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Effect of strain-specific maternally-derived antibodies on influenza A virus infection dynamics in nursery pigs

  • Fabian Orlando Chamba Pardo,

    Roles Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Supervision, Validation, Visualization, Writing – original draft, Writing – review & editing

    Affiliation Veterinary Population Medicine Department, University of Minnesota, St. Paul, MN, United States of America

  • Spencer Wayne,

    Roles Conceptualization, Methodology, Supervision, Writing – review & editing

    Affiliation Health Services, Pipestone Veterinary Services, Pipestone, MN, United States of America

  • Marie Rene Culhane,

    Roles Conceptualization, Funding acquisition, Investigation, Methodology, Supervision, Writing – review & editing

    Affiliation Veterinary Population Medicine Department, University of Minnesota, St. Paul, MN, United States of America

  • Andres Perez,

    Roles Conceptualization, Formal analysis, Funding acquisition, Methodology, Writing – review & editing

    Affiliation Veterinary Population Medicine Department, University of Minnesota, St. Paul, MN, United States of America

  • Matthew Allerson,

    Roles Conceptualization, Funding acquisition, Investigation, Methodology, Writing – review & editing

    Affiliation Health and Research Division, Holden Farms Inc., Northfield, MN, United States of America

  • Montserrat Torremorell

    Roles Conceptualization, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Writing – review & editing

    Affiliation Veterinary Population Medicine Department, University of Minnesota, St. Paul, MN, United States of America


Reducing the number of influenza A virus (IAV) infected pigs at weaning is critical to minimize IAV spread to other farms. Sow vaccination is a common measure to reduce influenza levels at weaning. However, the impact of maternally-derived antibodies on IAV infection dynamics in growing pigs is poorly understood. We evaluated the effect of maternally-derived antibodies at weaning on IAV prevalence at weaning, time of influenza infection, number of weeks that pigs tested IAV positive, and estimated quantity of IAV in nursery pigs. We evaluated 301 pigs within 10 cohorts for their influenza serological (seroprevalence estimated by hemagglutination inhibition (HI) test) and virological (prevalence) status. Nasal swabs were collected weekly and pigs were bled 3 times throughout the nursery period. There was significant variability in influenza seroprevalence, HI titers and influenza prevalence after weaning. Increase in influenza seroprevalence at weaning was associated with low influenza prevalence at weaning and delayed time to IAV infection throughout the nursery. Piglets with IAV HI titers of 40 or higher at weaning were also less likely to test IAV positive at weaning, took longer to become infected, tested IAV RT-PCR positive for fewer weeks, and had higher IAV RT-PCR cycle threshold values compared to piglets with HI titers less than 40. Our findings suggest that sow vaccination or infection status that results in high levels of IAV strain-specific maternally-derived antibodies may help to reduce IAV circulation in both suckling and nursery pigs.


Influenza A virus (IAV) is a primary cause of acute respiratory disease in pigs and it is also part of the porcine respiratory disease complex (PRDC), which includes other pathogens such as porcine reproductive and respiratory syndrome virus (PRRSV), porcine circovirus 2 (PCV2), and Mycoplasma hyopneumoniae [1]. IAV infection affects the performance of pigs by increasing feed conversion [2] and mortality [3], decreasing body weight gain [4] and reducing semen quality of boars [5]. IAV in pigs also represents a threat to public health since it is a zoonosis with pandemic potential.

Influenza is widespread in US pigs and breed-to-wean (BTW) pig farms play a central role in the spread of IAV across geographical regions [69]. Suckling pigs maintain, diversify and transmit IAV when moved to other farms [1013]. Commonly, pigs are weaned at 21 days of age and are moved to distant locations to grow to market. The significant IAV genetic diversity in pigs is a result not only from pig movements but also due to the rapid mutation rate of the virus (genetic drift) [1417] and the introduction of IAV from humans (reverse zoonosis) [1823] and birds [2426], which facilitates the emergence of novel reassorted strains (genetic shift). H1N1, H1N2 and H3N2 are the most common subtypes found in pigs and the introduction of gene segments from other species into a pool of endemic viruses has resulted in a complex landscape of IAV in pigs. Indeed, currently there are 16 genetically and antigenically distinct H1 clades (alpha, beta, gamma, gamma 2, delta 1a, delta 1b, delta 2 and pandemic 2009) [15, 17, 27] or H3 clusters (IV A-F, human-like 2011 and human-like 2016) [9, 14, 28, 29] of IAV co-circulating in US pigs. This broad genetic diversity and the common co-circulation of several clades within a farm or production system represents a critical hurdle for vaccines to induce cross-protective immunity effective against genetically diverse strains [30, 31].

Sow vaccination is the main measure to control influenza in BTW farms [3235]. Sow vaccination helps protecting the herd from severe clinical disease and enhances the transfer of maternally-derived antibodies (MDA) from sows to piglets through colostrum. MDA protect piglets from clinical disease shortly after weaning [3639] and strain-specific MDA can also decrease IAV transmission in weaned pigs [40]. However, although the use of IAV vaccines in BTW farms is common, IAV still circulates in pigs likely due to the limitations for generating sufficient levels of immunity against the multiple strains co-circulating [11, 41].

In addition, pigs have complex IAV infection dynamics, such as recurrent infections, multiple waves of infection, co-circulation of genetically distinct viruses and frequent reassortment events observed in growing-finishing pigs [4249]. However, quantitative knowledge on the factors that are associated with the complex IAV infection dynamics in nursery pigs is limited. Also, there is limited information about how IAV MDA may impact infection dynamics in nursery pigs. Filling this knowledge-gap is critical to guide successful intervention strategies for BTW farms that aim to reduce IAV circulation in pigs before and after weaning. In our study, we evaluated the effect of IAV seroprevalence and individual MDA levels at weaning on IAV prevalence at weaning, time to IAV infection after weaning, number of weeks that pigs tested IAV RT-PCR positive, and IAV RT-PCR cycle threshold values in pigs after weaning. Results from our study contributed to determine whether sow interventions in the breeding herd have a direct benefit in piglets at weaning and throughout the growing period in the nursery.

Materials and methods

Ethics statement

This study was carried out after the protocol was approved by the Institutional Animal Care and Use Committee (Protocol Number: 1510-33054A) and the Institutional Biosafety Committee (Protocol Number: 1508-32918H) of the University of Minnesota. The participating producers provided written consent to conduct the study in their farms. All samples were collected by trained veterinarians and researchers. Pigs were raised indoors in conventional mechanically ventilated nursery barns with climate control. Pigs had ad libitum access to fresh food and water and farmers monitored the health of the pigs twice daily to detect and treat any sick pig. Pigs were cared according to recommendations by the herd veterinarian which included antipyretic and antibiotic treatments on as needed basis.

Study population and design

Piglets (n = 301) were identified at weaning and tested during the growing period in the nursery which included piglets from 3 to 10 weeks of age. Five batches of weaned piglets were placed into 2 separate nursery farms. Pigs in each batch were divided into 2 rooms for a total of 10 cohorts. Approximately 30 pigs within each cohort were selected and pigs were placed in a randomly selected pen within each room. Each nursery farm was a single-barn site with two rooms with all-in/all-out flow by site (all pigs entered and exited at once to facilitate cleaning and disinfection of the entire facility). Piglets were vaccinated at weaning against Mycoplasma hyopneumoniae, PCV2 and PRRSV (Ingelvac 3FLEX® Vaccine, Boehringer Ingelheim Vetmedica, Inc., St. Joseph, MO).

Piglets originated from a single air-filtered 3,200 sow farm located in Minnesota with a history of testing IAV positive by RT-PCR in piglets prior to weaning for at least the last 2 months before the study began. The sow farm was located within 10 miles from the nursery farms and known to be negative for wild-type PRRSV strains but positive for M. hyopneumoniae and PCV2. Additionally, incoming replacement females (gilts) were vaccinated twice with an inactivated IAV commercial vaccine (FluSure XP®; Zoetis Inc, Parsippany, NJ).

The study was conducted from November 2015 to April 2016. During that time, sows were vaccinated against PRRSV using a modified-live vaccine (Ingelvac PRRS® MLV vaccine, Boehringer Ingelheim Vetmedica, Inc., St. Joseph, MO) as part of their biannual vaccination against PRRSV. During 8 weeks within the study period (December 2015 to January 2016), sows were also vaccinated with 2 commercial IAV inactivated vaccines (Maxivac Excell 5.0®, Merck Animal Health, Madison, NJ; and FluSure XP®, Zoetis Inc, Parsippany, NJ) with cohorts 3a, 3b, 4a and 4b originating from vaccinated sows. There was no sow influenza vaccination before and after the described 8-week period during the study timeframe. Cohort details are explained in Table 1.

Sampling, testing and influenza characterization

Piglets in each cohort were ear-tagged, nasal swabbed and blood samples collected at weaning. Nasal swabs were collected weekly post-weaning until the end of the growing period in the nursery and blood samples collected at 3- and 6-weeks post-weaning. Nasal swabs were processed by suspending them in 2mL of viral transport media (Minimum Essential Media plus bovine serum albumin, antibiotics, antifungals and trypsin TCPK) and stored at -80°C until testing. Swabs were tested by a real-time RT-PCR targeting the IAV matrix gene [50].

Briefly, sample viral RNA was extracted using a MagMaxTM—96 viral RNA extraction kit (Applied Biosystems, Foster City, CA) following manufacturer’s instructions using an automatized robotic extraction equipment: MagMaxTM–Express 96 Deep well magnetic particle processor (Applied Biosystems, Foster City, CA). RT-PCR test reagents were from AgPath-ID one-step RT-PCR kit (Life Technologies, Grand Island, NY) and reactions were run in a 7500 Fast RT-PCR system (Life Technologies, Grand Island, NY). Thermal cycles for IAV detection were 10 min at 45°C, 10 min at 95°C, 45 cycles of 1 sec at 94°C and 30 sec at 60°C within a 25μL of total volume [50]. A sample was considered IAV rRT-PCR positive if the cycle threshold (ct) value was 37.5 or lower.

Additionally, two nasal swab samples with the lowest ct values in each cohort from weeks when IAV prevalence was the highest were selected for virus isolation in Madin-Darby Canine Kidney (MDCK) cells [51] and whole-genome sequenced (WGS) using the MiSeq Ilumina platform [52]. When IAV isolation was not possible, WGS was performed directly from selected nasal swab samples (cohorts 4a, 4b, 5a and 5b). Raw sequence reads were cleaned, trimmed and assembled against both H1 (FJ789832) and H3 (KC992248) reference strains obtained from the Influenza Virus Resource at the National Center for Biotechnology Information (NCBI) [53]. Hemagglutinin (HA) gene assembly was done using the Map to a Reference function in Geneious 8.1 software [54].

HA gene consensus sequences were annotated for completeness, functionality and subtype classification using the influenza virus sequence annotation tool (FLAN) [55]. Complete HA gene sequences were further characterized using BLAST tools and the global swine H1 clade was inferred using the swine H1 clade classification tool from the Influenza Research Database (IRD) [56].

HA gene sequences were aligned and translated into amino acid sequences to further analyze the amino acid similarities between viruses from the BTW farm, the studied cohorts and the IAV commercial vaccines used in sows. Alignment, amino acid translation and sequence comparisons were done using the ClustalW algorithm, Neighbor Joining method and a Jukes-Cantor substitution model in Geneious software [54]. H1 antigenic sites (Sa, Sb, Ca1, Ca2 and Cb) [5761] were compared among the obtained sequences using the above described analysis and the Identify Sequence Features in Segments tool of the IRD [56].

Finally, blood samples were left at room temperature for serum separation and then refrigerated overnight. Samples were centrifuged for 10 min at 1500 rpm and 4°C. Sera then were stored at -20°C until testing. Sera were tested by hemagglutination inhibition (HI) assay using a representative and dominant IAV isolate (H1 delta 2) among all cohorts and BTW farm. Briefly, sera were pretreated with receptor destroying enzyme (RDE) and incubated at 37°C for 14 hours overnight. Incubated sera were hemadsorpted by adding a 20% turkey red blood cell solution, centrifuged, and stored at -20°C until titration. Two-fold serial dilutions from 1:10 to 1:640 were tested for each serum sample using an antigen solution containing 16 hemagglutination units (HAU) in 50uL and a 0.5% turkey red blood cell solution. Positive and negative controls were used in each plate to confirm the results obtained in plate. Plates were read manually by tilting them and visualizing inhibition of agglutination (presence of a mat of red blood cells in the bottom of the well) [62]. Additionally, 15 pigs in each cohort were selected according to their HI titers (half of the piglets within each HI titer) at weaning and serum samples tested by IAV NP ELISA (IDEXX Influenza A Ab Test, IDEXX Laboratories, Inc., Westbrook, Maine). A sample was considered positive if the S/N value was less than 0.6. ELISA testing was done according to the manufacturer’s protocol.

Data analysis

Individual HI titer data was tabulated and classified as HI-positive or -negative using a reciprocal HI titer of 40 as the cutoff point (≥40 were considered positive). RT-PCR data were tabulated and classified as positive or negative based on a cycle threshold (ct) cutoff point of 37.5. Then, the percentage of IAV RT-PCR and HI serum antibody positive pigs over time in the nursery was calculated and values reported as prevalence and seroprevalence, respectively. Prevalence and seroprevalence data were summarized by week for each cohort.

Time to IAV infection was calculated by counting the number of weeks from weaning until a pig tested IAV RT-PCR positive. The number of weeks that a given pig tested IAV RT-PCR positive during the growing period in the nursery was calculated by counting the total number of weeks that a pig tested IAV RT-PCR positive for the entire nursery period. Because the real-time RT-PCR used in this study is semi-quantitative, the estimated quantity of IAV virus was approximated from the RT-PCR ct values, with the lowest ct values having the highest estimated virus quantity and ct values of >37.5 having estimated virus quantity approaching zero. For a given pig, the lowest ct value during the entire study was chosen for the median calculations by cohort. Median values of time to IAV infection, number of weeks that a pig tested IAV RT-PCR positive and lowest ct values were calculated for each cohort and for each reciprocal HI titer group using the dplyr package in R 3.4.2 statistical software [63].

To test the association between IAV seroprevalence of the cohort and HI titer at weaning with prevalence at weaning, time to IAV infection, number of weeks that pigs tested IAV RT-PCR positive and lowest ct values during the nursery period, a Spearman non-parametric correlation test was used in STATA 12® statistical software (StataCorp, College Station, TX) [64]. Different regression models were used to test the association of HI titers at weaning with IAV RT-PCR positivity at weaning for each pig (logistic regressions), time to IAV infection and number of weeks that each pig tested IAV RT-PCR positive (Poison regressions), and lowest ct value of each pig during the growing period in the nursery (linear regressions). Regressions were modeled under generalized linear models (GLM) in the glimmix procedure of SAS 9.4® statistical software (SAS Foundation, Cary, NC) [65]. Models were adjusted by nursery farm and season effects as fixed covariates. Best models were chosen based on goodness of fit criteria using the lowest Bayesian Information Criterion (BIC) and visual inspection of the residuals to check model assumptions accordingly.


A total of 301 piglets from 10 cohorts were tested from weaning to the end of the growing period in the nursery. Overall, there were 885 serum samples of which 28% had a reciprocal HI titer of 40 or higher. Moreover, 348 of 2176 (16%) nasal swabs were IAV RT-PCR positive. In total, 185 of 301 pigs (62%) tested IAV RT-PCR positive at least once during the study (S1 Table).

We obtained 20 IAV HA gene sequences including 19 from the cohorts and 1 from the BTW farm prior to the start of the study. We characterized the HA gene sequences as H1 delta 2 (1B.2.1) viruses having 99.4% nucleotide and 99.3% amino acid similarity among themselves. The H1 delta 2 HA gene sequences obtained during the study shared 94.3% and 94.8% amino acid similarity to the H1 delta 2 commercial vaccine strains used in the BTW farm and there were no amino acid changes in the H1 antigenic sites (Sa, Sb, Ca1, Ca2 and Cb).

There was appreciable variability in prevalence and seroprevalence between cohorts over time (Fig 1). IAV prevalence at weaning changed from almost 100% in cohorts with low seroprevalence to about 0% in cohorts with high seroprevalence at weaning. Inversely, seroprevalence measured by HI titers against the H1N2 delta 2 IAV circulating farm strain at weaning changed over time from 0% in cohorts of piglets originating from non-vaccinated sows up to 80 and 90% in cohorts originating from vaccinated sows. Additionally, IAV prevalence during the growing period in the nursery at the cohort level varied. There were episodes of high prevalence of IAV infection at weaning (97–100%) with none or one recurrent infection after weaning to episodes of lower prevalence (~0%) at weaning with limited infection during the nursery, as shown in Fig 1.

Fig 1. Influenza A virus prevalence and seroprevalence by cohort during the nursery period.

Black bars are the seroprevalence (hemagglutination inhibition titer of 40 or higher), blue line is the prevalence (percentage of RT-PCR positives) and red dots is the percentage of NP ELISA positive pigs (S/N<0.6).

IAV NP ELISA results summarized as percentage of ELISA positive pigs over time in each cohort are also shown in Fig 1 (red dots). Almost all pigs at weaning tested IAV NP ELISA positive and percentage of positive pigs declined over time or increased slightly after the second peak of infection, similar to the HI titers dynamics observed in most of the cohorts. However, cohorts 1a, 1b, 2a and 2b had a high percentage of NP ELISA positive pigs at weaning and pigs had low HI titers and high levels of IAV infection. In these cohorts, pigs did not appear to have HI strain-specific antibodies that could prevent IAV infection.

Per cohort, high seroprevalence at weaning was significantly associated with lower IAV prevalence at weaning (correlation value of -0.71) and it took a longer time, more than 6 weeks, to become IAV infected after weaning. The number of weeks that pigs tested IAV positive, and lowest ct values during the growing period in the nursery were not significantly associated with seroprevalence at weaning although, there was a numerical trend that reflected limited circulation of IAV after weaning. Table 2 illustrates the association between IAV seroprevalence at weaning and our calculated IAV infection data after weaning for each cohort.

Table 2. Cohort-level correlation of influenza A virus (IAV) seroprevalence at weaning with infection parameters in the nursery.

HI titer group at weaning was significantly associated with infection dynamics after weaning as detailed in Table 3. According to their HI titer group at weaning, reciprocal HI titers of 40 or higher were significantly associated with lower IAV levels at weaning, longer time to IAV infection, fewer weeks that pigs tested IAV RT-PCR positive, and less estimated quantity of virus in groups of pigs during the growing period in the nursery.

Table 3. Correlation of influenza A virus (IAV) hemagglutination inhibition (HI) titer group at weaning with infection parameters in nursery pigs.

Individual HI titer at weaning was significantly associated with the IAV RT-PCR positive status during the growing period in the nursery as shown in Table 4. Pigs with a reciprocal HI titer of 40 or higher at weaning were less likely to test IAV RT-PCR positive at weaning, took longer to become IAV infected, tested IAV RT-PCR positive for fewer weeks and had higher ct values. We estimated a 5% probability of infection for pigs with reciprocal HI titers ≥ 40. Overall, high MDA levels at weaning decreased the likelihood of IAV infection and circulation at weaning and during the growing period in the nursery.

Table 4. Pig-level association of influenza A virus (IAV) hemagglutination inhibition titer at weaning with individual infection parameters during the nursery period.


Control of influenza in BTW farms is needed to minimize the spread of IAV across pig production systems and regions given that piglets infected at weaning covertly transmit IAV to other farms. Sow vaccination is the most common IAV control measure although the impact of MDA on infection after weaning is poorly understood. Our study showed that high levels of strain-specific MDA at weaning can decrease prevalence of IAV at weaning and more importantly, decrease the overall IAV circulation during the growing period in the nursery. Reducing IAV circulation at weaning should help decrease IAV transmission between farms.

Our study supports previous experimental work that demonstrated high MDA measured by strain-specific HI titers reduced significantly the transmission of IAV in weaned pigs [40]. It also supports case reports where IAV elimination was attempted using IAV strain-specific sow vaccination strategies that were successful in decreasing the detection of IAV in piglets at weaning for certain periods of time [34, 35]. Notably, our results help clarify the understanding of the effect of high levels of strain-specific MDA over IAV infection in the nursery phase under field conditions.

Our study was also similar to other studies that reported variable levels of IAV circulation and seroconversion in growing pigs and had one or more peaks of infection after weaning [37, 4247]. In contrast, our results differed from studies where heterologous antibodies had been measured. In those studies, heterologous MDA had no effect on IAV transmission in weaned pigs, prolonged the infectious period and blocked the active immune response to infection [36, 39, 40, 6669]. The immunological mechanism behind these latter observations is not well defined and B cell epitope masking has been proposed as one of the possible mechanisms. Maternal antibodies could cover some viral epitopes so B cells would not recognize those viral antigens and not mount the expected humoral response after infection or vaccination [70]. Nonetheless, our results indicate that high levels of homologous MDA decreased IAV levels at weaning and likely subsequent transmission during the growing period in the nursery.

Sow vaccination and MDA can protect sows and piglets from IAV clinical disease [36, 37, 39, 68, 69, 71]. Although sow vaccination is a common strategy to control IAV in BTW farms [32], IAV is still widespread in pigs. One of the major challenges to sow vaccination is the increasing IAV genetic diversity with about 16 different HA genetic clades currently circulating in US pig populations [1417, 1921, 2527, 31, 7280]. In this study, we found that homologous MDA against the circulating delta 2 H1 virus decreased delta 2 H1 IAV circulation in pigs after weaning. Since we did not sequence all viruses isolated from every pig nor did we sequence directly from every positive nasal swab, there is a possibility that we missed other viruses that might have been co-circulating in the study cohorts. Indeed, in one of the cohorts (Cohort 5a) we found partial sequences of 2 other viruses (H1 gamma (1A.3.3.3) and H3 subclade IV B) that had been historically identified in the sow farm. Why these viruses circulated at a low prevalence and did not spread throughout the entire population is uncertain. Immunity, viral spread and dominance of co-circulating viruses is puzzling and needs to be further studied in pig populations [8185].

Our sequencing approach demonstrated the circulation of one predominant strain (H1 delta 2) in the 10 cohorts and in the BTW farm. We found no differences in the H1 antigenic sites between the delta 2 H1viruses detected in our study and the vaccine strains. Our results indicated that the HI antibodies at weaning affected IAV infection dynamics in the nursery. We considered that pigs originating from vaccinated sows had higher levels of antibodies able to cross-react with the farm specific delta 2 H1 strain and that these were, at least in part, responsible for the decrease in IAV infection. Although almost all pigs tested ELISA IAV NP positive at weaning, NP antibodies did not appear to be protective against IAV infection in the cohorts with high infection at weaning. This may be an additional indication that HI strain-specific antibodies were not present to protect pigs from IAV infection against the circulating H1 farm strain prior or at weaning, especially in the first 4 cohorts originating from non-vaccinated sows. Finally, we cannot fully discard the possibility that sow vaccination may have boosted IAV natural infection before vaccination.

Understanding IAV immunity is critical when trying to optimize the vaccination strategies currently used in swine production systems. There are still gaps in our understanding of the quality and quantity of antibodies being generated after IAV vaccination and/or natural infection. This is particularly important in pigs where we have several strains cocirculating in the same population. The complex evolution and landscape of IAV in pigs highlights the need for better vaccines and comprehensive vaccination strategies [30, 31, 80].


In conclusion, we found that high levels of strain-specific hemagglutination inhibition antibodies at weaning decreased IAV infection at weaning, delayed time to become influenza-positive, decreased the number of weeks that pigs tested influenza-positive during the nursery stage, and decreased the overall estimated virus quantity during the infection period. Our results suggest that IAV infection and circulation in nursery pigs may be decreased by using adequate influenza sow vaccination protocols.

Supporting information


We would like to thank Dr. Joel Nerem from Pipestone Veterinary Services for his help in this study. We also thank Dr. Aaron Rendahl for his help on the statistical analysis. Thanks to Jay Nirmala and My Yang for their help in the laboratory work including the sequencing work. Finally, thanks to all the producers and farm personnel that helped us with this study activities.


  1. 1. Van Reeth K, Brown IH, Olsen CW. Influenza virus. In: Zimmerman J, Karriker L, Ramirez A, Schwartz K, Stevenson G, editors. Diseases of swine. 10. Ames, IA: Wiley-Blackwell; 2012. p. 557–71.
  2. 2. Er C, Lium B, Tavornpanich S, Hofmo PO, Forberg H, Hauge AG, et al. Adverse effects of Influenza A(H1N1)pdm09 virus infection on growth performance of Norwegian pigs—a longitudinal study at a boar testing station. BMC Vet Res. 2014;10:284. pmid:25472551; PubMed Central PMCID: PMCPMC4300606.
  3. 3. Alvarez J, Sarradell J, Kerkaert B, Bandyopadhyay D, Torremorell M, Morrison R, et al. Association of the presence of influenza A virus and porcine reproductive and respiratory syndrome virus in sow farms with post-weaning mortality. Preventive veterinary medicine. 2015;121(3–4):240–5. pmid:26210012.
  4. 4. Er C, Skjerve E, Brun E, Hofmo PO, Framstad T, Lium B. Production impact of influenza A(H1N1)pdm09 virus infection on fattening pigs in Norway. J Anim Sci. 2016;94(2):751–9. pmid:27065145.
  5. 5. Lugar DW, Ragland D, Stewart KR. Influenza outbreak causes reduction in semen quality of boars. Journal of Swine Health and Production. 2017;25(6):303–7. WOS:000413890100006.
  6. 6. Corzo CA, Culhane M, Juleen K, Stigger-Rosser E, Ducatez MF, Webby RJ, et al. Active surveillance for influenza A virus among swine, midwestern United States, 2009–2011. Emerging infectious diseases. 2013;19(6):954–60. pmid:23735740; PubMed Central PMCID: PMCPMC3713829.
  7. 7. Allerson MW. Transmission and control of influenza virus in pig populations [Thesis or Dissertation]. St. Paul, MN: University of Minnesota; 2013.
  8. 8. Kyriakis CS, Zhang M, Wolf S, Jones LP, Shim BS, Chocallo AH, et al. Molecular epidemiology of swine influenza A viruses in the Southeastern United States, highlights regional differences in circulating strains. Veterinary microbiology. 2017;211:174–9. pmid:29102115; PubMed Central PMCID: PMCPMC5709717.
  9. 9. Walia RR, Anderson TK, Vincent AL. Regional patterns of genetic diversity in swine influenza A viruses in the United States from 2010 to 2016. Influenza and other respiratory viruses. 2018. pmid:29624873.
  10. 10. Allerson MW, Davies PR, Gramer MR, Torremorell M. Infection dynamics of pandemic 2009 H1N1 influenza virus in a two-site swine herd. Transboundary and emerging diseases. 2014;61(6):490–9. pmid:23294593.
  11. 11. Diaz A, Marthaler D, Culhane M, Sreevatsan S, Alkhamis M, Torremorell M. Complete Genome Sequencing of Influenza A Viruses within Swine Farrow-to-Wean Farms Reveals the Emergence, Persistence, and Subsidence of Diverse Viral Genotypes. Journal of virology. 2017;91(18):JVI. 00745–17. pmid:28659482; PubMed Central PMCID: PMCPMC5571239.
  12. 12. Diaz A, Perez A, Sreevatsan S, Davies P, Culhane M, Torremorell M. Association between Influenza A Virus Infection and Pigs Subpopulations in Endemically Infected Breeding Herds. PloS one. 2015;10(6):e0129213. pmid:26076494; PubMed Central PMCID: PMCPMC4468154.
  13. 13. Kaplan BS, DeBeauchamp J, Stigger-Rosser E, Franks J, Crumpton JC, Turner J, et al. Influenza Virus Surveillance in Coordinated Swine Production Systems, United States. Emerging infectious diseases. 2015;21(10):1834–6. pmid:26402228; PubMed Central PMCID: PMCPMC4593420.
  14. 14. Lewis NS, Russell CA, Langat P, Anderson TK, Berger K, Bielejec F, et al. The global antigenic diversity of swine influenza A viruses. Elife. 2016;5:e12217. Epub 2016/04/27. pmid:27113719; PubMed Central PMCID: PMCPMC4846380.
  15. 15. Anderson TK, Campbell BA, Nelson MI, Lewis NS, Janas-Martindale A, Killian ML, et al. Characterization of co-circulating swine influenza A viruses in North America and the identification of a novel H1 genetic clade with antigenic significance. Virus research. 2015;201:24–31. pmid:25701742.
  16. 16. Lewis NS, Anderson TK, Kitikoon P, Skepner E, Burke DF, Vincent AL. Substitutions near the hemagglutinin receptor-binding site determine the antigenic evolution of influenza A H3N2 viruses in U.S. swine. Journal of virology. 2014;88(9):4752–63. pmid:24522915; PubMed Central PMCID: PMCPMC3993788.
  17. 17. Rajao DS, Anderson TK, Kitikoon P, Stratton J, Lewis NS, Vincent AL. Antigenic and genetic evolution of contemporary swine H1 influenza viruses in the United States. Virology. 2018;518:45–54. pmid:29453058.
  18. 18. Nelson MI, Vincent AL. Reverse zoonosis of influenza to swine: new perspectives on the human-animal interface. Trends Microbiol. 2015;23(3):142–53. pmid:25564096; PubMed Central PMCID: PMCPMC4348213.
  19. 19. Nelson MI, Stratton J, Killian ML, Janas-Martindale A, Vincent AL. Continual Reintroduction of Human Pandemic H1N1 Influenza A Viruses into Swine in the United States, 2009 to 2014. Journal of virology. 2015;89(12):6218–26. pmid:25833052; PubMed Central PMCID: PMCPMC4474294.
  20. 20. Nelson MI, Wentworth DE, Culhane MR, Vincent AL, Viboud C, LaPointe MP, et al. Introductions and evolution of human-origin seasonal influenza a viruses in multinational swine populations. Journal of virology. 2014;88(17):10110–9. pmid:24965467; PubMed Central PMCID: PMCPMC4136342.
  21. 21. Rajao DS, Gauger PC, Anderson TK, Lewis NS, Abente EJ, Killian ML, et al. Novel Reassortant Human-Like H3N2 and H3N1 Influenza A Viruses Detected in Pigs Are Virulent and Antigenically Distinct from Swine Viruses Endemic to the United States. Journal of virology. 2015;89(22):11213–22. Epub 2015/08/28. pmid:26311895; PubMed Central PMCID: PMCPMC4645639.
  22. 22. Kitikoon P, Vincent AL, Gauger PC, Schlink SN, Bayles DO, Gramer MR, et al. Pathogenicity and transmission in pigs of the novel A(H3N2)v influenza virus isolated from humans and characterization of swine H3N2 viruses isolated in 2010–2011. Journal of virology. 2012;86(12):6804–14. pmid:22491461; PubMed Central PMCID: PMCPMC3393545.
  23. 23. Ciacci Zanella JR, Vincent AL, Zanella EL, Lorusso A, Loving CL, Brockmeier SL, et al. Comparison of Human-Like H1 (delta-Cluster) Influenza A Viruses in the Swine Host. Influenza research and treatment. 2012;2012:329029. Epub 2012/10/18. pmid:23074664; PubMed Central PMCID: PMCPMC3447287.
  24. 24. Mancera Gracia JC, Van den Hoecke S, Saelens X, Van Reeth K. Effect of serial pig passages on the adaptation of an avian H9N2 influenza virus to swine. PloS one. 2017;12(4):e0175267. pmid:28384328; PubMed Central PMCID: PMCPMC5383288.
  25. 25. Abente EJ, Gauger PC, Walia RR, Rajao DS, Zhang J, Harmon KM, et al. Detection and characterization of an H4N6 avian-lineage influenza A virus in pigs in the Midwestern United States. Virology. 2017;511:56–65. pmid:28841443; PubMed Central PMCID: PMCPMC5623641.
  26. 26. Obadan AO, Kimble BJ, Rajao D, Lager K, Santos JJ, Vincent A, et al. Replication and transmission of mammalian-adapted H9 subtype influenza virus in pigs and quail. The Journal of general virology. 2015;96(9):2511–21. pmid:25986634; PubMed Central PMCID: PMCPMC4635494.
  27. 27. Gao S, Anderson TK, Walia RR, Dorman KS, Janas-Martindale A, Vincent AL. The genomic evolution of H1 influenza A viruses from swine detected in the United States between 2009 and 2016. The Journal of general virology. 2017;98(8):2001–10. pmid:28758634; PubMed Central PMCID: PMCPMC5817270.
  28. 28. United States Department of Agriculture. Influenza A Virus in Swine Surveillance (accessed 24 January 2018)2017 [updated 07/31/2015; cited 2017 March]. Available from:
  29. 29. Bolton MJ, Abente EJ, Venkatesh D, Stratton JA, Zeller M, Anderson TK, et al. Antigenic evolution of H3N2 influenza A viruses in swine in the United States from 2012 to 2016. Influenza and other respiratory viruses. 2018. pmid:30216671.
  30. 30. Rajao DS, Perez DR. Universal Vaccines and Vaccine Platforms to Protect against Influenza Viruses in Humans and Agriculture. Frontiers in microbiology. 2018;9:123. pmid:29467737; PubMed Central PMCID: PMCPMC5808216.
  31. 31. Vincent AL, Perez DR, Rajao D, Anderson TK, Abente EJ, Walia RR, et al. Influenza A virus vaccines for swine. Veterinary microbiology. 2017;206:35–44. pmid:27923501.
  32. 32. USDA. Swine 2012. Part II: Reference of Swine Health and Health Management in the United States, 2012 Fort Collins, CO: USDA–APHIS–VS–CEAH–NAHMS; 2017 [updated 02/23/2017; cited 2017 March]. NAHMS Swine 2012]. Available from:
  33. 33. Chamba Pardo FO, Schelkopf A, Allerson M, Morrison R, Culhane M, Perez A, et al. Breed-to-wean farm factors associated with influenza A virus infection in piglets at weaning. Preventive veterinary medicine. 2018;161:33–40. pmid:30466656
  34. 34. Mughini-Gras L, Beato MS, Angeloni G, Monne I, Buniolo F, Zuliani F, et al. Control of a Reassortant Pandemic 2009 H1N1 Influenza Virus Outbreak in an Intensive Swine Breeding Farm: Effect of Vaccination and Enhanced Farm Management Practices. PLoS Curr. 2015;7. pmid:25932349; PubMed Central PMCID: PMCPMC4405187.
  35. 35. Corzo CA, Gramer M, Kuhn M, Mohr M, Morrison R. Observations regarding influenza A virus shedding in a swine breeding farm after mass vaccination. Journal of Swine Health and Production. 2012;20(6):283–9. WOS:000310652600006.
  36. 36. Choi YK, Goyal SM, Joo HS. Evaluation of transmission of swine influenza type A subtype H1N2 virus in seropositive pigs. Am J Vet Res. 2004;65(3):303–6. pmid:15027676.
  37. 37. Loeffen WL, Heinen PP, Bianchi AT, Hunneman WA, Verheijden JH. Effect of maternally derived antibodies on the clinical signs and immune response in pigs after primary and secondary infection with an influenza H1N1 virus. Veterinary immunology and immunopathology. 2003;92(1–2):23–35. pmid:12628761.
  38. 38. Blaškovič D, Jamrichová O, Rathova V, Kočišková D, Kaplan M. Experimental infection of weanling pigs with A/Swine influenza virus: 2. The shedding of virus by infected animals. Bulletin of the World Health Organization. 1970;42(5):767. pmid:5311062
  39. 39. Deblanc C, Hervé S, Gorin S, Cador C, Andraud M, Quéguiner S, et al. Maternally-derived antibodies do not inhibit swine influenza virus replication in piglets but decrease excreted virus infectivity and impair post-infectious immune responses. Veterinary microbiology. 2018.
  40. 40. Allerson M, Deen J, Detmer SE, Gramer MR, Joo HS, Romagosa A, et al. The impact of maternally derived immunity on influenza A virus transmission in neonatal pig populations. Vaccine. 2013;31(3):500–5. pmid:23174202; PubMed Central PMCID: PMCPMC3534892.
  41. 41. Diaz A, Marthaler D, Corzo C, Munoz-Zanzi C, Sreevatsan S, Culhane M, et al. Multiple Genome Constellations of Similar and Distinct Influenza A Viruses Co-Circulate in Pigs During Epidemic Events. Sci Rep. 2017;7(1):11886. pmid:28928365; PubMed Central PMCID: PMCPMC5605543.
  42. 42. Rose N, Herve S, Eveno E, Barbier N, Eono F, Dorenlor V, et al. Dynamics of influenza A virus infections in permanently infected pig farms: evidence of recurrent infections, circulation of several swine influenza viruses and reassortment events. Veterinary research. 2013;44:72. pmid:24007505; PubMed Central PMCID: PMCPMC3846378.
  43. 43. Pileri E, Martin-Valls GE, Diaz I, Allepuz A, Simon-Grife M, Garcia-Saenz A, et al. Estimation of the transmission parameters for swine influenza and porcine reproductive and respiratory syndrome viruses in pigs from weaning to slaughter under natural conditions. Preventive veterinary medicine. 2017;138:147–55. pmid:28237230.
  44. 44. Kyriakis CS, Rose N, Foni E, Maldonado J, Loeffen WL, Madec F, et al. Influenza A virus infection dynamics in swine farms in Belgium, France, Italy and Spain, 2006–2008. Veterinary microbiology. 2013;162(2–4):543–50. pmid:23201246.
  45. 45. Loeffen WL, Hunneman WA, Quak J, Verheijden JH, Stegeman JA. Population dynamics of swine influenza virus in farrow-to-finish and specialised finishing herds in the Netherlands. Veterinary microbiology. 2009;137(1–2):45–50. pmid:19181461.
  46. 46. Loeffen WLA, Nodelijk G, Heinen PP, van Leengoed LAMG, Hunneman WA, Verheijden JHM. Estimating the incidence of influenza-virus infections in Dutch weaned piglets using blood samples from a cross-sectional study. Veterinary microbiology. 2003;91(4):295–308. WOS:000180328400001. pmid:12477644
  47. 47. Simon-Grife M, Martin-Valls GE, Vilar MJ, Busquets N, Mora-Salvatierra M, Bestebroer TM, et al. Swine influenza virus infection dynamics in two pig farms; results of a longitudinal assessment. Veterinary research. 2012;43:24. pmid:22452923; PubMed Central PMCID: PMCPMC3353254.
  48. 48. Ferreira JB, Grgic H, Friendship R, Nagy E, Poljak Z. Influence of microclimate conditions on the cumulative exposure of nursery pigs to swine influenza A viruses. Transboundary and emerging diseases. 2018;65(1):e145–e54. pmid:28940764.
  49. 49. Ferreira JB, Grgic H, Friendship R, Wideman G, Nagy E, Poljak Z. Longitudinal study of influenza A virus circulation in a nursery swine barn. Veterinary research. 2017;48(1):63. pmid:29017603; PubMed Central PMCID: PMCPMC5634873.
  50. 50. Zhang J, Harmon KM. RNA extraction from swine samples and detection of influenza A virus in swine by real-time RT-PCR. In: Spackman E, editor. Animal Influenza Virus. Methods Mol Biol. 1161. New York: Humana Press; 2014. p. 277–93.
  51. 51. Zhang J, Gauger PC. Isolation of Swine Influenza Virus in Cell Cultures and Embryonated Chicken Eggs. In: Spackamn E, editor. Animal Influenza Virus. Methods in Molecular Biology. 1161. New York, NY: Humana Press; 2014. p. 265–76.
  52. 52. Rutvisuttinunt W, Chinnawirotpisan P, Simasathien S, Shrestha SK, Yoon IK, Klungthong C, et al. Simultaneous and complete genome sequencing of influenza A and B with high coverage by Illumina MiSeq Platform. Journal of virological methods. 2013;193(2):394–404. pmid:23856301.
  53. 53. Bao Y, Bolotov P, Dernovoy D, Kiryutin B, Zaslavsky L, Tatusova T, et al. The influenza virus resource at the National Center for Biotechnology Information. Journal of virology. 2008;82(2):596–601. pmid:17942553; PubMed Central PMCID: PMCPMC2224563.
  54. 54. Kearse M, Moir R, Wilson A, Stones-Havas S, Cheung M, Sturrock S, et al. Geneious Basic: an integrated and extendable desktop software platform for the organization and analysis of sequence data. Bioinformatics. 2012;28(12):1647–9. pmid:22543367; PubMed Central PMCID: PMCPMC3371832.
  55. 55. Bao Y, Bolotov P, Dernovoy D, Kiryutin B, Tatusova T. FLAN: a web server for influenza virus genome annotation. Nucleic acids research. 2007;35(Web Server issue):W280–4. Epub 2007/06/05. pmid:17545199; PubMed Central PMCID: PMCPMC1933127.
  56. 56. Zhang Y, Aevermann BD, Anderson TK, Burke DF, Dauphin G, Gu Z, et al. Influenza Research Database: An integrated bioinformatics resource for influenza virus research. Nucleic acids research. 2017;45(D1):D466–D74. pmid:27679478; PubMed Central PMCID: PMCPMC5210613.
  57. 57. Caton AJ, Brownlee GG, Yewdell JW, Gerhard W. The antigenic structure of the influenza virus A/PR/8/34 hemagglutinin (H1 subtype). Cell. 1982;31(2 Pt 1):417–27. pmid:6186384.
  58. 58. Gerhard W, Yewdell J, Frankel ME, Webster R. Antigenic structure of influenza virus haemagglutinin defined by hybridoma antibodies. Nature. 1981;290(5808):713–7. pmid:6163993.
  59. 59. Huang JW, Lin WF, Yang JM. Antigenic sites of H1N1 influenza virus hemagglutinin revealed by natural isolates and inhibition assays. Vaccine. 2012;30(44):6327–37. pmid:22885274.
  60. 60. Lubeck MD, Gerhard W. Topological mapping of antigenic sites on the influenza A/PR/8/34 virus hemagglutinin using monoclonal antibodies. Virology. 1981;113(1):64–72. WOS:A1981MC21600006. pmid:6791373
  61. 61. Xu H, Yang Y, Wang S, Zhu R, Qiu T, Qiu J, et al. Predicting the Mutating Distribution at Antigenic Sites of the Influenza Virus. Sci Rep. 2016;6:20239. pmid:26837263; PubMed Central PMCID: PMCPMC4738307.
  62. 62. Kitikoon P, Gauger PC, Vincent AL. Hemagglutinin Inhibition Assay with Swine Sera. In: Spackman E, editor. Animal Influenza Virus, Second Edition. Methods in Molecular Biology. 11612014. p. 295–301.
  63. 63. R Core Team. R: A language and environment for statistical computing. 3.4.2 ed. Vienna, Austria: R Foundation for Statistical Computing; 2017. p. Statisitical Software.
  64. 64. StataCorp. Stata Statistical Software: Release 12. 12 ed. College Station, TX: StataCorp LP; 2011.
  65. 65. SAS. SAS 9.4 ®. 9.4m5 ed. Cary, NC, USA: SAS Institute Inc.; 2017. p. Statistical Software.
  66. 66. Cador C, Herve S, Andraud M, Gorin S, Paboeuf F, Barbier N, et al. Maternally-derived antibodies do not prevent transmission of swine influenza A virus between pigs. Veterinary research. 2016;47(1):86. pmid:27530456; PubMed Central PMCID: PMCPMC4988049.
  67. 67. Corzo CA, Allerson M, Gramer M, Morrison RB, Torremorell M. Detection of airborne influenza a virus in experimentally infected pigs with maternally derived antibodies. Transboundary and emerging diseases. 2014;61(1):28–36. pmid:22827737.
  68. 68. Renshaw HW. Influence of antibody-mediated immune suppression on clinical, viral, and immune responses to swine influenza infection. Am J Vet Res. 1975;36(1):5–13. pmid:123140.
  69. 69. Kitikoon P, Nilubol D, Erickson BJ, Janke BH, Hoover TC, Sornsen SA, et al. The immune response and maternal antibody interference to a heterologous H1N1 swine influenza virus infection following vaccination. Veterinary immunology and immunopathology. 2006;112(3–4):117–28. pmid:16621020.
  70. 70. Niewiesk S. Maternal antibodies: clinical significance, mechanism of interference with immune responses, and possible vaccination strategies. Front Immunol. 2014;5:446. pmid:25278941; PubMed Central PMCID: PMCPMC4165321.
  71. 71. Sandbulte MR, Platt R, Roth JA, Henningson JN, Gibson KA, Rajao DS, et al. Divergent immune responses and disease outcomes in piglets immunized with inactivated and attenuated H3N2 swine influenza vaccines in the presence of maternally-derived antibodies. Virology. 2014;464–465:45–54. pmid:25043588.
  72. 72. Rajao DS, Walia RR, Campbell B, Gauger PC, Janas-Martindale A, Killian ML, et al. Reassortment between Swine H3N2 and 2009 Pandemic H1N1 in the United States Resulted in Influenza A Viruses with Diverse Genetic Constellations with Variable Virulence in Pigs. Journal of virology. 2017;91(4). pmid:27928015; PubMed Central PMCID: PMCPMC5286888.
  73. 73. Nelson MI, Viboud C, Vincent AL, Culhane MR, Detmer SE, Wentworth DE, et al. Global migration of influenza A viruses in swine. Nat Commun. 2015;6:6696. Epub 2015/03/31. pmid:25813399; PubMed Central PMCID: PMCPMC4380236.
  74. 74. Lorusso A, Ciacci-Zanella JR, Zanella EL, Pena L, Perez DR, Lager KM, et al. Polymorphisms in the haemagglutinin gene influenced the viral shedding of pandemic 2009 influenza virus in swine. The Journal of general virology. 2014;95(Pt 12):2618–26. pmid:25127710.
  75. 75. Lorusso A, Vincent AL, Gramer MR, Lager KM, Ciacci-Zanella JR. Contemporary epidemiology of North American lineage triple reassortant influenza A viruses in pigs. In: Richt JA, Webby R, editors. Swine Influenza. Current Topics in Microbiology and Immunology. Berlin, Germany: Springer Berlin Heildelberg; 2013. p. 113–31.
  76. 76. Harris KA, Freidl GS, Munoz OS, von Dobschuetz S, De Nardi M, Wieland B, et al. Epidemiological Risk Factors for Animal Influenza A Viruses Overcoming Species Barriers. Ecohealth. 2017;14(2):342–60. pmid:28523412.
  77. 77. Kitikoon P, Nelson MI, Killian ML, Anderson TK, Koster L, Culhane MR, et al. Genotype patterns of contemporary reassorted H3N2 virus in US swine. The Journal of general virology. 2013;94(Pt 6):1236–41. pmid:23695819.
  78. 78. Anderson TK, Nelson MI, Kitikoon P, Swenson SL, Korslund JA, Vincent AL. Population dynamics of cocirculating swine influenza A viruses in the United States from 2009 to 2012. Influenza and other respiratory viruses. 2013;7 Suppl 4(s4):42–51. pmid:24224819; PubMed Central PMCID: PMCPMC5655888.
  79. 79. Nelson MI, Gramer MR, Vincent AL, Holmes EC. Global transmission of influenza viruses from humans to swine. The Journal of general virology. 2012;93(Pt 10):2195–203. pmid:22791604; PubMed Central PMCID: PMCPMC3541789.
  80. 80. Sandbulte MR, Spickler AR, Zaabel PK, Roth JA. Optimal Use of Vaccines for Control of Influenza A Virus in Swine. Vaccines (Basel). 2015;3(1):22–73. pmid:26344946; PubMed Central PMCID: PMCPMC4494241.
  81. 81. Na W, Lyoo KS, Song EJ, Hong M, Yeom M, Moon H, et al. Viral dominance of reassortants between canine influenza H3N2 and pandemic (2009) H1N1 viruses from a naturally co-infected dog. Virology journal. 2015;12(1):134. pmid:26336880; PubMed Central PMCID: PMCPMC4559257.
  82. 82. Brown VL, Drake JM, Barton HD, Stallknecht DE, Brown JD, Rohani P. Neutrality, cross-immunity and subtype dominance in avian influenza viruses. PloS one. 2014;9(2):e88817. pmid:24586401; PubMed Central PMCID: PMCPMC3934864.
  83. 83. Skowronski DM, Janjua NZ, De Serres G, Purych D, Gilca V, Scheifele DW, et al. Cross-reactive and vaccine-induced antibody to an emerging swine-origin variant of influenza A virus subtype H3N2 (H3N2v). The Journal of infectious diseases. 2012;206(12):1852–61. pmid:22872731.
  84. 84. Sriwilaijaroen N, Kondo S, Yagi H, Takemae N, Saito T, Hiramatsu H, et al. N-glycans from porcine trachea and lung: predominant NeuAcalpha2-6Gal could be a selective pressure for influenza variants in favor of human-type receptor. PloS one. 2011;6(2):e16302. pmid:21347401; PubMed Central PMCID: PMCPMC3036579.
  85. 85. Laurie KL, Guarnaccia TA, Carolan LA, Yan AW, Aban M, Petrie S, et al. Interval Between Infections and Viral Hierarchy Are Determinants of Viral Interference Following Influenza Virus Infection in a Ferret Model. The Journal of infectious diseases. 2015;212(11):1701–10. pmid:25943206; PubMed Central PMCID: PMCPMC4633756.