Experimental H1N1pdm09 infection in pigs mimics human seasonal influenza infections.

Pigs are anatomically, genetically and physiologically comparable to humans and represent a natural host for influenza A virus (IAV) infections. Thus, pigs may represent a relevant biomedical model for human IAV infections. We set out to investigate the systemic as well as the local immune response in pigs upon two subsequent intranasal infections with IAV H1N1pdm09. We detected decreasing numbers of peripheral blood lymphocytes after the first infection. The simultaneous increase in the frequencies of proliferating cells correlated with an increase in infiltrating leukocytes in the lung. Enhanced perforin expression in αβ and γδ T cells in the respiratory tract indicated a cytotoxic T cell response restricted to the route of virus entry such as the nose, the lung and the bronchoalveolar lavage. Simultaneously, increasing frequencies of CD8αα expressing αβ T cells were observed rapidly after the first infection, which may have inhibited uncontrolled inflammation in the respiratory tract. Taking together, the results of this study demonstrate that experimental IAV infection in pigs mimics major characteristics of human seasonal IAV infections.

Introduction

Influenza A virus (IAV) infections cause low mortality but high morbidity in humans worldwide annually, while pandemics that occur at irregular intervals may have a disastrous impact on global human health [1-3]. Because newly emerging IAV were often of swine origin or arose from reassortments in the pig as mixing vessel [4-7], pigs have increasingly been evaluated as a biomedical model for human influenza [8]. Pigs are anatomically, physiologically and genetically similar to humans indicating a closer mimic of the situation in humans than rodent models. Studies on the immune response in pigs have long been biased by the lack of specific reagents. Especially the variety of antibodies available for the pig is still far lower to that of mice. As a result, the knowledge of the porcine immune response in general is much smaller compared to that of mice. Despite these disadvantages, pigs have decisive advantages as a model for human IAV infection: they are themselves a natural host for influenza viruses and source of new IAV pathogenic for humans. Thus, understanding the immune response of pigs upon IAV infection and protection by tailored vaccines would assist in the burden on human health by minimizing the emergence of new viruses in swine and zoonotic transmission [4, 5, 9]. Initially, these approaches primarily aimed at identification of antigen preparations that elicit a protective antibody response against surface glycoproteins [10-13], which are effective in controlling IAV infection. However, constant antigenic drift requires an incessant update of the vaccines to match circulating strains. Thus and by expanding the number of reagents for porcine immunological analyses, in the past two decades research increasingly focused on analysis of the immune response against conserved antigenic regions unlikely to change extensively, including cellular immunity. These studies provided evidence for the importance of the cellular immune response in eliminating IAV, also in pigs. These studies have in common that they either only investigated blood-derived cells [14-16] or analyzed the systemic and local immune response only at a very early [17] or late time points of infection [15]. The situation is further complicated by the use of different IAV strains. Recent work recorded for the first time the kinetics of T cell responses and their phenotyping, which supported the previous assumption of the induction of IAV-specific CD4+ and CD8+ T cells [18, 19]. This is in line with several publications reporting IAV-specific CD8+ T cell responses in humans associated with cytolytic [20, 21] as well as memory characteristics [22]. Further, IAV-specific memory T cells were reported to reside in human lungs [23]. Altogether, these porcine studies provided further evidence for similarities of the IAV immune response in pigs and humans [22, 24, 25]. However, it is important to note that the prominent porcine populations of CD4+/CD8+ double positive T cells [26] as well as the high number of peripheral γδ T [27] cells are virtually absent in humans [28, 29], representing a major difference. Besides the numerical difference, the functionality of the two cell populations is comparable in both human and swine. CD4+/CD8+ double positive T cells are mature effector cells with memory characteristics that rapidly mount antigen-specific responses upon antigenic challenge [18, 30]. Besides acting as innate immune cells via pattern recognition receptors and direct killing of infected cells, γδ T cells do play a major role in antigen processing and presentation in human and swine [31, 32]. The two most detailed characterizations of T cell dynamics in influenza-infected pigs were performed by Gerner’s group [18, 19]. They reported increased capacity of CD4+/CD8α+ T cells to co-produce IFN-γ, TNF-α and IL-2 as well as the appearance of blood-derived IAV-specific CD8β+ T cells [18]. Further investigations revealed increased frequencies of IFN-γ and/or TNF-α producing, influenza-specific CD4+ and CD8+ T cells in the lungs of infected pigs, resulting in an accumulation of memory cells in the lungs at 6 weeks post infection [19]. Although these porcine IAV studies report a pronounced involvement of the cellular immune response, a global analysis and comparison of immune cells and their functions along the route of entry (nose, lung and lung lymph node) as well as the systemic responses (blood) after infection with H1N1pdm09 were still missing. To ascertain the value of the pig as a reliable model mimicking human IAV infection, we compared the immune response in pigs after a natural IAV infection without inducing major clinical signs, as usually occurs in seasonal human influenza.

Material and methods

Ethical statement and study design

All animal experiments were approved by the State Office for Agriculture, Food Safety and Fishery in Mecklenburg-Western Pomerania (LALFF M-V) with reference numbers 7221.3-1-035/17.

In total 29 four-week old German landrace pigs were obtained from a commercial high health status herd (BHZP-Basiszuchtbetrieb Garlitz-Langenheide, Germany). This farm is free from the following diseases or pathogens: Pseudorabies, classical swine fever virus (CSFV), Porcine Reproductive and Respiratory Syndrome Virus (PRRSV), Actinobacillus pleuropneumoniae, Mycoplasma hypopneumoniae, ascaris, mange, Brachyspira hyodysenteriae and salmonella category I. Vaccination program does not include vaccination against influenza viruses but the following vaccines were administered: Sows were vaccinated against porcine circovirus type 2 (once), Erysipelothrix rhusiopathiae/Porcine parvovirus (twice), salmonella (twice) and Haemophilus parasuis (twice). Piglets were vaccinated against PCV2 once and Mycoplasma hypopneumoniae twice. Piglets for this study were kept under BSL2 conditions in the animal facilities of the Friedrich-Loeffler-Institut, Greifswald-Riems. IAV infections were performed twice during study period (Table 1), whereby first infection occurred three weeks after transport to our facility by intranasal administration of 2 ml virus suspension (106 TCID50/ml) using mucosal atomization devices (MAD) (Wolfe Tory Medical, USA) with 26 animals. A second infection was performed on day 21 after the first infection. Three animals were mock infected with medium only and served as controls. Blood samples from the same six randomly selected animals were taken on day 0, 2, 4, 7, 14, 21 (prior to second infection), 22, 25 and 31 after first infection for kinetics of blood cells. After euthanasia with Release® (IDT, Germany), necropsy was performed on five animals on day 4, 7, 21 and 25 post first infection. Control animals were euthanized on day 30 after first mock-infection.

Table 1

Summary of sampling days and animals during study.

	d01st infection	d2pi	d4pi	d7pi	d14pi	d21pi2nd infection	d22pi	d25pi	d31pi
Blood samples	6	6	6	6	6	6	6	6	6
Organ samples			5	5		5		5	3

Blood samples were taken from the same six animals, randomly chosen on day 0. Three control animals underwent necropsy on day 30 post infection.

Virus

Influenza virus A/Bayern/74/2009 (hereafter referred to as H1N1pdm09) was propagated on Madin-Darby canine kidney cells (MDCKII) in MEM supplemented with 0.56% bovine serum albumin, 100 U/ml Penicillin, 100 μg/ml Streptomycin and 2 μg/ml L-1-Tosylamide-2-phenylethyl chloromethyl ketone (TPCK)-treated trypsin (Sigma-Aldrich, USA). For viral titration by TCID50 assay, serial ten-fold dilutions of virus suspensions were prepared, added to MDCKII cells in 96-well plates, and incubated for three days at 37°C and 5% CO2. Cytopathic effect was examined microscopically. Titers were calculated according to Spearman-Kärber [33, 34]. An acute IAV infection of pigs acquired prior to the study was excluded by real-time PCR (AgPath.ID™ One-Step RT-PCR Kit, Applied Biosystems, USA) of nasal swabs immediately before transport to the experimental facility (modified from [35]).

Sample preparation

Necropsy and pathological gross examination was performed according to standard guidelines under BSL3** conditions. For extracorporeal bronchoalveolar lavage (BAL), the left main bronchus was cut with sterile scissors and 200 ml of sterile PBS solution was injected with a syringe through the main bronchus into the left lung lobe, which was then kneaded softly. BAL fluid was recovered by syringe.

For flow cytometric analyses organ samples from the following organs were taken during necropsy and stored in ice-cold PBS until further use: mucosa of nasal cavity, lung tissue (lobus dexter medius) and lung lymph node (nodi lymphatici tracheobronchiales inferiores).

Specimen from the following organs were collected for histopathology and fixed in 4% neutral buffered formaldehyde: nasal cavity with conchae, trachea, lungs (lobus sinister cranialis pars cranialis and caudalis, lobus sinister caudalis, lobus dexter cranialis, lobus dexter medius, lobus dexter accessorius, lobus dexter caudalis), lymph node from the lymphocentrum bronchiale.

Lungs were scored for macroscopically detectable atelectasis (reddish-tan, consolidated, lobular to lobar parenchyma) and scored as follows: (shown as percentage of the total parenchyma analyzed for each pulmonary lobe): 0 = no atelectasis, 1 = mild atelectasis (0–30%), 2 = moderate atelectasis (30–60%), 3 = severe atelectasis (60–100%). The max atelectasis score of all seven scored pulmonary lobes was taken to set up one gross lesion score for each individual pig. For histopathological examination, formaldehyde-fixed tissue samples were embedded in paraffin and cut at 3μm. The sections were mounted on Super-Frost-Plus-Slides (Carl Roth GmbH, Karlsruhe, Germany) and stained with hematoxylin-eosin for light microscopical examination using a Zeiss Axio Scope.A1 microscope equipped with 5x, 10x, 20x, and 40x N-ACHROPLAN objectives (Carl Zeiss Microscopy GmbH, Jena, Germany). All tissue sections were scored blind and investigated for signs and severity levels of inflammation (rhinitis, tracheitis, bronchointerstial pneumonia) as follows: 0 = no inflammation, 1 = mild inflammation, 2 = moderate inflammation, 3 = severe inflammation. The final score for each organ of an individual animal was raised on the basis of the max value of the respective scores.

Immunohistochemistry was performed to detect Influenza A virus antigen in paraffin embedded tissue sections using a mouse monoclonal antibody against the matrixprotein of influenza A virus (M21C64R3, ATCC, Manassas, VA) as previously described [36]. Briefly, tissue sections were dewaxed and rehydrated, and endogenous peroxidase was blocked with 3% H2O2 (Merck, Darmstadt, Germany) for 10 min. After demasking of antigens with 10mM Na-citrate buffer (pH = 6, 700 W) for 20 min in a microwave oven, sections were incubated with undiluted normal goat serum for 30 min at room temperature to block nonspecific binding sites. Thereafter, section were incubated with anti-Influenza A matrix protein antibodies, diluted 1: 200 in Tris-buffered saline (TBS) at 4°C overnight. This incubation was followed by a biotinylated goat anti-mouse IgG (1:200, LINARIS biologische Produkte, Dossenheim, Germany) and the avidin-biotin-peroxidase complex (Vector Laboratories, Burlingame, CA) for 30 min each at room temperature. Positive antigen-antibody reactions were visualized by incubation with AEC-substrate (Dako, Hamburg, Germany) for 10 min. Sections were washed with deionized water and counterstained with Mayer`s hemalaun for 30 s. Sections that stained positive for IAV matrix protein were investigated for cell-specific viral antigen localization and scored for viral antigen distribution in each inspected organ: 0 = negative, 1 = focal to oligofocal, 2 = multifocal, 3 = diffuse as previously described [37]. For each organ the max values of all scores were taken into analysis.

Cell preparation and antibody staining for flow cytometric analysis

Whole blood was diluted with PBS 1:10 and overall number of white blood cells and different leukocyte subpopulations were determined by blood counting device. Flow cytometric analysis of single cell suspensions from mucosa of nasal cavity, BAL, lung tissue and lung lymph node were performed. Cell from BAL fluid were enriched by centrifugation and discarding the supernatant. Leukocytes from mucosa of nasal cavity, lung lymph node and spleen were prepared by mechanical disruption on metal sieves with plungers and washed with PBS. For isolation of leukocytes from the lung, tissue was minced with scissors, resuspended in PBS-EDTA supplemented with 100 μM CaCl2, and digested with Collagenase D (1 mg/ml; Sigma-Aldrich) for 40 min at 37°C. After pressing the digested tissue gently through a cell strainer with a plunger, remaining tissue was removed by short centrifugation.

Cell pellets were suspended in FACS buffer for flow cytometric antibody staining. Antibodies used in this study are listed in S1 Table. Unless otherwise stated, all incubation steps were carried out at 4 °C in the dark for 15 min in case of extracellular staining and for 30 min for intracellular staining. Staining of whole blood required erythrocyte lysis after surface staining by conventional lysis buffer (1.55 M NH4Cl, 100 mM KHCO3, 12.7 mM Na4EDTA, pH 7.4, in A. dest.). For intracellular staining, the True-Nuclear™ Transcription Factor Buffer Set was used according to manufacturer’s instructions (BioLegend, Germany).

Software and statistics

Flow cytometric analyses were run on BD FACS CantoII with BD FACS DIVA Software and analyzed using FlowJo Software. GraphPad Prism was used to visualize data and perform statistical analyses. All animal groups examined on days 4, 7, 21, 25 and 30 (control) were analyzed by the nonparametric Kruskal-Wallis test followed by pairwise Dunn’s post hoc tests compared to control. For blood analyses same tests were used but post hoc test was compared to day 0. Statistical significance was designated as p ≤ 0.05 indicated by an asterisk (*) in the graphs.

Results

Intranasal infection of pigs with H1N1pdm09 induced macroscopic and microscopic lesions in the lungs

After intranasal primary IAV infection, multifocal, reddish-tan consolidated areas (pulmonary atelectasis) of different sizes were macroscopically observed in inoculated animals after 4, 7 and 21 days (Fig 1A), mainly in the lobus dexter medius and in the lobus sinister cranialis pars caudalis (Fig 1B). 4 dpi, the animals reached the highest atelectasis score compared to pigs which were analyzed after 25 dpi (Fig 1C). One control pig showed a minimal, focal atelectasis in the lobus dexter cranialis.

Fig 1

Gross pathologic changes after macroscopic investigation of lungs from H1N1pdm09-infected pigs.

At indicated time points, three to five animals were subjected to necropsy for macroscopic investigation of artelectasis. (A) Lung from a pig inoculated with H1N1, 4 dpi. Acute, diffuse atelectasis of the lobus dexter medius (arrow). (B) Frequency distribution of macroscopic lesions in different lung lobes. (C) Atelectasis scores after 4, 7, 21 and 25 days of H1N1-inoculated and mock-infected animals. l. = lobus; x in graph axis indicates infection.

Inflammatory changes were detected in the nasal mucosa, trachea and lung. Results from histopathological investigations of nasal mucosa and lungs are summarized in Fig 2. Starting at 4 dpi pigs showed mild, focal, necrotizing rhinitis with loss of epithelial cells (Fig 2 left panel) and IAV matrix protein-positive respiratory epithelial cells within the lesions. Mild, focal, subacute, lymphohistiocytic rhinitis have been observed 7, 21 and 25 dpi. Until 25 dpi inflammation decreased constantly whereas control pigs were free of rhinitis. One infected pig showed mild, necrotizing tracheitis at 4 dpi compared to all other infected and control pigs, which lacked similar lesions. Lung lesions were mainly localized in bronchi, bronchioles and bronchioloalveolar transition zone leading to mild bronchiolointerstitial pneumonia as shown in Fig 2 (right panel). 4 dpi, mild necrosis and loss of bronchial and bronchiolar epithelium was evident in H1N1pdm09 inoculated pigs followed by the infiltration of lymphocytes, macrophages and few neutrophils into the affected tissue (Fig 2C, right panel). At 7 dpi, mild alveolar edema was present whereas necrosis extended to the bronchi-alveolar transition zone (Fig 2E, right panel). At that time, lymphocytes and macrophages increasingly infiltrated the pulmonary interstitium (Fig 2E, right panel), but Influenza A matrix protein was not detectable at any time point later than 4 dpi (Fig 2B, 2D, 2F, 2H and 2J, right panel). 21 dpi, inflammatory cells were still evident (Fig 2G, right panel). Still negative for viral antigen (Fig 2J, right panel), the amount of infiltrating inflammatory cells slightly decreased at 25 dpi (Fig 2I, right panel). Data from histopathological scoring are summarized in Fig 3. As indicated, IAV matrixprotein was only detectable 4 dpi in the nose, trachea and lung (Fig 3A). At 7 dpi, infected animals showed the highest inflammation score in the nose and lung which then slightly decreased and remained constant until the end of the experiment (Fig 3B). Of note, a moderate significant positive correlation (Spearmann r = 0.464; p<0.0001) was found between macroscopic (atelectasis) and microscopic lesions in the lung (p < 0.0001) (Fig 4).

Fig 2

Histopathology from nose (left panel) and lung (right panel) of H1N1-infected pigs. At indicated time points, three to five animals were subjected to necropsy. Lungs, trachea and conchae were fixed in 4% formaldehyde, embedded in paraffin and cut at 3μm. Hematoxylin-Eosin (A, C, E, G, I) and anti-Influenza matrixprotein immunohistochemistry (B, D, F, H, J) were performed on nose and lung tissue. A-B) mock-control. (C-D) 4 dpi. (E-F) 7 dpi. (G-H) 21 dpi. (I-J) 25 dpi. White arrows: infiltration of inflammatory cells; black arrows: Influenza A matrix protein-positive cells; arrowheads: flattening and loss of epithelial cells; asterisks: necrotic lung tissue.

Fig 3

IAV matrixprotein score (A) and inflammation score (B) obtained from the nose, trachea and lung. At indicated time points, three to five animals were subjected to necropsy. Lungs, trachea and conchae were fixed in 4% formaldehyde, embedded in paraffin and cut at 3μm. Hematoxylin-eosin staining was used to determine inflammatory score, immunohistochemistry allowed the detection of IAV matrixprotein. x in graph axis indicates infection.

Fig 4

Correlation between macroscopic and microscopic lesions.

Correlation between scores of atelectasis and simultaneous inflammatory processes in the lung obtained by histopathological investigation. Spearman r = 0,4642 (p < 0,0001).

IAV H1N1pdm09 infection led to a shift in distribution of blood immune cell subsets but not to leucopenia

Counts of total and different subpopulations of white blood cells did not change significantly during the course of infection (Fig 5). A slight decrease in total cell numbers was observed two days after the first infection but they recovered until day four and stayed at a comparable number until day 14 post first infection. On day 21, prior to second infection, cell number was slightly elevated, stayed at a comparable level the following day and returned to cell numbers comparable to those on day 0 and remained stable until the end of the study (Fig 5A).

Fig 5

Counts of peripheral blood leukocytes after infection with H1N1pdm09.

At indicated time points, blood was collected from the same six pigs, randomly chosen on day 0. Blood counting device revealed total count of A) white blood cells and B) myeloid cells (monocytes, dendritic cells and neutrophils). C) Within the granulocytic population frequency of mature granulocytes is indicated by expression of CD14. D) Total count of lymphocytes and E) main subpopulations: αβ T cells, γδ T cells and B cells. x in graph axis indicates infection.

Of the myeloid cells in blood, dendritic cell and neutrophil count decreased initially on day two post infection but their numbers as well as monocyte count increased on day four post infection (Fig 5B). On average, monocyte and neutrophil counts remained stable from day 14 post infection until the end of the study, regardless of a second infection, whereas dendritic cell numbers decreased to a hardly detectable level after the second infection and did not recover until the end of the study (Fig 5B). Although neutrophil counts decreased initially after first infection, the frequency of CD14 expressing cells among them increased after first as well as after second infection (Fig 5C).

Regarding cell number of lymphocytes, we observed a decrease for αβ and γδ T cells, as well as for B cells initially after first infection that lasted until day seven and for the latter until the end of the study (Fig 5D). On day 14 post infection cell numbers of γδ T cells and B cells recovered, but αβ T cells numbers were elevated compared to day 0. After second infection αβ T cells decreased to basal level and remained unchanged for the rest of the study, whereas second infection led to another increase in γδ T cells on day 22. Three days later (day 25) and on day 31 numbers of these cells reflected basal levels (Fig 5D).

Proliferation of CD8+ αβ T cells in the blood was increased upon infection with H1N1pdm09

In line with absolute cell numbers obtained from blood counting device (Fig 5), we also observed a decreased frequency of αβ T cells from lymphocytes both after first and second infection with H1N1pdm09 in blood of pigs (Fig 6A). 14 days after first infection, they returned to normal levels. After the second infection recovery time was faster and original frequencies were reconstituted on day 25 (four days after second infection).

Fig 6

Frequencies of αβ T cells (A), their subpopulations (B) expressing Ki67 (C). At indicated time points, blood was collected from the same six pigs, randomly chosen on day 0. Flow cytometric analyses were performed to determine frequency of CD3+/ γδTCR- T cells (A). Further classification (B) was made based on the expression of CD4+ single positive cells (naive Th cells), CD4+/CD8+ (memory as well as cytotoxic T cells) and CD4-/CD8+ cytolytic T cells. Expression of Ki67 (C) indicated proliferative capacity of cells. x in graph axis indicates infection.

To determine subpopulations, frequencies of T cells expressing CD4 and/or CD8 were distinguished into naïve Th cells (CD4+/CD8-), extrathymic CD4+/CD8+ cells (including memory as well as cytotoxic cells) and cytolytic T cells (CD4-/CD8+) (Fig 6B). Overall frequencies of different subpopulations remained stable until day 14 after first infection. Afterwards frequency of cytolytic T cells in bloodstream increased at the expense of naïve cell frequencies. From day 25 (4 days after second infection) increasing frequencies of double-positive Th cells were detected with further decreasing naïve cells. Frequency of proliferating CD8+ cells–characterized by Ki-67 expression–increased both after first and to a lesser extent after second infection with H1N1pdm09. CD8- αβ T cells did not show signs of proliferation (Fig 6C).

γδ T cells proliferated and increased CD8 expression in the blood after infection with H1N1pdm09

Because γδ T cells do play a major role in pigs during infections, we analyzed these cells and the distribution of different subtypes in the course of IAV infection. Frequencies remained stable during the first infection but were increased immediately after the second infection, which was congruent with decreased frequencies of αβ T cells (Fig 7A).

Fig 7

Frequencies of γδ T cells (A), their subpopulations (B) expressing Ki67 (C). At indicated time points, blood was collected from the same six pigs, randomly chosen on day 0. Flow cytometric analyses were performed to determine frequency of CD3+/ γδTCR+ T cells. Further classification (B) was made based on the expression of CD2 and CD8: CD2-/CD8- = naïve γδ T cells, CD2+/CD8- = activated γδ T cells and CD2+/CD8+ = differentiated effector γδ T cells. Expression of Ki67 (C) indicated proliferative capacity of cells. * = p≤0.05 Kruskal-Wallis test followed by Dunn’s post hoc test compared to day 0. x in graph axis indicates infection.

CD2 and subsequent CD8 expression on the cell surface of γδ T cells resembles activation and maturation steps, the former being observed 14 days after first and one day after second infection with H1N1pdm09 (Fig 7B). Differentiated effector γδ T cells (characterized by simultaneous expression of CD2 and CD8) increased after second infection only. In addition, Ki-67 expressing cells increased among activated and differentiated effector memory T cells after the first infection (Fig 7C). After the second infection, only the latter showed higher frequencies of proliferating cells. Naïve γδ T cells did not show proliferative activity.

Frequency of αβ T cells along the respiratory tract increased after the first infection only and expressed mainly CD8αα homodimers

To investigate the immune response along the proposed route of infection, frequencies and functional properties of αβ T cells from lymphocytes in mucosa from nasal cavity, BAL, lung and lung lymph node were analyzed. Increasing frequencies for these cells could be observed four days after first but not after second infection in mucosa of nasal cavity, BAL, lung tissue and lung lymph node and until day seven after first infection in the former two (Fig 8A). Frequencies were still elevated on day 21 compared to control animals and returned to normal levels on day 25, four days after second infection in all organ samples.

Fig 8

Frequencies of αβ T cells (A) and CD8αα and CD8αβ subpopulations (B) expressing perforin (C and D). At indicated time points, three to five animals were subjected to necropsy. After preparation of single cell suspensions from nasal mucosa, BAL, lung tissue and lung lymph node, flow cytometric analyses were performed to determine frequency of CD3+/ γδTCR- T cells (A). Further classification (B) was made based on the expression of CD8αα homo- or CD8αβ heterodimers. MFI (mean fluorescence intensity) indicated cytolytic activity of CD8αα (C) and CD8αβ (D) T cells. * = p≤0.05 Kruskal-Wallis test followed by Dunn’s post hoc test compared to control. x in graph axis indicates infection.

Differentiation in subtypes expressing either CD8αα homo- or CD8αβ heterodimers revealed that under physiological conditions (control) CD8+ αβ T cells from tissues of the respiratory tract–nose, BAL and lung–are mainly composed of cells expressing the CD8αα homodimers (Fig 8B). In contrast, in the lymph node CD8αβ expressing T cells were the predominant population. After an initial slight drop in nose and BAL four days after first infection, CD8αα expressing cells increased until day seven in all organs, whereby the ratio in the lymph node was completely reversed towards the homodimer expressing CD8+ T cells. On day 21, prior to second infection, ratio of CD8αα to CD8αβ expressing cells was the same as for control animals. Four days after second infection, the previously observed ratio shift was repeated and resembled the ratio on day seven except for cells in the lung lymph node, for which the ratio was the same as on day four (Fig 8B).

In both αβ T cell subsets expression of perforin increased after the first infection

To investigate the functionality of αβ T cells, expression levels of perforin were investigated along the infection route. In all organs, expression of perforin in CD8αα+ cells increased on the fourth day after first infection but returned to levels as observed in control animals already three days later and did not change during the rest of the study (Fig 8C). CD8αβ+ T cells showed a similar pattern, although the level of expression was higher, both under basal (control) as well as in infection conditions (day 4) (Fig 8D). For CD8αβ+ T cells in lung and BAL expression of perforin returned to basal levels on day 7 and remained unchanged. In lung and to a greater extent in the lymph node, expression of perforin was lowest on day 7, returned to basal levels on day 21 but decreased once again after the second infection (Fig 8D).

Frequency of γδ T cells in the nose and CD8 expressing subpopulations were increased after first infection

Because frequencies of γδ T cells remained stable in the blood, we investigated whether there would be changes in organs from the respiratory tract. In the nose, frequency of cells increased slightly over the fourth to seventh day, stayed elevated until day 21 and returned to levels comparable to those in control animals on day 25 (four days after second infection) (Fig 9A). In BAL, frequencies continuously decreased over study period. This also applies to the lymph node, albeit to a lesser extent. In the lung, frequency of γδ T cells halved after first and decreased by 1/3 after second infection (Fig 9A).

Fig 9

Frequencies of γδ T cells (A) and CD8α subpopulation (B) expressing perforin (C). At indicated time points, three to five animals were subjected to necropsy. After preparation of single cell suspensions from nasal mucosa, BAL, lung tissue and lung lymph node, flow cytometric analyses were performed to determine frequency of CD3+/ γδTCR+ T cells (A). The CD8α expressing subpopulation (B) was further tested for intracellular perforin expression (C). * = p≤0.05 Kruskal-Wallis test followed by Dunn’s post hoc test compared to control. x in graph axis indicates infection.

Investigations on the activation status revealed that in the nose around 40% and in BAL, around 60% of γδ T cells already express the CD8α molecule under physiological conditions (Fig 9B). In contrast to cells in BAL, where frequencies decreased by 10%, activated γδ T cells increased to 60% in the nose four days after infection. In lung and lymph node, frequencies increased from 20% to more than 40% on day four after first infection. For nose, lung and the lymph node, frequency of activated γδ T cells returned to basal levels on day seven and stayed there for every other time point (Fig 9B).

Nasal and BAL γδ T cells show increased expression of perforin and/or transcription factors associated with activation

Related to effector functions of γδ T cells in nose perforin expression was increased only at day four after first infection and did not change at any other day of study period (Fig 9C). For γδ T cells in BAL, lung and lung lymph node, expression of effector molecule perforin was slightly increased on the same day.

Because activation of γδ T cells does not necessarily lead to upregulation of CD8 or perforin, the transcription factors T-bet and EOMES of these cells were also investigated in mucosa from the nasal cavity as well as the BAL. Frequency of γδ T cells expressing T-bet only increased in the nose four days after infection, whereas EOMES single-positive cells increased four days after second infection only (Fig 10A). Frequency of cells expressing both transcription factors did not change during study period. In BAL T-bet expressing cells increased both after first and second infection, whereas EOMES as well as double positive cells remained unchanged (Fig 10B).

Fig 10

Frequencies of γδ T cells expressing transcription factors EOMES and/or T-bet.

At indicated time points, three to five animals were subjected to necropsy. After preparation of single cell suspensions from nasal mucosa and BAL, flow cytometric analyses were performed to determine frequency of T-bet+/EOMES-, T-bet+/EOMES+ and T-bet-/EOMES+ γδ T cells in nose (A) and BAL (B). * = p≤0.05 Kruskal-Wallis test followed by Dunn’s post hoc test compared to control. x in graph axis indicates infection.

Discussion

Using two different approaches, continuous peripheral blood cell analysis from the same animals as well as analysis of organ samples from the respiratory tract at specific time points after infection, we characterized in detail the kinetics of the porcine immune response after intranasal infection with H1N1pdm09. In contrast to previous studies reporting fever as well as clinical sings after intranasal infection of pigs with IAV [10, 14, 38, 39], pigs in our study did not develop either. Since different virus strains as well as doses were used, this finding was not unexpected. While a study by Pomorska-Mól et al. reported mild clinical signs without fever in intranasally H1N1-infected pigs they neither observed significant changes in hematological parameters nor in numbers of subsets of T and B cells [40]. However, the decreasing lymphocyte count caused by decreasing numbers of T cells and B cells within the first week after infection, correlated well with the other studies [17, 41]. Khatri et al. detected also increasing frequencies of CD8+ αβ as well as γδ T cells in lungs and lung LN of H1N1-infected pigs [17], which is in line with our findings. We could further show that this also holds true at the site of first contact, the nose. Increased numbers correlated with an increased amount of perforin pointing toward enhanced cytotoxic function. An increase in CD4+ T cells in the lungs, which was reported by two groups that observed high fever and strong clinical signs [14, 17], was not observed in our study.

The overall leukocyte count was only slightly decreased on day two after the first infection, which was mainly due to a decrease of the neutrophil population and to a lesser extent to decreasing numbers of αβ T cells and B cells. Two days later, absolute numbers of WBC were restored to normal levels because of elevated numbers of myeloid cells–monocytes, dendritic cells and neutrophils–whereas numbers of lymphocytes decreased further until day seven. These observations parallel the mild lymphopenia with monocytosis in human patients infected with IAV [42-44], which was proposed as a screening tool for influenza infection [45, 46]. An increase in neutrophils of IAV patients within the first days of infection described by some groups [42, 47, 48] could not be detected in our study. However, we observed an increase in frequency of CD14 expressing cells after both infections. CD14 upregulation is associated with activation in granulocytes [49, 50] and serves as a coreceptor for TLR7 mediated recognition of ssRNA [51]. We detected a higher number of dendritic cells in the blood in infected pigs four days after the first infection with simultaneously increased number of monocytes, which was not reported for human patients infected with H1N1pdm09. This might be because in most of the studies characterizing the immune response in H1N1 patients definite time points are hard to determine e.g. time of the infection. In contrast, the decrease in αβ T cells and B cells in pigs covers a period of four to seven days with recovery until day 14 and is comparable with decreasing frequencies in patients infected with H1N1pdm09 although the period of mild lymphopenia varies between studies [43, 46, 52]. However, after the second infection, recovery was faster compared to the first infection and thus comparable to vaccinated humans [53, 54]. Further, we detected an increased proliferative activity in CD4+/CD8+ T cells (constituting of cytotoxic and memory cells) and cytotoxic (CD8+) subtypes of αβ T cells, which was pronounced after first infection with a lesser increase after second infection resembling a memory response. These observations are in line with the findings in an experimental human influenza experiment that additionally reported those CD4+ T cells to have cytolytic and thus direct antiviral characteristics like perforin expression [55, 56]. Description of prominent CD4+ T cell responses being strong in numbers, by contrast, are primarily associated with severe influenza cases [57].

Given that the H1N1pdm09 infection of the pigs in our setup with MAD leads to an infection that is almost exclusively localized to the respiratory tract, the increase in αβ T cells frequencies in nose and BAL were more prominent starting as early as day 4 post infection and peaking at day 7. Interestingly, in 14 weeks old healthy pigs the ratio of CD8αα to CD8αβ expressing αβ T cells isolated from nasal mucosa, the BAL or lung tissue is shifted towards CD8αα cells with frequencies of 85%, 93% and 85% respectively. This is the first description of the distribution of these two subtypes in respiratory organs of pigs and is in line with finding in humans and mice investigating the role of CD8αα expressing cells among the intraepithelial leukocyte population [58-60]. Because CD8αα is known to be a corepressor of TCR avidity and diminishes activation [61, 62], it is tempting to speculate that infection with H1N1pdm09 leads to activation whilst upregulation of CD8αα inhibits at the same time an excessive immune response. In line with this, due to homeostasis, frequencies of cytolytic CD8αβ T cells decreased further enhancing an anti-inflammatory environment. Several studies in mice and humans do further attribute CD8αα expressing T cells to have memory function [63-65]. Our findings point in the same direction, as CD8αα expression of αβ T cells in the respiratory tract of pigs increased more rapidly after second infection. In control (healthy) pigs, the expression level of perforin per cell within the CD8αα subpopulation in respiratory tract samples is only a third of the expression level in CD8αβ expressing T cells supporting in addition that they do not primarily have cytolytic activity. CD8αα expressing T cells arise mainly from CD8αβ T cells by downregulation of the β-chain and thus diminishing activation [64, 65]. Therefore, it is conceivable that the absence of perforin expression after the second infection is due to inhibition by increased frequencies of CD8αα T cells.

Because lymphocytes in the blood of pigs comprise up to 50% of γδ T cells (~3000 cells/μl blood), which strongly contrasts the maximum of 5% (<500 cells/μl blood) in healthy humans [66], it is challenging to draw conclusions with regard to comparability. Nevertheless, absolute numbers of these cells remained relatively constant throughout the study decreasing only slightly after the first infection but increasing after the second infection at the expense of αβ T cells in the blood. The latter observation might be explained by the different time points of blood sampling: after the first infection, blood was taken at day two whereas after the second infection it was taken on the first day p.i., to gather a potential memory response. The hypothesis is supported by a pronounced increase in proliferative activity in both activated subtypes of γδ T cells after first infection and a subsequent increase of activated γδ T cells in the blood, supporting that this cell population serves as first line defense. The rapid decrease in frequency of proliferative cells might be explained by the recruitment to the site of inflammation (nose and lung), where their frequencies increased as early as day four after infection with a pronounced perforin expression. Frequencies of γδ T cells in lungs of pigs are comparable to those of humans [67] and they are known not only to maintain pulmonary homeostasis [68] but also to efficiently kill IAV infected epithelial cells and macrophages [69, 70]. In line with these findings from human in vitro experiments [70, 71], we observed a notable increase in perforin expression of γδ T cells after first infection in mucosa of nasal cavity constituting an entrance for the virus. Furthermore, concurred increase in frequency of CD8 expressing γδ T cells was observed along the route of entry from nose to lung and lung lymph node, in the latter most probably constituting antigen-presenting cells. Because perforin expression did not increase after the second infection and frequencies of γδ T cells in blood where increased only at day 14 p.i., it is likely that these cells play a major role in recovery from influenza infections as described earlier [29]. Finally, also T-bet expression in γδ T cells increased after both infections in nose and BAL, which is associated with improved recovery from influenza infections [72].

Given that in a natural H1N1 infection in humans the day of infection is not exactly clear, it seems obvious that a day-to-day comparison of patient data with the predefined time points in this study is challenging. Further, for obvious reasons, analyses of lung or respiratory epithelial tissue from patients with mild influenza virus infections are limited. However, transiently decreased numbers of lymphocytes along with increased monocyte counts in the blood seems to be a common feature in mild human and porcine subclinical H1N1pdm09 infections. Further, we could show the presence of and increase in CD8αα expressing αβ as well as CD8α+ γδ T cells in mucosa from respiratory tract after infection in pigs, indicating that these cells have the same dual role as in humans. They do rapidly respond with perforin expression to H1N1pdm09 but simultaneously increase expression of the inhibitory CD8αα molecule to prevent excessive harmful immune responses. Therefore, even though pigs in our study did not show overt clinical signs, underlying immune and pathogenic mechanisms seem to be similar. The results from this study further expand the knowledge of the porcine immune response to pandemic IAV infection and thus, support the use of the pig as a large animal model for human seasonal influenza infections. This offers not only a model for testing the efficacy of new influenza vaccines regarding cellular immune responses but also to expand the model for further investigations of influenza induced pneumonia especially in bacto-viral coinfection scenarios.

Antibodies used in flow cytometric analyses.

(TIF)

Click here for additional data file.

16 Jul 2019

PONE-D-19-15088

Experimental H1N1pdm09 infection in pigs mimics human seasonal influenza infections

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Reviewer #1: 1. Please furnish information on the pre-screen done on the pigs. This should include both qPCR and serology (ELISA) for IAV, PRRSV, PPV, PCV 2, TTV. if animals were negative prior to D0, what was their status at the end of the study?

2. How were the mock and challenge groups housed? could comparative counts and FACS data for mocks also be clearly identified and shown along with challenge groups?

Reviewer #2: The manuscript by Schwaiger and colleagues describes the characterization of swine immune cell population changes following an experimental infection with a human H1N1 influenza virus. The results are of interest because swine are a natural host of influenza viruses and can also play a role in the zoonotic transmission of non-human influenza strains into the human population. In addition, (as stated by the authors), because of comparable sizes, immunity and physiology, pigs are a valuable animal model for evaluating immune responses, vaccines, antivirals, etc. In general, the manuscript is well written and provides detailed methods and results that can serve as a guide to other groups. To be fair, the authors should also point out some important differences between pigs and humans such as the much shorter “childhood” and general lifespan of pigs; important differences in immune cell populations such as the double positive CD4+CD8+ lymphocytes and strikingly different proportions of gamma-delta+ T cells in blood, or the lack of reagents and general knowledge of the pig’s immunity in comparison to other animal models (mice).

The authors should also consider the following specific comments:

Line 68. The animals were obtained from a commercial herd. If known, it would be important to state the serological status (from natural infection and from vaccination) of the mothers, since the presence of maternal antibodies against H1N1 influenza would have a strong impact in the evolution of the infection.

Line 73. Animals were re-infected with the same influenza strain 21 days after the first infection. In the context of modelling influenza in humans, it is not clear what was the purpose of this infection. In humans, re-infection occurs in the following influenza seasons (at least 1 year later) and by an antigenically drifted strain.

Line 122. “…antibody against the anti-matrixprotein…” this sentence is incorrect. It should be either “…antibody against the matrixprotein…” or “…anti-matrixprotein monoclonal antibody…”. Please correct.

Line 128. It might be useful to describe what TBS stands for.

Line 149. In this paragraph there is a sentence repeated “Antibodies used in this study are listed in Supplementary table 1”. Please correct.

Line 175 and beyond. Please consider using the term “necropsy” instead of “autopsy”.

Line 197. I believe the panels from figure 2 are cited wrongly: panel 2I shows H&E staining (infiltrating inflammatory cells) and panel 2J shows immunohistochemistry (viral antigen).

Line 334. “…as well as in infectious conditions” I believe it should say “…as well as in infection conditions”.

Line 404. This is probably a typo, “deceased” should say “decreased”

Line 423. “…reported those CD4+ T cells to have cytolytic…” please double check. Did you mean CD8+?

Reviewer #3: The manuscript by Schwaiger and colleagues described experimental H1N1pdm09 infection in pigs in order to mimic human seasonal influenza infections. The authors used 2009 pandemic H1N1 virus to intranasally infect a group of 4-week-old pigs for 21 days, then re-infected them with the same virus again to investigate systemic and local immune responses by testing blood, mucosa of nasal cavity and lung tissues bronchoalveolar lavage fluid samples. Result showed that decreasing numbers of peripheral blood lymphocytes were detected after the first infection, the simultaneous increase in the frequencies of proliferating cells correlated with an increase in infiltrating leukocytes in the lung. Furthermore, a cytotoxic T cell response was detected but restricted to the respiratory route of virus entry such as the nose, the lung and the bronchoalveolar lavage according to detected enhanced perforin expression in αβ and γδ T cells in the respiratory tract. Increasing frequencies of CD8αα expressing αβ T cells were also observed rapidly after the first infection, which could play a role in inhibition of uncontrolled inflammation in the respiratory tract. Authors conclude that the results from this study demonstrate that experimental influenza A virus infection in pigs mimics major characteristics of human seasonal influenza A virus infections. The manuscript is well written and provides interesting information which complements findings of former studies.

Comments:

The pigs used in the study were purchased from a commercial high health status herd. The detailed information on these pigs should be provided such as which pathogens (influenza A, D virus, PRRSV, PCV2 and mycoplasma etc) were tested prior to infection as any prior infection will impact the obtained results and further interpret.

In the figures 5, 6, 9 and 10, the data from only infected pigs were presented despite of the data at d0 as controls. If the pigs were infected other pathogens before, these data could not reflect the reality of only infection of 2009 pandemic H1N1 virus. Therefore, it is necessary to provide more detailed information on pigs

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9 Sep 2019

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2. We note that you have included the phrase “data not shown” in your manuscript. Unfortunately, this does not meet our data sharing requirements. PLOS does not permit references to inaccessible data. We require that authors provide all relevant data within the paper, Supporting Information files, or in an acceptable, public repository. Please add a citation to support this phrase or upload the data that corresponds with these findings to a stable repository (such as Figshare or Dryad) and provide and URLs, DOIs, or accession numbers that may be used to access these data. Or, if the data are not a core part of the research being presented in your study, we ask that you remove the phrase that refers to these data.

We thank the editor for this note. Because the data are not a core part of our research, we removed the phrase “data not shown” from line 195.

Reviewers' comments:

Reviewer #1:

1. Please furnish information on the pre-screen done on the pigs. This should include both qPCR and serology (ELISA) for IAV, PRRSV, PPV, PCV 2, and TTV. If animals were negative prior to d0, what was their status at the end of the study?

Prior to d0, pigs were tested negative for IAV genome by qPCR. During the trial, we were able to detect IAV only on day 2 and 4 after first infection in nasal swabs (see Table 1). Neutralizing antibodies were detected in sera of infected pigs only from day 14 with very low titres until the end of the study (Table 2). Control pigs did not show antibodies against H1N1.

Table 1 Viral titres during trial in nasal swabs. Nasal swabs were taken throughout the study period and tested for replicable virus by TCID50 assay. Results are given as TCID50/ml supernatant. First three rows are mock-infected control animals (grey filling). neg = negative, nt = not tested.

Animal d0 d2 d4 d7 d22 d23 d25 Necropsy at day

7936 neg neg neg neg neg neg neg 30

7444 neg neg neg neg neg neg neg

7460 neg neg neg neg neg neg neg

7477 neg neg 1,5x 102 nt nt nt nt 4

7470 neg neg 3,2x 103 nt nt nt nt

7964 neg neg 3,2x 103 nt nt nt nt

7993 neg 6,8x 103 1,5x 104 nt nt nt nt

7445 neg 3,2x 103 6,8x 104 nt nt nt nt

7370 neg 3,2x 102 6,8x 103 neg nt nt nt 7

7410 neg 3,2x 102 6,7x 101 neg nt nt nt

7468 neg 6,8x 102 1,4x 103 neg nt nt nt

7479 neg 1,4x 103 3,2x 103 neg nt nt nt

7965 neg 1,4x 102 1,5x 102 neg nt nt nt

7403 neg 1,4x 103 7,1x 102 neg nt nt nt 21

7970 neg neg 6,8x 103 neg nt nt nt

7478 neg 1,4x 104 1,5x 105 neg nt nt nt

7939 neg neg 6,7x 101 neg nt nt nt

7434 neg neg 1,5x 104 neg nt nt nt

7443 neg 6,8x 102 6,8x 102 neg neg neg neg 25

7367 neg 1,4x 102 6,8x 102 neg neg neg neg

7402 neg 1,5x 104 1,5x 104 neg neg neg neg

7480 neg 1,4x 102 3,2x 104 neg neg neg neg

7471 neg 1,4x 102 6,8x 104 neg neg neg neg

7407 neg 6,8x 102 1,5x 104 neg neg neg neg 31

7408 neg neg 1,5x 103 neg neg neg neg

7995 neg neg 1,5x 103 neg neg neg neg

7991 neg neg 3,2x 103 neg neg neg neg

7441 neg neg neg neg neg neg neg

7433 neg neg 1,5x 103 neg neg neg neg

Table 2. Titers of neutralizing antibodies. During trial serum of naïve (rows 1-3, gray filling) and infected (rows 4-9) was tested by Hemagglutination inhibition test. neg = negative, nt = not tested.

Animal d7 d14 d21 d22 d25 d31

7936 nt nt nt nt nt neg

7444 nt nt nt nt nt neg

7460 nt nt nt nt nt neg

7408 neg 1:160 1:160 1:320 1:320 1:320

7407 neg 1:160 1:160 1:160 1:320 1:160

7991 neg 1:160 neg neg 1:160 neg

7995 neg 1:160 neg neg neg neg

7441 neg 1:320 1:160 1:160 1:160 1:160

7433 neg neg 1:160 1:160 1:320 1:320

We have not yet provided this information, as we do not consider it a core part of this study, which focuses on the cellular mucosal immune response. If desired, we can include viral titres and titration of neutralizing antibodies in new supplementary tables to provide additional information.

The farm from which the pigs were purchased is free of PRRSV, which obviates the need for re-testing by our laboratory. Furthermore, the sows were vaccinated against PPV on the 170th and 190th day of life and against PCV 2 on day 160. Because we started our study when the animals were 8 weeks old, the maternal antibodies against these viruses were probably degraded as far as possible [1]. Piglets themselves received vaccines against PCV2 and H. parasuis at their 14th day of life, which makes it likely that they already started to generate antibodies against these viruses, which we did not screen for. For the occurrence of TTV was tested neither by us nor by the farm. We included a section on pathogen status and vaccination program of the farm in Material and Methods section (lines 81-89): “This farm is free from the following diseases or pathogens: Pseudorabies, classical swine fever virus (CSFV), Porcine Reproductive and Respiratory Syndrome Virus (PRRSV), Actinobacillus pleuropneumoniae, Mycoplasma hypopneumoniae, ascaris, mange, Brachyspira hyodysenteriae and salmonella category I. Vaccination program does not include vaccination against influenza viruses but the following vaccines were administered: Sows were vaccinated against porcine circovirus type 2 (once), Erysipelothrix rhusiopathiae/Porcine parvovirus (twice), salmonella (twice) and Haemophilus parasuis (twice). Piglets were vaccinated against PCV2 once and Mycoplasma hypopneumoniae twice.”

2. How were the mock and challenge groups housed?

After arrival of the animals (4-weeks of age), the piglets were randomly separated in three groups and housed in three different sheds under BSL2 conditions. One week prior to infection (7-weeks of age) pigs were divided in six groups, with the control group held in a shed at the other end of the building. Five groups were infected one week later. Contact between the animals of different sheds was strictly avoided not only structurally but also by different hygiene interventions (showers between the sheds, disposable clothing and masks within the sheds).

3. Could comparative counts and FACS data for mocks also be clearly identified and shown along with challenge groups?

With this question, the reviewer addresses one of the central points of our study. The characteristic feature of this study is that the animals intended for organ removal (four infected groups plus control group) were only subjected to blood sample drawing on the day of necropsy. This also includes the control group whose section day coincided with the last day of the study. We deliberately chose this scheme because regular blood collection has a significant effect on the stress level of the animals and thus on the immune response. In order to avoid this effect, the same six pigs randomly selected at the beginning of the experiment were used for the blood kinetics only. Blood from control animals was only drawn at day of their euthanasia. This is why we cannot provide comparative counts and FACS data for peripheral blood leukocytes of mock animals from time points correlating with those of infected blood-donating group. From our point of view, the d0 sample of the latter animals represents the adequate control to all blood samples of these animals after infection.

Reviewer #2:

The manuscript by Schwaiger and colleagues describes the characterization of swine immune cell population changes following an experimental infection with a human H1N1 influenza virus. The results are of interest because swine are a natural host of influenza viruses and can also play a role in the zoonotic transmission of non-human influenza strains into the human population. In addition, (as stated by the authors), because of comparable sizes, immunity and physiology, pigs are a valuable animal model for evaluating immune responses, vaccines, antivirals, etc. In general, the manuscript is well written and provides detailed methods and results that can serve as a guide to other groups.

1. To be fair, the authors should also point out some important differences between pigs and humans such as the much shorter “childhood” and general lifespan of pigs; important differences in immune cell populations such as the double positive CD4+CD8+ lymphocytes and strikingly different proportions of gamma-delta+ T cells in blood, or the lack of reagents and general knowledge of the pig’s immunity in comparison to other animal models (mice).

The reviewer is right. We have added information about drawbacks when using the pig as a model compared to the mouse model (lines 33-37 and lines 44-45): “Studies on the immune response in pigs have long been biased by the lack of specific reagents. Especially the variety of antibodies available for the pig is still far lower to that of mice. As a result, the knowledge of the porcine immune response in general is much smaller compared to that of mice. Despite these disadvantages, pigs have decisive advantages as a model for human IAV infection.”, and “[…] and by expanding the number of reagents for porcine immunological analyses, […]”. We also included a paragraph on the major differences in peripheral blood cell composition between human and swine (lines 57-63 and references 26-32): “However, it is important to note that the prominent porcine populations of CD4+/CD8+ double positive T cells [26] as well as the high number of peripheral γδ T [27] cells are virtually absent in humans [28, 29], representing a major difference. Besides the numerical difference, the functionality of the two cell populations is comparable in both human and swine. CD4+/CD8+ double positive T cells are mature effector cells with memory characteristics that rapidly mount antigen-specific responses upon antigenic challenge [18, 30]. Besides acting as innate immune cells via pattern recognition receptors and direct killing of infected cells, γδ T cells do play a major role in antigen processing and presentation in human and swine [31, 32].”

With regard to the shorter "childhood" of pigs, we cannot agree: compared to the life span of pigs, the "childhood" is not much shorter. Especially regarding the risk and the frequency of infections of the young animals in the flat deck area, the herd veterinarians report amazing parallels to children who come to kindergarten.

2. The authors should also consider the following specific comments:

a. Line 68. The animals were obtained from a commercial herd. If known, it would be important to state the serological status (from natural infection and from vaccination) of the mothers, since the presence of maternal antibodies against H1N1 influenza would have a strong impact in the evolution of the infection.

The reviewer is right, that maternal antibodies have a strong impact in the evolution of infection. In our study, serological status of sows regarding antibodies directed against H1N1 viruses was not taken into account, because our study started, when pigs were 8 weeks of age. By this time, all maternal antibodies should have been largely metabolized [1].

b. Line 73. Animals were re-infected with the same influenza strain 21 days after the first infection. In the context of modelling influenza in humans, it is not clear what was the purpose of this infection. In humans, re-infection occurs in the following influenza seasons (at least 1 year later) and by an antigenically drifted strain.

The reviewer is right that re-infection in humans occurs primarily around a year later with an antigenically drifted strain in the vast majority of cases. The purpose of the re-infection in our study three weeks after first infection was to analyse a potential memory response. This re-infection was not intended to imitate the seasonal flu that occurs in humans a year later. The period of three weeks between the first and second infection in our study allows at most a comparison with the seasonal infection after vaccination in humans.

c. Line 122. “…antibody against the anti-matrixprotein…” this sentence is incorrect. It should be either “…antibody against the matrixprotein…” or “…anti-matrixprotein monoclonal antibody…”. Please correct.

We corrected the sentence as follows: “…antibody against the matrixprotein of influenza A virus…” (line 139).

d. Line 128. It might be useful to describe what TBS stands for.

We added the full form “Tris-buffered saline (TBS)” in the material and methods section (line 145).

e. Line 149. In this paragraph there is a sentence repeated “Antibodies used in this study are listed in Supplementary table 1”. Please correct.

We have deleted one of the duplicate sentences (line 169).

f. Line 175 and beyond. Please consider using the term “necropsy” instead of “autopsy”.

We exchanged the term “autopsy” to “necropsy” throughout the document (lines 190, 221, 229, 325, 365, 392 and 520).

g. Line 197. I believe the panels from figure 2 are cited wrongly: panel 2I shows H&E staining (infiltrating inflammatory cells) and panel 2J shows immunohistochemistry (viral antigen).

We fixed the wrong citation and used the correct labelling to the panels of figure 2 in the text. The new text is as follows: “Still negative for viral antigen (Fig 2J, right panel), the amount of infiltrating inflammatory cells slightly decreased at 25 dpi (Fig 2I, right panel). “ (lines 212-213).

h. Line 334. “…as well as in infectious conditions” I believe it should say “…as well as in infection conditions”.

We corrected the expression “…infectious conditions”. It now reads as follows: “…infection conditions” (line 349).

i. Line 404. This is probably a typo, “deceased” should say “decreased”

We have corrected the typo. The word shall read as follows: “decreased” (line 418).

j. Line 423. “…reported those CD4+ T cells to have cytolytic…” please double check. Did you mean CD8+?

Although it may sound unusual at first, Wilkinson and colleagues showed that influenza-specific CD4+ T cells were able to kill autologous cells in a peptide specific manner [2]. This finding was later confirmed in a study performed by Zhou et al. [3]. Growing amount of evidence suggest CD4+ T cells to have direct antiviral activity and thus, to play a critical role in numerous viral infections in humans and animal models in vivo. These findings are summarized in e.g. these reviews: [4-6].

Reviewer #3:

The manuscript by Schwaiger and colleagues described experimental H1N1pdm09 infection in pigs in order to mimic human seasonal influenza infections. The authors used 2009 pandemic H1N1 virus to intranasally infect a group of 4-week-old pigs for 21 days, then re-infected them with the same virus again to investigate systemic and local immune responses by testing blood, mucosa of nasal cavity and lung tissues bronchoalveolar lavage fluid samples. Result showed that decreasing numbers of peripheral blood lymphocytes were detected after the first infection, the simultaneous increase in the frequencies of proliferating cells correlated with an increase in infiltrating leukocytes in the lung. Furthermore, a cytotoxic T cell response was detected but restricted to the respiratory route of virus entry such as the nose, the lung and the bronchoalveolar lavage according to detected enhanced perforin expression in αβ and γδ T cells in the respiratory tract. Increasing frequencies of CD8αα expressing αβ T cells were also observed rapidly after the first infection, which could play a role in inhibition of uncontrolled inflammation in the respiratory tract. Authors conclude that the results from this study demonstrate that experimental influenza A virus infection in pigs mimics major characteristics of human seasonal influenza A virus infections. The manuscript is well written and provides interesting information, which complements findings of former studies.

1. The pigs used in the study were purchased from a commercial high health status herd. The detailed information on these pigs should be provided such as which pathogens (influenza A, D virus, PRRSV, PCV2 and mycoplasma etc) were tested prior to infection as any prior infection will impact the obtained results and further interpret.

Piglets were tested negative by qPCR for influenza A virus genome. Neutralizing antibodies were detected in sera of infected pigs only from day 14 post infection until the end of the study. We have not yet provided this information, as we do not consider it a core part of this study, which focuses on the cellular mucosal immune response. If desired, we can include viral titres and titration of neutralizing antibodies in new supplementary tables to provide additional information.

The farm from which the pigs were purchased is free from PRRSV, Pseudorabies, classical swine fever virus (CSFV), Porcine Reproductive and Respiratory Syndrome Virus (PRRSV), Actinobacillus pleuropneumoniae, Mycoplasma hypopneumoniae, ascaris, mange, Brachyspira hyodysenteriae and salmonella category I. which obviates the need for re-testing by our laboratory. Furthermore, the sows were vaccinated against PPV on the 170th and 190th day of life and against PCV 2 on day 160. Additionally, piglets themselves received vaccines against PCV2 and H. parasuis at their 14th day of life. This vaccination program together with the strict hygiene management (no contact to other animals, showers between the sheds, disposable clothing and masks within the sheds) makes a prior infection rather unlikely.

To address this issue, we included a section on pathogen status and vaccination program of the farm in Material and Methods section (lines 81-89): “This farm is free from the following diseases or pathogens: pseudorabies, classical swine fever virus (CSFV), Porcine Reproductive and Respiratory Syndrome Virus (PRRSV), Actinobacillus pleuropneumoniae, Mycoplasma hypopneumoniae, ascaris, mange, Brachyspira hyodysenteriae and salmonella category I. Vaccination program does not include vaccination against influenza viruses but the following vaccines were administered: Sows were vaccinated against porcine circovirus type 2 (once), Erysipelothrix rhusiopathiae/Porcine parvovirus (twice), salmonella (twice) and Haemophilus parasuis (twice). Piglets were vaccinated against PCV2 once and Mycoplasma hypopneumoniae twice.”

2. In the figures 5, 6, 9 and 10, the data from only infected pigs were presented despite of the data at d0 as controls. If the pigs were infected other pathogens before, these data could not reflect the reality of only infection of 2009 pandemic H1N1 virus. Therefore, it is necessary to provide more detailed information on pigs.

Yes, the reviewer is right, in Fig. 8, 9 and 10 we compared animals at different times of infection with only one control group at one time point. We have opted for this method to avoid excessive animal numbers. It is true that the control animals could also develop infections during the experimental period, so the control group was subjected to necropsy at the end of the experiment and not at the beginning. The animals all come from the same delivery, have the same age and were kept under comparable conditions (see reviewer 1, comment #2), so the error is considered to be minor. We have included information about health status of the pigs and the vaccination regime of the breeding farm in the manuscript. Thank you very much for this note.

The choice of correct controls is always difficult and controversial in these experiments: is the individual animal control (d0 before infection) superior to an untreated control group or vice versa? Animal-individual control has the error that an incalculable handling error (capture, frequent bleeding) is to be considered. In order best to present both situations equally well, we conducted two independent experiments in parallel with animals of the same origin and age: animal-individual non-lethal data from blood (Fig 5-7) and data for the organs after necropsy with an uninfected comparison group (Fig 8-10).

As stated above (reviewer 3, comment #1), pigs were tested negative for IAV genome by qPCR prior to infection. Additionally the vaccine program of the farm from which we purchased the piglets almost eliminates a prior infection with the mentioned porcine pathogens. Because contact to other animals was strictly avoided and hygiene management (also see reviewer 1, comment #2) was rigorously followed, it is rather unlikely that pigs were infected with other pathogens in the time of acclimatization in our animal facility under BSL2 conditions (four weeks prior to start of experiment). In addition, necropsies did only reveal a single lung lesion in one animal (in blood donating group) that could not be attributed to IAV infection. As already stated above, we included a section in Material and Methods section on pathogen status and vaccination program of the farm (lines 81-89).

References:

1. Reeth KV, Labarque G, Pensaert M. Serological Profiles after Consecutive Experimental Infections of Pigs with European H1N1, H3N2, and H1N2 Swine Influenza Viruses. Viral Immunology. 2006;19(3):373-82. doi: 10.1089/vim.2006.19.373.

2. Wilkinson TM, Li CKF, Chui CSC, Huang AKY, Perkins M, Liebner JC, et al. Preexisting influenza-specific CD4+ T cells correlate with disease protection against influenza challenge in humans. Nature Medicine. 2012;18:274. doi: 10.1038/nm.2612

https://www.nature.com/articles/nm.2612#supplementary-information.

3. Zhou X, McElhaney JE. Age-related changes in memory and effector T cells responding to influenza A/H3N2 and pandemic A/H1N1 strains in humans. Vaccine. 2011;29(11):2169-77. doi: 10.1016/j.vaccine.2010.12.029. PubMed PMID: 21353149.

4. Juno JA, van Bockel D, Kent SJ, Kelleher AD, Zaunders JJ, Munier CM. Cytotoxic CD4 T Cells-Friend or Foe during Viral Infection? Front Immunol. 2017;8:19. Epub 2017/02/09. doi: 10.3389/fimmu.2017.00019. PubMed PMID: 28167943; PubMed Central PMCID: PMCPMC5253382.

5. Soghoian DZ, Streeck H. Cytolytic CD4(+) T cells in viral immunity. Expert Rev Vaccines. 2010;9(12):1453-63. doi: 10.1586/erv.10.132. PubMed PMID: 21105780.

6. Brown DM, Lampe AT, Workman AM. The Differentiation and Protective Function of Cytolytic CD4 T Cells in Influenza Infection. Frontiers in immunology. 2016;7:93-. doi: 10.3389/fimmu.2016.00093. PubMed PMID: 27014272.

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11 Sep 2019

Experimental H1N1pdm09 infection in pigs mimics human seasonal influenza infections

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13 Sep 2019

PONE-D-19-15088R1

Experimental H1N1pdm09 infection in pigs mimics human seasonal influenza infections

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