%)b |
|---|---|---|
| Farm and/or year(s) | ||
| Farm 1 (2004) | 32 | 15.6 |
| Farm 2 | 253 | 8.7 |
| 2004 | 71 | 1.4 |
| 2011 | 59 | 6.8 |
| 2011 | 123 | 13.8 |
| Farm 3 (2004) | 15 | 6.7 |
| Farm 4 (2011) | 37 | 18.9 |
| Farm 5 (2011) | 34 | 0.0 |
| Total | 371 | 9.4 |
| Year and seasons | ||
| 2004 | 118 | 5.9 |
| Spring | 46 | 13.0 |
| Summer | 31 | 0.0 |
| Winter | 41 | 2.4 |
| 2011 | 130 | 8.5 |
| Spring | 41 | 2.4 |
| Summer | 69 | 11.6 |
| Winter | 20 | 10.0 |
| 2012 | 123 | 13.8 |
| Spring | 112 | 13.4 |
| Winter | 11 | 18.2 |
| Seasons | ||
| Spring | 199 | 11.1 |
| Summer | 100 | 8.0 |
| Winter | 72 | 6.9 |
| Diarrheal status | ||
| Diarrhea | 62 | 17.7 |
| Nondiarrhea | 61 | 9.8 |
| Age group | ||
| Nursing | 76 | 21.1 |
| Weaned | 47 | 2.1 |
Information on diarrheal status and age group was available only for 123 samples collected in 2012.
There were significant differences in prevalence rates among the farms (P < 0.01) and the years (P < 0.05) studied. There were no significant differences (P > 0.05) in RVA detection rates between the seasons during the years surveyed. Diarrheal status was also not a significant factor (P > 0.05), whereas detection in nursing piglets and weaned piglets differed significantly (P < 0.01).
The relationship between RV detection and diarrhea was evaluated in 123 samples collected in 2012 since information about the diarrheal status was not available for samples collected in 2004 and 2011. Although the diarrhea rate was higher in the RVA-positive pigs, diarrheal status had no significant influence (P > 0.05) on RVA incidence in the surveyed farms. RVA was detected in 17.7% (11/62) and 9.8% (6/61) of the diarrheic and nondiarrheic piglets, respectively (Table 3). Additionally, of the 123 samples collected in 2012, RVA was detected significantly (P < 0.01) more often in nursing piglets (21.1%, 16/76) than in weaned piglets (2.1%, 1/47), as shown in Table 3. Within the nursing piglets, our study showed that RVA detection was higher in piglets without diarrhea (27.8%) than in piglets with diarrhea (19%), whereas in weaned piglets, RVA was detected in only one sample from an asymptomatic piglet.
Assessment of the specificity of gene-specific primers for G genotyping.
The specificity of the G-typing primers designed in this study (for G3, G4, G5, G9, and G11) was evaluated. There were no cross-reactions among the genotypes tested; hence, these primers were used for the G typing of RVAs. However, it is important to note that these primers were used separately in a single reaction for the detection of specific genotypes.
Distribution of G and P genotypes.
Our results indicated that several genotypes of RVA are currently circulating in swine in Ohio. Of the 35 RVA-positive samples, 23 were further characterized by G and P typing, and 12 samples were of insufficient quantity/quality for further molecular characterization. The G types were determined for 95.7% (22/23) of the samples using our genotype-specific primers (Table 2), whereas one sample was G-genotype untypeable. The P types were determined for 91.3% (21/23) of the samples, whereas two samples were untypeable (Table 4). We used a combination of BLAST (blastn) search, phylogenetic tree analyses (MEGA5), and RotaC2.0 web-based tool (20) analysis of the partial sequences of VP7 and VP4 genes of the 23 samples with those of established RVA genotypes.
Table 4.
Combinations of G and P genotypes of 23 porcine RVAs identified in U.S. farms
| Genotype | No. of samples (yr of isolation) |
||||
|---|---|---|---|---|---|
| G4 | G9 | G11 | Unknown | Total | |
| P[7] | 0 | 2 (2012) | 1 (2004) | 0 | 3 |
| P[13] | 2 (2011) | 11 (2011), 3 (2012) | 1 (2004) | 1 | 18 |
| Unknown | 0 | 1 (2011), 1 (2012) | 0 | 0 | 2 |
| Total | 2 | 18 | 2 | 1 | 23 |
Using G-type-specific primers or an established nucleotide cutoff value of 80% for delineation of VP7 and VP4 genotypes (if universal primers followed by direct sequencing were used) (9), the 23 strains were classified into three G-genotypes (G4, G9, and G11) in combination with two P genotypes (P[7] and P[13]) that are either commonly or exclusively associated with porcine RVAs. Among the G types, G9 was predominant in 78% of the RVA-positive samples, while G4 and G11 each were detected in 9%. The P[13] genotype was dominant among the P types, constituting 78% of the RVA-positive samples, followed by P[7] (13%). Five G-P type combinations were identified, with G9P[13] being the dominant combination (61%), followed by G9P[7] and G4P[13] at 9% each, while G11P[7] and G11[13] were identified in one sample each (4%). Two samples were untypeable for P type, and one sample was untypeable for G type (Table 4). The G9 genotype was detected in all 3 years while G4 and G11 genotypes were detected only in samples from 2011 and 2004, respectively. The P[13] genotype was detected in all the years studied, while P[7] was detected only in 2004 and 2012.
Sequence analysis of the partial VP7 genes of G4, G9, and G11 porcine RVAs.
From the 22 strains that were G typed, seven randomly selected strains (two G4, three G9, and two G11, from farms 1, 2, and 4, from 2004 and 2011) were sequenced for further characterization of the partial VP7 gene (316 nt) and the assessment of the genetic identity within each G type. The two G4 strains (RV0120 and RV0132 [farm 2]) detected in 2011 were closely related to one another (95%), and they formed a separate branch in the phylogenetic tree (Fig. 1). The RV0120 and RV0132 strains displayed 79 to 85% and 82 to 87% nucleotide identity with G4 human RVAs and 77 to 82% and 77 to 85% identity with porcine RVAs, respectively, with the highest nucleotide identity (85% to 87%) to human RV D151 identified in Mexico. Interestingly, the two strains showed the lowest nucleotide identity (77%) to porcine G4 strain 1/07/Ire identified in Ireland. Three G9 strains (RV0036 [farm 4], RV0084, and RV0146 [farm 2]) detected in 2011 and 2012 were closely related to each other, with 90 to 97% nt identity. They were also closely related (89 to 91%) to both human and porcine G9 RVAs. However, they formed a separate branch in the phylogenetic tree (Fig. 1). Two strains (QW359 and QW362 [farm 1]) from historical samples (2004) that were characterized as G11 strains were identical to each other, with 100% nucleotide identity. They displayed 85 to 86% nucleotide identity with human G11 RVAs and 83% nucleotide identity with other G11 porcine RVAs. Both the QW359 and QW362 strains clustered in a monophyletic branch in the phylogenetic tree (Fig. 1).
Fig 1.
Phylogenetic dendrogram of the partial (nt 73 to 388) sequence of the RVA VP7 gene of field strains (boldface) compared with available VP7 gene sequences for human and porcine RVA G genotypes. Bootstrap values less than 50% are not shown.
Sequence analysis of the partial VP4 genes of P[7] and P[13] porcine RVAs.
The VP4 type was determined for 21 of the 23 RVA samples. A BLAST search for the 21 strains identified them as P[7] or P[13] types. A phylogenetic tree was constructed using the partial VP4 gene sequences (635 bp) of the 14 selected strains identified in this study and those of the existing 35 P genotypes in GenBank (Fig. 2). The quality of seven samples was insufficient for sequence analysis. The three P[7] strains shared high nucleotide identity (91 to 100%) with one another and with other P7 porcine strains. Two P[7] strains (QW362 and RV0161) clustered in a monophyletic branch together with the OSU (X13190) strain, which was identified in the United States, while the RV0263 strain clustered with the PRG9121 (JF796737) strain detected in South Korea (Fig. 2). Additionally, a BLAST (blastn) search and ClustalW analysis of the VP4 segment of our P[7] field strains demonstrated that they were closely related (92 to 99%) to bovine RVA strains (KJ25-1_HM988968 and KV0407_EU873009) isolated in South Korea and to porcine-like RVA strain CH-1 (HQ641296) detected in a panda in China.
Fig 2.
Phylogenetic dendrogram of the partial (nt 859 to 1493; 635 bp) nucleotide sequence of the RVA VP4 gene of field strains (boldface) compared with available VP4 gene sequences for RVA P genotypes. (Insets) Trees for the P[13] (a) and P[7] (b) strains. The porcine OSU strain (boldface) is the prototype for the P[7] genotype. Bo, bovine; Po, porcine.
The P[13] genotypes appeared to be genetically diverse in the surveyed farms. The phylogenetic analysis of the partial VP4 gene showed five genetic sublineages of the P[13] genotype: one sublineage formed by strains identified in Japan, a second sublineage by strains from Thailand and India, a third sublineage formed by a strain from South America, a fourth sublineage branch formed by an Australian strain, and the fifth new sublineage formed by some of our strains (Fig. 2). Analysis of P[13] strains identified in this study and VP4 gene sequences available in GenBank showed that they were from porcine RVAs sharing variable nucleotide identity rates among themselves of 81% to 99%. On the phylogenetic tree, they clustered together with other porcine P[13] strains, forming three distinct lineages: (i) India/Thailand lineage (HP113 and CMP213), (ii) Japanese lineage (FGP28), and (iii) a new lineage, U.S. strains (Fig. 2).
Upon analysis using the RotaC2.0 tool, a close relationship was also observed between our P[13] strains and a number of P[13] porcine strains, of which strain A46 (AY050274) shared the highest nucleotide identity (83.9 to 86.1%). On the other hand, another close relationship was found between P[7] strains, of which strain OSU (X13190) shared the highest nt identity (92 to 99.6%).
DISCUSSION
Group A rotaviruses are an important cause of acute gastroenteritis in children and young animals. Epidemiological information related to the prevalence and genotype specificities of porcine RVAs are beneficial for the development of accurate diagnostic tools and effective prophylactic measures against RVA infections in swine and to survey RVA strains closely related to human strains with potential for zoonotic transmission. In this study, evidence was collected for the occurrence of RVAs in nursing and weaned piglets in U.S. pig farms, and the genetic heterogeneity among the strains identified was examined. The RVAs were detected in 9.4% of the fecal samples from four of the five swine herds over the 3 years of the study, suggesting the continual circulation of RVA in swine farms. Surprisingly, no RVA strains were detected from farm 5. The prevalence of RVAs (9.4%) in this study was low compared to previous reports: 63.6% in the United States (21), 26% in Spain (22), 38.3% in South Korea (23), 67.3% in Japan (1), 17.2% (2009 to 2010) and 22.3% (2000 to 2001) in Thailand (24, 25), 17% in Canada (26), 35.3% in Brazil (27), and 25.7% in India (28). However, the RVA prevalence reported in this study was higher than that reported in preweaned piglets in Argentina (3.3%) (29), in suckling and weaned piglets in Germany (4%) (30), and in weaned asymptomatic piglets in Ireland (6.5%) (3). Our results are similar to the prevalence reported in Denmark, (10%) (31), between 2006 to 2008 in Thailand (10.7%) (10), and in Canada (9.2%) (32). It is noteworthy that the above reports affirm variable RVA prevalence rates in the same geographic regions in different years. Consistent with findings by others, we observed that 25.7% (9/35) of RVA-positive samples, all from symptomatic nursing piglets, were mixed infections with RVB, RVC, or SaV (14), suggesting that a number of enteric pathogens, either singly or in combination, may have synergistic effects on the clinical course of porcine RVA infections (21, 33–36). In contrast, RVA infections alone in nursing pigs receiving sow colostral and milk antibodies to RVA may cause only transient diarrhea that may have subsided at the time of sample collection in our study. Recently, Marthaler et al. (21) reported higher mixed infection rates (86.4%) of RVA, RVB, and RVC in piglets in the United States; however, they examined only diagnostic diarrheic pig samples.
There was a significant difference in the RVA detection rates among the five farms (P < 0.01) and the 3 years (P < 0.05) studied. An increase in RVA detection was observed from 5.9% in 2004 to 13.8% in 2012; however, the seasonal influences (P > 0.05) on the prevalence of RVA infections were unclear among the years sampled (Table 3). To date, there is a lack of information showing a clear seasonal distribution of porcine RVAs in the United States and also globally; hence, further epidemiological studies are required to fully understand the seasonal pattern of RVA to implement effective intervention or management strategies for the porcine RVA infections.
Porcine RVAs have been detected in diarrheic and nondiarrheic fecal samples from nursing and postweaning pigs either alone or in combination with other enteric pathogens (7, 37–39). In this study, we detected RVAs in 17.7% and 9.8% of the diarrheic and nondiarrheic fecal samples, respectively, in 2012. Asymptomatic infections with RVA in swine have been reported in previous studies (3, 29, 31). The prevalence of RVAs in asymptomatic piglets observed in this study (9.8%) was slightly higher than that reported in Spain (7%) (22) and Ireland (6.5%) (3). However, a higher prevalence of RVA in asymptomatic pigs was reported in Slovenia (20%) (31) and Italy (71.5%) (35). When we compared the data with diagnostic results for RVB and RVC, RVA was detected more frequently (9.8%) in asymptomatic pigs than RVC (6.6%) and RVB (0%) (unpublished data). This study also revealed that RVAs are more frequently detected in nursing piglets (21.1%, 16/76) than in weaned piglets (2.1%, 1/47), which is consistent with prior reports (2, 3). Among the nursing piglets, RVA was more often detected in asymptomatic (27.7%, 5/18) than in symptomatic (19%, 11/58) piglets although due to low sample numbers the differences were not statistically significant. Higher maternal immunity levels (40), the circulation of naturally attenuated RVA strains (41, 42), or RV shedding before the onset or after the resolution of diarrhea could account for the occurrence of RVA infections in asymptomatic animals.
The VP4- and VP7-encoding genes have different levels of nucleotide conservation since they are targets of host neutralization antibodies. Thus, evaluating the differences in terms of pathogenicity, virulence, and the nucleotide diversity in these genes is important in understanding RV infections and in the prediction of efficiency of vaccines (43). In this study, we investigated the genetic heterogeneity of RVAs circulating in swine farms in Ohio. The G9 genotypes were detected at higher levels (78%), which was consistent with a recent study in Thailand (64.3%) and Spain (33%), where it was found to be the most common genotype (10, 31). However, this was in contrast with a recent study in Denmark and Slovenia (31) and South Korea (23), where G9 was detected at low levels (4% and 9.2%, respectively) in swine. The P[13] genotype was the most common P type (78%) in the U.S. swine farms surveyed. The P[13] genotypes showed higher diversity among themselves, forming three different sublineages, with one sublineage clustering with the India/Thailand strains and another sublineage clustering with the Japanese strains while the third group clustered separately on their own branch. To our knowledge this is the first time RVA P[13] has been reported in U.S. swine at this high level. The RVA strains bearing P[13] VP4 specificity are host restricted, commonly detected among pigs, and have not been identified from other animal sources or humans (12, 44–46). Recent studies have reported the P[6] genotype as the most common P type in Slovenia (41%) and Spain (56%) in Europe (31). We detected three samples with P[7] genotypes, with two associated with G9 and one with G11 genotypes. Interestingly, regardless of the G genotypes, P[7] genotypes were phylogenetically closer to each other and to P[7] genotypes detected in swine, cattle, and pandas than to P[13] genotypes. The G11P[13] and G11P[7] strains contained almost identical G11 sequences, whereas the G11P[7] and G9P[7] strains had nearly identical P[7] sequences. This further supports the possibility of genetic reassortment between RVA strains within these farms.
In this study, we reported fewer distinct G-P genotype combinations (17), which was in contrast to recent studies in Europe (3, 13, 31), where more genotype combinations were reported. However, recent studies in Asia also reported fewer G-P genotype combinations in swine (10, 23), while in Canada a single genotype combination (G4P[6]) was most commonly detected (26). Although the detected G and P genotypes are common in pig populations worldwide (3, 6–8), the strains identified in this study displayed marked genetic variation, especially G9 and P[13] strains, suggesting genetic variability in RVA strains circulating in the U.S. pig population. The genetic diversity, such as reported in this study, in the same region (and sometimes the same farm) may have resulted from the introduction of new RVA strains.
Since RV infections cannot be diagnosed solely based on clinical signs, laboratory tests are necessary for reliable RV surveillance and can be useful in clinical settings to avoid inappropriate use of antibiotics which cannot prevent viral diarrheas. However, since it was not possible to characterize G and P genotypes for some RVA-positive samples, which may be attributed to either low viral load in the fecal sample or variability in the primer binding sites due to genetic diversity at the nucleotide level (3, 43, 47), it is necessary to constantly review and update genotyping primers as more sequencing data become available, as we had to do in this study.
To date, the only RVA vaccine (ProSystem Rota; Intervet Inc./Merck Animal Health) licensed for swine in the United States contains only G4 and G5 genotypes of RVA, according to the manufacturer, with no information about P genotype. However, none of the farms surveyed used the RVA vaccine; hence, it is unclear whether the diversity reported in this study may pose a challenge for future prophylactic strategies for prevention and control of RV-associated gastroenteritis in nursing and weaned piglets. Therefore, given the low frequency of RVA detected and the low number of farms surveyed within only Ohio, large-scale epidemiological investigations are needed to better evaluate both the ecology and genetic diversity of porcine RVAs circulating in U.S. swine farms. Acquiring information on the epidemiology of RVAs, non-group A RVs, and other viral enteric pathogens will be important to design effective prophylactic measures, to decrease the impact of enteric infections in swine, and to minimize the financial loss for swine producers.
ACKNOWLEDGMENTS
We gratefully acknowledge the cooperation and assistance of the Ohio swine farm veterinarians and managers in sample collection. We are grateful to Baoming Jiang (CDC) for providing us with the characterized human RVA G9P[6] strain.
This work was supported in part by the Ohio Agricultural Research and Development Center, Ohio State University SEED grant (2011-077), and a National Pork Board grant (NPB 12-094) for swine health. Salaries and research support were provided by state and federal funds provided to the Ohio Agricultural Research and Development Center, The Ohio State University.
APPENDIX
The GenBank accession numbers of all field strains and representative known RVA strains for each gene mentioned in this study are listed in Table A1.
Table A1.
GenBank accession numbers for each gene segment (VP7 and VP4) of RVA field strains and available known strains used in this studya
| Gene | Reference strain | Host | Accession no. |
|---|---|---|---|
| VP7 | B3458 | Human | EF990708 |
| CAU-1 | Human | HQ198807 | |
| D151 | Human | AJ488586 | |
| CIT-176RV | Human | AF254139 | |
| Cr117 | Human | AF450294 | |
| CUK1 | Human | EF121951 | |
| IAL-28 | Human | L79916 | |
| JP8485 | Human | HQ230034 | |
| Melb-G9-19 | Human | AY307089 | |
| MI09 | Human | HQ018934 | |
| MS037-03 | Human | AB436831 | |
| R479 | Human | DQ873680 | |
| VN-9 | Human | FJ598040 | |
| 1/07/Ire | Porcine | FJ617255 | |
| A46 | Porcine | L35054 | |
| CMP003 | Porcine | AY707787 | |
| CMP121 | Porcine | DQ683522 | |
| CMP178 | Porcine | DQ515961 | |
| CMP213 | Porcine | DQ786576 | |
| CMP77 | Porcine | DQ683521 | |
| FGP28 | Porcine | AB573873 | |
| FGP35 | Porcine | AB573875 | |
| FGP36 | Porcine | AB573877 | |
| GUB72 | Porcine | AB573649 | |
| Hokkaido-14 | Porcine | AB176677 | |
| HP113 | Porcine | DQ003292 | |
| HP140 | Porcine | DQ003293 | |
| Ire/60/2007 | Porcine | FJ617260 | |
| JP13-3 | Porcine | AB176679 | |
| JP32-4 | Porcine | AB176682 | |
| JP35-7 | Porcine | AB176683 | |
| Arg/P3 | Porcine | AY115862 | |
| Arg/P14 | Porcine | AY115861 | |
| Arg4671 | Human | FJ712692 | |
| RVA/Pig-wt/USA/QW359/2004 | Porcine | KC244310 | |
| RVA/Pig-wt/USA/QW362/2004 | Porcine | KC244311 | |
| RVA/pig-wt/USA/RV0036/2011 | Porcine | KC244285 | |
| RVA/pig-wt/USA/RV0084/2011 | Porcine | KC244286 | |
| RVA/pig-wt/USA/RV0120/2011 | Porcine | KC244312 | |
| RVA/pig-wt/USA/RV0132/2011 | Porcine | KC244313 | |
| RVA/pig-wt/USA/RV0146/2012 | Porcine | KC244287 | |
| VP4 | VMRI | Bovine | U53923 |
| GRV | Caprine | AB055967 | |
| CH3 | Equine | D25228 | |
| FI23 | Equine | D16342 | |
| L338 | Equine | JF712558 | |
| Cat97 | Feline | EU708948 | |
| FRV64 | Feline | D14723 | |
| A_B10 | Human | HM627556 | |
| RMC321 | Human | AF523677 | |
| 116E | Human | L07934 | |
| 69 M | Human | M60600 | |
| AU-1 | Human | D10970 | |
| B4106 | Human | AY740738 | |
| Dhaka6 | Human | GU199520 | |
| DS-1 | Human | HQ650119 | |
| Ecu534 | Human | EU805773 | |
| G9_F45 | Human | U30716 | |
| Ch-04V0027 | Avian | EU486961 | |
| AvRV-2 | Avian | JQ085405 | |
| Ch-06V0661 | Avian | EU486962 | |
| PO-13 | Avian | AB009632 | |
| Tu-03V0002E10 | Avian | EU486958 | |
| 993-83 | Bovine | D16352 | |
| A5 | Bovine | D13395 | |
| AzuK-1 | Bovine | AB454420 | |
| C486 | Bovine | Y00127 | |
| CACC | Bovine | AB486010 | |
| Dai-10 | Bovine | AB513836 | |
| Hg18 | Bovine | AF237665 | |
| KJ25-1 | Bovine | HM988968 | |
| KV0407 | Bovine | EU873009 | |
| M63267 | Bovine | M63267 | |
| M33516 | Porcine | M33516 | |
| MDR-13 | Porcine | L07886 | |
| I321 | Human | L07657 | |
| K8 | Human | D90260 | |
| KTM368 | Human | GU199495 | |
| Mc323 | Human | D38052 | |
| Mc35 | Human | D14032 | |
| TB-Chen | Human | AY787644 | |
| US1205 | Human | AF079356 | |
| Wa | Human | JX406750 | |
| Murine_EB | Murine | U08419 | |
| Murine_EC | Murine | U08421 | |
| Murine_EHP | Murine | U08424 | |
| Lamb_L11599 | Ovine | L11599 | |
| Lamb_Lamb-NT | Ovine | FJ031027 | |
| 134/04-15 | Porcine | DQ061053 | |
| 344–04-1 | Porcine | DQ242615 | |
| 61/07/Ire | Porcine | FJ492835 | |
| A46 | Porcine | AY050274 | |
| BMI-1 | Porcine | L07887 | |
| CMP034 | Porcine | DQ534016 | |
| CMP178 | Porcine | QD536362 | |
| CMP213 | Porcine | DQ786578 | |
| FGP28 | Porcine | AB573874 | |
| FGP35 | Porcine | AB573876 | |
| FGP36 | Porcine | AB573878 | |
| GUB72 | Porcine | AB573650 | |
| HP113 | Porcine | DQ003290 | |
| HP140 | Porcine | QD003291 | |
| JL94 | Porcine | AY523636 | |
| PRG921 | Porcine | JF796715 | |
| OSU | Porcine | X13190 | |
| PRG9121 | Porcine | JF796737 | |
| Rhesus_TUCH | Simian | FJ816611 | |
| SA11 | Simian | D16346 | |
| RVA/pig-wt/USA/QW359/2004 | Porcine | KC244301 | |
| RVA/pig-wt/USA/QW362/2004 | Porcine | KC244290 | |
| RVA/pig-wt/USA/RV0036/2011 | Porcine | KC244291 | |
| RVA/pig-wt/USA/RV0042/2011 | Porcine | KC244292 | |
| RVA/pig-wt/USA/RV0044/2011 | Porcine | KC244302 | |
| RVA/pig-wt/USA/RV0047/2011 | Porcine | KC244293 | |
| RVA/pig-wt/USA/RV0084/2011 | Porcine | KC244294 | |
| RVA/pig-wt/USA/RV0120/2011 | Porcine | KC244295 | |
| RVA/pig-wt/USA/RV0132/2011 | Porcine | KC244296 | |
| RVA/pig-wt/USA/RV0145/2012 | Porcine | KC244297 | |
| RVA/pig-wt/USA/RV0146/2012 | Porcine | KC244298 | |
| RVA/pig-wt/USA/RV0147/2011 | Porcine | KC244303 | |
| RVA/pig-wt/USA/RV0148/2011 | Porcine | KC244304 | |
| RVA/pig-wt/USA/RV0154/2011 | Porcine | KC244305 | |
| RVA/pig-wt/USA/RV0160/2012 | Porcine | KC244299 | |
| RVA/pig-wt/USA/RV0161/2012 | Porcine | KC244288 | |
| RVA/pig-wt/USA/RV0254/2011 | Porcine | KC244306 | |
| RVA/pig-wt/USA/RV0258/2011 | Porcine | KC244307 | |
| RVA/pig-wt/USA/RV0262/2011 | Porcine | KC244300 | |
| RVA/pig-wt/USA/RV0263/2012 | Porcine | KC244289 | |
| RVA/pig-wt/USA/RV0264/2012 | Porcine | KC244308 |
Field strains are shown in boldface.
Footnotes
Published ahead of print 30 January 2013
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