Characterization of the Cross-Species Transmission Potential for Porcine Deltacoronaviruses Expressing Sparrow Coronavirus Spike Protein in Commercial Poultry

Avian species often serve as transmission vectors and sources of recombination for viral infections due to their ability to travel vast distances and their gregarious behaviors. Recently a novel deltacoronavirus (DCoV) was identified in sparrows. Sparrow deltacoronavirus (SpDCoV), coupled with close contact between sparrows and swine carrying porcine deltacoronavirus (PDCoV) may facilitate recombination of DCoVs resulting in novel CoV variants. We hypothesized that the spike (S) protein or receptor-binding domain (RBD) from sparrow coronaviruses (SpCoVs) may enhance infection in poultry. We used recombinant chimeric viruses, which express S protein or the RBD of SpCoV (icPDCoV-SHKU17, and icPDCoV-RBDISU) on the genomic backbone of an infectious clone of PDCoV (icPDCoV). Chimeric viruses were utilized to infect chicken derived DF-1 cells, turkey poults, and embryonated chicken eggs (ECEs) to examine permissiveness, viral replication kinetics, pathogenesis and pathology. We demonstrated that DF-1 cells in addition to the positive control LLC-PK1 cells are susceptible to SpCoV spike- and RBD- recombinant chimeric virus infections. However, the replication of chimeric viruses in DF-1 cells, but not LLC-PK1 cells, was inefficient. Inoculated 8-day-old turkey poults appeared resistant to icPDCoV-, icPDCoV-SHKU17- and icPDCoV-RBDISU virus infections. In 5-day-old ECEs, significant mortality was observed in PDCoV inoculated eggs with less in the spike chimeras, while in 11-day-old ECEs there was no evidence of viral replication, suggesting that PDCoV is better adapted to cross species infection and differentiated ECE cells are not susceptible to PDCoV infection. Collectively, we demonstrate that the SpCoV chimeric viruses are not more infectious in turkeys, nor ECEs than wild type PDCoV. Therefore, understanding the cell and host factors that contribute to resistance to PDCoV and avian-swine chimeric virus infections may aid in the design of novel antiviral therapies against DCoVs.


Introduction
Coronaviruses (CoVs) are enveloped viruses possessing the largest positive sense, single-stranded RNA genomes [1]. CoVs belong to the family Coronaviridae and the order Nidovirales [2]. According to the phylogenetic relationships and genomic structures, there are four genera of the Orthocoronavirinae subfamily: Alphacoronavirus, Betacoronavirus, Gammacoronavirus and Deltacoronavirus. Viruses from each coronavirus genus have been found in diverse host species, but only DCoVs in multiple mammalian and avian species [1]. chickens [19]. Boley et al. demonstrated that PDCoV can infect 14-day-old chicks and 14-day-old turkey poults [25]. Jung et al. experimentally infected gnoto-biotic calves with PDCoV [30]. More recently it was shown that replacement of the S protein of PDCoV with the spike protein from SpDCoV resulted in asymptomatic infection of pigs with altered tissue tropism from the gastrointestinal tract to the respiratory tract [31]. The consequence of recombination of PDCoV with SpDCoV S in infection of poultry has not been reported. Therefore, this study was conducted to investigate the effects of SpDCoV S protein or RBD replacement in an infectious clone of PDCoV on viral susceptibility and replication in various cell lines, on pathogenesis and pathology in specific pathogen free (SPF) turkey poults, and on the infection of embryonated chicken eggs.

Viruses
icPDCoV, icPDCoV-S HKU17 and icPDCoV-RBD ISU were produced and characterized by Dr Q Wang and colleagues as described previously [31]. These viruses were validated for infectivity in gnotobiotic pigs as described [31].

Replication Kinetics in Various Cell Lines
The replication of icPDCoV, icPDCoV-S HKU17 and icPDCoV-RBD ISU in DF-1 and LLC-PK1 cell lines was compared using a multiplicity of infection (MOI) of up to 10 and 0.01, respectively. Viruses were incubated with the indicated cells in triplicate wells of a 96-well plate or 24 well plate at 37 • C and 5% CO 2 for 1 h, as previously described [31]. After removal of virus inocula, cells were washed twice with PBS. Cells were covered with serum-free media with 10 µg/mL trypsin (LLC-PK-1) or no trypsin (most DF-1 assays) (Gibco) and incubated at 37 • C and 5% CO 2 . Supernatants were harvested at 0, 6, 12, 24, 48, and 72 h and stored at −80 • C until use or entire wells were scraped at 6,12,24,48, and 72 hpi. The replication kinetics were performed in triplicate. CCIF or TCID 50 was used to titrate the viruses as described previously [32]. The results are presented as mean and standard deviation of fluorescent foci-forming units or TCID 50 per mL.

Cell-Associated and Cell-Free Virus RNA Loads in DF-1 Cell Line
The cell-associated and cell-free viral loads were assessed as genome equivalent of viral RNA in inoculated DF-1 cells collected during the experiment. DF-1 cells were cultured in growth medium as described earlier in a 96 well plate at 37 • C and 5% CO 2 . Cells that had attained 90% confluence were infected with icPDCoV-, icPDCoV-S HKU17 -, and icPDCoV-RBD ISU viruses at an MOI of 1. Viruses were incubated with the indicated cells in duplicate wells at 37 • C and 5% CO 2 for 1 h, after which inocula were removed, and cells were washed twice with PBS. Cells were then covered with serum-free media with 10 µg/mL trypsin (Gibco) and incubated at 37 • C and 5% CO 2 . Cell-associated and cell-free virus were harvested at 0-, 6-, 12-, and 24-h post infection (hpi) and stored at −80 • C until assayed ( Figure 1). Before extraction of the cell-associated fraction, the cells in the monolayer were washed twice with PBS then dissolved in 200 µL of lysis buffer containing RXV and RNA carrier from RNA extraction kit and scraped from the plate. Cell-associated and cell-free viral loads were analyzed by quantification of viral RNA using a TaqMan-probe-based RT-qPCR with PDCoV-M gene-specific primers. The percentage of release was calculated using the following ratio: well plate or 24 well plate at 37 °C and 5% CO2 for 1 h, as previously described [31]. After removal of virus inocula, cells were washed twice with PBS. Cells were covered with serum-free media with 10 µg/mL trypsin (LLC-PK-1) or no trypsin (most DF-1 assays) (Gibco) and incubated at 37 °C and 5% CO2. Supernatants were harvested at 0, 6, 12, 24, 48, and 72 h and stored at −80 °C until use or entire wells were scraped at 6, 12, 24, 48, and 72 hpi. The replication kinetics were performed in triplicate. CCIF or TCID50 was used to titrate the viruses as described previously [32]. The results are presented as mean and standard deviation of fluorescent foci-forming units or TCID50 per ml.

Cell-Associated and Cell-Free Virus RNA Loads in DF-1 Cell Line
The cell-associated and cell-free viral loads were assessed as genome equivalent of viral RNA in inoculated DF-1 cells collected during the experiment. DF-1 cells were cultured in growth medium as described earlier in a 96 well plate at 37 °C and 5% CO2. Cells that had attained 90% confluence were infected with icPDCoV-, icPDCoV-SHKU17-, and icPDCoV-RBDISU viruses at an MOI of 1. Viruses were incubated with the indicated cells in duplicate wells at 37 °C and 5% CO2 for 1 h, after which inocula were removed, and cells were washed twice with PBS. Cells were then covered with serum-free media with 10 µg/mL trypsin (Gibco) and incubated at 37 °C and 5% CO2. Cell-associated and cellfree virus were harvested at 0-, 6-, 12-, and 24-h post infection (hpi) and stored at −80 °C until assayed ( Figure 1). Before extraction of the cell-associated fraction, the cells in the monolayer were washed twice with PBS then dissolved in 200 µL of lysis buffer containing RXV and RNA carrier from RNA extraction kit and scraped from the plate. Cell-associated and cell-free viral loads were analyzed by quantification of viral RNA using a Taq-Man-probe-based RT-qPCR with PDCoV-M gene-specific primers. The percentage of release was calculated using the following ratio: Release ratio = cell-free (cell-associated + cell-free) × 100

Experimental Inoculation Turkey Poults
In vivo experimentation was undertaken within the guidelines of the Ohio State University. Protocols and chicken sampling procedures were approved by the Institutional Animal Care and Use Committee (IACUC).
The pathogenesis and pathology of the infectious clone PDCoV and recombinant viruses were evaluated in SPF turkey poults (Meleagris gallopavo). A total of 56 seven-day-

Experimental Inoculation Turkey Poults
In vivo experimentation was undertaken within the guidelines of the Ohio State University. Protocols and chicken sampling procedures were approved by the Institutional Animal Care and Use Committee (IACUC).
The pathogenesis and pathology of the infectious clone PDCoV and recombinant viruses were evaluated in SPF turkey poults (Meleagris gallopavo). A total of 56 seven-dayold SPF turkey poults were obtained from the Ohio Agricultural Research and Development Center of The Ohio State University (Wooster, Ohio, USA) flock (OARDC flock), which has no history of exposure to swine or symptoms related to PDCoV, TGEV, or PEDV. Poults were floor housed in temperature-controlled biosafety level 2 (BSL-2) rooms for the period of the experiment with sustained artificial light, floor covered with wood litter shavings and birds were continuously provided access to food and water. Birds appeared healthy and showed no diarrhea or other clinical signs during the acclimation period. Turkey poults were randomly assigned to one of the four infection groups, which were housed in separate rooms: (1) icPDCoV-infected (n = 10); (2) icPDCoV-S HKU17 -infected (n = 10); icPDCoV-RBD ISU -infected (n = 10); or MEM-infected (control) (n = 10). Birds were inoculated through the choanal cleft with 4.4 × 10 5 FFU/poult of virus in a volume of 195 µL. Ten control turkey poults were also inoculated with the same amount of MEM. Four uninfected birds per group, at 1-day post inoculation (DPI), were randomly assigned to serve as sentinels and allowed to commingle with each infected or control group ( Figure 2).  ), which has no history of exposure to swine or symptoms related to PDCoV, TGEV, or PEDV. Poults were floor housed in temperature-controlled biosafety level 2 (BSL-2) rooms for the period of the experiment with sustained artificial light, floor covered with wood litter shavings and birds were continuously provided access to food and water. Birds appeared healthy and showed no diarrhea or other clinical signs during the acclimation period. Turkey poults were randomly assigned to one of the four infection groups, which were housed in separate rooms: (1) icPDCoV-infected (n = 10); (2) icPDCoV-SHKU17-infected (n = 10); icPDCoV-RBDISU-infected (n = 10); or MEM-infected (control) (n = 10). Birds were inoculated through the choanal cleft with 4.4 × 10 5 FFU/poult of virus in a volume of 195 µL. Ten control turkey poults were also inoculated with the same amount of MEM. Four uninfected birds per group, at 1-day post inoculation (DPI), were randomly assigned to serve as sentinels and allowed to commingle with each infected or control group ( Figure  2).

Clinical Swabs and Tissue Collection
To evaluate viral shedding, tracheal and cloacal swabs were collected from challenged and control poults daily. Fecal consistency produced during swabbing and via room observation was monitored and scored as follows: 0, solid (No diarrhea); 1, pasty (Likely normal); 2, semiliquid (Some potential diarrhea); and 3, liquid (Diarrhea) as previously published [25]. Fecal scores >2 were considered as diarrhea. Tracheal and fecal samples were diluted in 1 mL MEM and centrifuged at 1832× g at 4 °C for 20 min, and the supernatants were collected for viral RNA isolation. Five poults from each group were euthanized and their duodenum, jejunum, ileum, and lung were collected at 3-and 5-DPI and fixed with 4% paraformaldehyde during the bird necropsy. All sentinel birds were euthanized at 3-day post commingling. Paraformaldehyde-fixed tissues were preserved for immunohistochemical examination to detect lesions.

Inoculation of Chicken Embryos with the Chimeric Viruses
A total number of 29 11-day-old SPF ECEs and 55 5-day-old SPF ECEs from the OARDC flock were divided into 4 or 3 groups, respectively. Viruses icPDCoV, icPDCoV-SHKU17, icPDCoV-RBDISU were injected into the 29 11-day-old SPF ECEs using a titer of 6 log10 fluorescent focus units (FFU)/egg, in 200 µL. Five ECEs for each virus were

Clinical Swabs and Tissue Collection
To evaluate viral shedding, tracheal and cloacal swabs were collected from challenged and control poults daily. Fecal consistency produced during swabbing and via room observation was monitored and scored as follows: 0, solid (No diarrhea); 1, pasty (Likely normal); 2, semiliquid (Some potential diarrhea); and 3, liquid (Diarrhea) as previously published [25]. Fecal scores >2 were considered as diarrhea. Tracheal and fecal samples were diluted in 1 mL MEM and centrifuged at 1832× g at 4 • C for 20 min, and the supernatants were collected for viral RNA isolation. Five poults from each group were euthanized and their duodenum, jejunum, ileum, and lung were collected at 3-and 5-DPI and fixed with 4% paraformaldehyde during the bird necropsy. All sentinel birds were euthanized at 3-day post commingling. Paraformaldehyde-fixed tissues were preserved for immunohistochemical examination to detect lesions.

Inoculation of Chicken Embryos with the Chimeric Viruses
A total number of 29 11-day-old SPF ECEs and 55 5-day-old SPF ECEs from the OARDC flock were divided into 4 or 3 groups, respectively. Viruses icPDCoV, icPDCoV-S HKU17 , icPDCoV-RBD ISU were injected into the 29 11-day-old SPF ECEs using a titer of 6 log 10 fluorescent focus units (FFU)/egg, in 200 µL. Five ECEs for each virus were inoculated with viable icPDCoV, icPDCoV-S HKU17 , or icPDCoV-RBD ISU . As a background control, three ECEs for each virus were similarly inoculated with killed (heat inactivated at 60 • C for 20 min) viruses. The ECEs of the mock group were inoculated with 200 µL MEM. For the 5-day-old SPF ECEs, icPDCoV, icPDCoV-SHKU17, or icPDCoV-RBDISU were serially diluted from neat to up to 10 −6 in MEM and used to inoculate 5 eggs per infectious dose, 100 µL/egg. In brief, the embryos were inoculated via the allantoic cavity route and incubated at 37.5 • C, 55 ± 2% relative humidity. The viability of embryos was examined by using an egg candler. An ECE was harvested by putting at 4 • C for 2-6 h when it was found dead. The remaining 11-day-old and 5-day-old SPF ECEs were harvested at 3 DPI and 5 DPI, respectively. The eggshells above the air sac were disinfected with 70% ethanol, and the allantoic fluids were collected and centrifuged at 4122× g at 4 • C for 20 min to remove the cell debris. For the trial using 11-day-old ECEs, after removing heads, wings, and legs, embryos were dissected into two portions, and the whole thoracic and abdominal tissues in each cavity were harvested separately and homogenized using an electric homogenizer (OMNI GLH International/USA) for 1 min in equal amounts of MEM (1 g of tissue:1 mL of MEM). The allantoic fluids and tissues were analyzed by quantification of viral RNA using a TaqMan RT-qPCR with PDCoV-M gene-specific primers as reported previously [34]. The allantoic fluid samples of the 5-day-old ECEs (at no dilution and 1:10 dilution) were tested for infectious virus in LLC-PK1 cells using 96-well plates.

RNA Isolation and RT-qPCR Analysis
Total viral RNA was extracted from 150 µL of swab supernatants using TRIzol reagent (Life Technologies, Carlsbad CA, USA) according to the manufacturer's instructions and eluted in 40 µL RNase free water. We determined the titers of viral RNA shed in swabs by carrying out one step TaqMan-probe-based RT-qPCR as described previously [35] targeting the PDCoV-specific M gene with the primers (forward 5 -ATC GAC CAC ATG GCT CCA A-3 and reverse 5 -CAG CTC TTG CCC ATG TAG CTT-3 ), which were designed based on the sequence of PDCoV strain US, Illinois121/2014 (GenBank accession no. KJ481931) and the probe (5 -/56-FAM/CAC ACC AGT/ZEN/CGT TAA GCA TGG CAA GCT/3IABkFQ/3 ). The reaction system was set up with 12.5 µL nuclease free water, 5 µL of 4X TaqMan (Applied Biosystems, Waltham MA, USA), 0.5 µL of F primer, R primer, and probe, and 1 µL of template RNA. The amplified fragment was 541-bp with the following thermal cycling profile: 50 • C for 5 min and 95 • C for 20 s, followed by 45 cycles of 95 • C for 3 s and 55 • C for 30 s. The detection limit of the RT-qPCR was 1.47 × 10 5 genomic equivalents (GEs)/mL, which corresponded to 5.17 log 10 GE/mL of PDCoV.

Immunohistochemistry (IHC) and Immunofluorescence (IF) Staining of Tissues from Turkey Poults
At necropsy, tissues including small intestines, duodenum to ileum, and lung were collected from euthanized poults and then fixed in 4% paraformaldehyde for 24 h at room temperature. After trimming, processing, embedding, and sectioning (4.5 µm), tissues were first incubated at 60 • C for 30 min and deparaffinized using xylenes followed by rehydrating through graded alcohol and deionized (DI) water [36,37].
For IHC analysis, dewaxed sections were treated with pronase reagent (GeneTex), for 18 min at RT for antigen retrieval followed by peroxide and power block (BioGenex, Fremont, CA, USA) to quench endogenous peroxide and non-specific binding for 10 and 30 min at RT, respectively. Sections were then incubated with a primary antibody as above at 4 • C overnight, followed by a non-biotin polymerized horseradish peroxidase system (BioGenex Laboratories, Fremont, CA, USA) as described previously [31]. PBS or deionized water was used for rinsing cells between incubations. Slides were imaged using an Olympus IX-70 microscope (Olympus, Tokyo, Japan).
For IF staining, DF-1 and LLC-PK1 cells were grown on sterilized glass coated with poly-L-lysine (Fisher Scientific, Waltham MA, USA) in a 6-well plate and inoculated at an MOI of 0.01. Cell lines were subsequently fixed in 4% paraformaldehyde at 18 hpi and 12 hpi, respectively, followed by epitope unmasking using 0.05% tween phosphate buffer solution (TPBS) for 10 min at room temperature (RT). Non-specific binding sites on cells were blocked with 0.1% power block universal blocking reagent X-10 (BioGenex, Fremont, Viruses 2022, 14, 1225 7 of 18 CA, USA) in distilled water for 30 min at RT. Cells were then incubated with mouse anti-N monoclonal antibody (mAb) SD55-197 diluted 1:500 for 2 h at RT, followed by incubation with goat anti-mouse IgG antibody conjugated with Alexa Fluor 488 at a dilution of 1:400 (Invitrogen, Carlsbad, CA, USA) for 90 min at RT and nuclei were visualized using DAPI (Thermo Fisher, Waltham, MA, USA) for 3 min at RT. Phosphate buffered saline (PBS) was used for rinsing cells between incubations and PBS with 0.2% bovine serum albumin (BSA) was used for antibody dilution. Cells were observed using confocal microscopy (Leica, Wetzlar, Germany).

Statistical Analysis
Statistical analysis was performed with two-way ANOVA and Bonferroni post hoc test or one-way ANOVA with Tukey's post hoc test using GraphPad Prism 5. Differences in means between groups were considered significant when the p value was less than 0.05. Results are expressed as mean ± SD of the means.

icPDCoV and Recombinant Viruses in In Vitro Cultures
To determine if icPDCoV, icPDCoV-S HKU17 , and icPDCoV-RBD ISU viruses would infect chicken and swine cell lines, mock and infected chicken DF-1 or swine LLC-PK1 cells were stained for viral antigen. Notably, we demonstrated that DF-1 and confirmed that the LLC-PK1 cells are susceptible to infection by the icPDCoV and chimeric viruses. Viral NP localized specifically at cytoplasmic regions in DF-1 cells (Figure 3C,E,G) and in LLC-PK1 cells ( Figure 3D,F,H) consistent with previous results for LLC-PK1 [31], while mock-inoculated cells had no NP staining ( Figure 3A,B). Taken together, these results indicate that DF-1 and LLC-PK1 cells are susceptible to icPDCoV, icPDCoV-S HKU17 , and icPDCoV-RBD ISU viral infections.
For IF staining, DF-1 and LLC-PK1 cells were grown on sterilized glass coated with poly-L-lysine (Fisher Scientific, Waltham MA, USA) in a 6-well plate and inoculated at an MOI of 0.01. Cell lines were subsequently fixed in 4% paraformaldehyde at 18 hpi and 12 hpi, respectively, followed by epitope unmasking using 0.05% tween phosphate buffer solution (TPBS) for 10 min at room temperature (RT). Non-specific binding sites on cells were blocked with 0.1% power block universal blocking reagent X-10 (BioGenex, Fremont CA, USA) in distilled water for 30 min at RT. Cells were then incubated with mouse anti-N monoclonal antibody (mAb) SD55-197 diluted 1:500 for 2 h at RT, followed by incubation with goat anti-mouse IgG antibody conjugated with Alexa Fluor 488 at a dilution of 1:400 (Invitrogen, Carlsbad, CA, USA) for 90 min at RT and nuclei were visualized using DAPI (Thermo Fisher, Waltham, MA, USA) for 3 min at RT. Phosphate buffered saline (PBS) was used for rinsing cells between incubations and PBS with 0.2% bovine serum albumin (BSA) was used for antibody dilution. Cells were observed using confocal microscopy (Leica, Wetzlar, Germany).

Statistical Analysis
Statistical analysis was performed with two-way ANOVA and Bonferroni post hoc test or one-way ANOVA with Tukey's post hoc test using GraphPad Prism 5. Differences in means between groups were considered significant when the p value was less than 0.05. Results are expressed as mean ± SD of the means.

icPDCoV and Recombinant Viruses in In Vitro Cultures
To determine if icPDCoV, icPDCoV-SHKU17, and icPDCoV-RBDISU viruses would infect chicken and swine cell lines, mock and infected chicken DF-1 or swine LLC-PK1 cells were stained for viral antigen. Notably, we demonstrated that DF-1 and confirmed that the LLC-PK1 cells are susceptible to infection by the icPDCoV and chimeric viruses. Viral NP localized specifically at cytoplasmic regions in DF-1 cells (Figure 3C,E,G) and in LLC-PK1 cells ( Figure 3D,F,H) consistent with previous results for LLC-PK1 [31], while mockinoculated cells had no NP staining ( Figure 3A,B). Taken together, these results indicate that DF-1 and LLC-PK1 cells are susceptible to icPDCoV, icPDCoV-SHKU17, and icPDCoV-RBDISU viral infections.

Replication Kinetics of Viruses in Chicken and Swine Cells
To examine whether viral replication kinetics differed among the viruses, we assessed and compared viral replication kinetics in DF-1 and LLC-PK1 cells across multiple, matched time-points. We observed that icPDCoV virus produced significantly higher viral titers in supernatant fractions harvested from LLC-PK1 cells than recombinant viruses when infected at an MOI of 0.01 ( Figure 4A). Attempts to infect DF1 cells at MOIs of 0.1, 1, 5, and 10 resulted in no detectable infectious particles released at any tested timepoints as assessed with CCIF (Data not shown), indicating that the replication of chimeric viruses in DF-1 cells was below the level of CCIF detection. Repeating the assay utilizing TCID 50 showed a low infectious titer in which icPDCoV produced minimal replication while icPDCoV-S HKU17 , and icPDCoV-RBD ISU were unable to produce additional infectious particles ( Figure 4A). A starting infectious dose of 10 MOI yielded infectious titers for each test virus which gradually declined over time, suggesting that virus particles attached to cells but failed to replicate ( Figure 4B). Distinct differences in observed cytopathic effect between each virus were noted with icPDCoV producing larger cellular syncytia than icPDCoV-S HKU17 and icPDCoV-RBD ISU which retained distinct cellular borders ( Figure 4C,E). Altogether, the results obtained showed that DF-1 cells are likely less permissive than LLC-PK1 for infection with icPDCoV-S HKU17 , and icPDCoV-RBD ISU with potential differences in Smediated syncytia formation.

Replication Kinetics of Viruses in Chicken and Swine Cells
To examine whether viral replication kinetics differed among the viruses, we assessed and compared viral replication kinetics in DF-1 and LLC-PK1 cells across multiple, matched time-points. We observed that icPDCoV virus produced significantly higher viral titers in supernatant fractions harvested from LLC-PK1 cells than recombinant viruses when infected at an MOI of 0.01 ( Figure 4A). Attempts to infect DF1 cells at MOIs of 0.1, 1, 5, and 10 resulted in no detectable infectious particles released at any tested timepoints as assessed with CCIF (Data not shown), indicating that the replication of chimeric viruses in DF-1 cells was below the level of CCIF detection. Repeating the assay utilizing TCID50 showed a low infectious titer in which icPDCoV produced minimal replication while icPDCoV-SHKU17, and icPDCoV-RBDISU were unable to produce additional infectious particles ( Figure 4A). A starting infectious dose of 10 MOI yielded infectious titers for each test virus which gradually declined over time, suggesting that virus particles attached to cells but failed to replicate ( Figure 4B). Distinct differences in observed cytopathic effect between each virus were noted with icPDCoV producing larger cellular syncytia than icPDCoV-SHKU17 and icPDCoV-RBDISU which retained distinct cellular borders ( Figure  4C,E). Altogether, the results obtained showed that DF-1 cells are likely less permissive than LLC-PK1 for infection with icPDCoV-SHKU17, and icPDCoV-RBDISU with potential differences in S-mediated syncytia formation.  [31]. The data for icPDCoV were included in 4A as a TCID50 kinetics reference. After 1 h of incubation at 37 °C, cells were washed and overlaid with MEM supplemented with 10 µg/mL trypsin (LLC-PK-1) or no trypsin (DF-1) due to cell sensitivity. The kinetics of viral replication were evaluated in supernatants collected at denoted timepoints post infection, and the viral titers were determined by TCID50 (A,B). Different lower-case letters (a, b, c) indicate significant differences between each group (n = 3 biologically independent samples) at each time point. All data points are mean ± SD. Two-way ANOVA with Bonferroni post hoc test, p ≤ 0.05. ns, not statistically significant.

In Vitro Assessment of Viral RNA Loads in Inoculated DF-1 Cells
Viral RNA loads at longitudinal time points through 24 h post inoculation were measured by RT-qPCR to assess in vitro viral RNA levels. We compared the viral RNA loads from the cell-associated fraction to the viral RNA loads in the same inoculated DF-1 cells of the cell-free fraction. Predictably, we found significant differences in the viral RNA loads between cell-associated fraction (n = 48) and cell-free fraction (n = 48) (p = 0.0  [31]. The data for icPDCoV were included in 4A as a TCID 50 kinetics reference. After 1 h of incubation at 37 • C, cells were washed and overlaid with MEM supplemented with 10 µg/mL trypsin (LLC-PK-1) or no trypsin (DF-1) due to cell sensitivity. The kinetics of viral replication were evaluated in supernatants collected at denoted timepoints post infection, and the viral titers were determined by TCID 50 (A,B). Different lower-case letters (a, b, c) indicate significant differences between each group (n = 3 biologically independent samples) at each time point. All data points are mean ± SD. Two-way ANOVA with Bonferroni post hoc test, p ≤ 0.05. ns, not statistically significant. (C-E) Light microscopy of DF-1 cells infected with specified virus at 72 h post infection with an MOI of 0.01.

In Vitro Assessment of Viral RNA Loads in Inoculated DF-1 Cells
Viral RNA loads at longitudinal time points through 24 h post inoculation were measured by RT-qPCR to assess in vitro viral RNA levels. We compared the viral RNA loads from the cell-associated fraction to the viral RNA loads in the same inoculated DF-1 cells of the cell-free fraction. Predictably, we found significant differences in the viral RNA loads between cell-associated fraction (n = 48) and cell-free fraction (n = 48) (p = 0.05) ( Figure 5A-D). Consistent with the findings from replication kinetics ( Figure 4B). (Cellassociated virus appeared to remain static over time ( Figure 6A) while cell free viral RNA increased from the 6-12 h timepoints but remained static from 12-24 h ( Figure 6B). We further examined the total release ratios of the DF-1 inoculated with icPDCoV-, icPDCoV-Viruses 2022, 14, 1225 9 of 18 S HKU17 -or icPDCoV-RBD ISU viruses to determine the presence of cell-associated virus in the inoculated DF-1 cells. Indeed, we found no differences in viral release between chimeric viruses ( Figure 7A-C). These results suggest that the reduction in infectious virus observed at MOIs lower than 40 was not attributable to viral release, suggesting that there might be other mechanistic pathways contributing to the decrease in the infectious virus in cell-free fractions; however, in this study we did not determine these mechanisms. (Figure 5A-D). Consistent with the findings from replication kinetics ( Figure 4B). (Cellassociated virus appeared to remain static over time ( Figure 6A) while cell free viral RNA increased from the 6-12 h timepoints but remained static from 12-24 h ( Figure 6B). We further examined the total release ratios of the DF-1 inoculated with icPDCoV-, icPDCoV-SHKU17or icPDCoV-RBDISU viruses to determine the presence of cell-associated virus in the inoculated DF-1 cells. Indeed, we found no differences in viral release between chimeric viruses ( Figure 7A-C). These results suggest that the reduction in infectious virus observed at MOIs lower than 40 was not attributable to viral release, suggesting that there might be other mechanistic pathways contributing to the decrease in the infectious virus in cell-free fractions; however, in this study we did not determine these mechanisms.    Figure 5A-D). Consistent with the findings from replication kinetics ( Figure 4B). (Cellassociated virus appeared to remain static over time ( Figure 6A) while cell free viral RNA increased from the 6-12 h timepoints but remained static from 12-24 h ( Figure 6B). We further examined the total release ratios of the DF-1 inoculated with icPDCoV-, icPDCoV-SHKU17or icPDCoV-RBDISU viruses to determine the presence of cell-associated virus in the inoculated DF-1 cells. Indeed, we found no differences in viral release between chimeric viruses ( Figure 7A-C). These results suggest that the reduction in infectious virus observed at MOIs lower than 40 was not attributable to viral release, suggesting that there might be other mechanistic pathways contributing to the decrease in the infectious virus in cell-free fractions; however, in this study we did not determine these mechanisms.   All data points (cts) were first converted to GE/mL, and then, the percentage of releases was calculated, according to the equation previously described. GE, genomic equivalent. One-way ANOVA with Tukey's multiple comparisons post hoc test, p ≤ 0.05, ns; not statistically significant.

Clinical Manifestations in SPF Turkey Poults and RT-qPCR of Viral RNA in Tracheal and Fecal Samples
Fecal consistency scores at 1, 3, and 5 PDI were significantly higher in groups treated with icPDCoV-SHKU17 virus or icPDCoV-RBDISU virus compared to control group ( Figure  8A). Moreover, uninfected sentinel birds that commingled with icPDCoV-SHKU17or icPD-CoV-RBDISUinfected group had higher fecal score at 2 DPI compared to control infection ( Figure 8B). Consistent with the results from the fecal consistency score, at 1 DPI, the groups infected with icPDCoV-SHKU17 or icPDCoV-RBDISU virus were more lethargic (movement less frequently) and poorly feeding more than the icPDCoV infected group, suggesting that the spike protein could play a role in virus infection. However, cloacal and tracheal swabs from experimentally infected turkey poults were negative for viral RNA throughout the study. At the cutoff value of 5.17 log10 GE/mL, no samples tested higher than control samples. The results indicate that both cloacal and tracheal swabs were negative for all viruses tested although mild clinical signs were noted. Figure 8. Fecal scores of turkey poults at 0-to 5-DPI. Fecal consistency was scored as follows: 0, normal; 1, pasty; 2, semiliquid; 3, watery), and the fecal score >2 was considered as diarrhea. Red circles, green squares, blue triangles, and black triangles represent (A) icPDCoV-inoculated, icPD-CoV-SHKU17-inoculated, icPDCoV-RBDISU-inoculated, and control groups (n = 10 biologically independent samples). (B) sentinel poults that commingled at 1 DPI (n = 4 biologically independent samples), respectively. Different lower-case letters (a; icPDCoV, b; icPDCoV-SHKU17, c; icPDCoV-RBDISU, d; mock) indicate significant differences between the given group at each time point. Significant diarrhea for each test virus was only observed at day 1 post infection but for groups icPDCoV-SHKU17 and icPDCoV-RBDISU diarrhea was also observed on days 3 and 5 in inoculated birds and day 2 in sentinel birds. Two-way ANOVA with Bonferroni post hoc test, p ≤ 0.05. ns, not statistically significant.

Clinical Manifestations in SPF Turkey Poults and RT-qPCR of Viral RNA in Tracheal and Fecal Samples
Fecal consistency scores at 1, 3, and 5 PDI were significantly higher in groups treated with icPDCoV-S HKU17 virus or icPDCoV-RBD ISU virus compared to control group ( Figure 8A). Moreover, uninfected sentinel birds that commingled with icPDCoV-S HKU17 -or icPDCoV-RBD ISU -infected group had higher fecal score at 2 DPI compared to control infection ( Figure 8B). Consistent with the results from the fecal consistency score, at 1 DPI, the groups infected with icPDCoV-S HKU17 or icPDCoV-RBD ISU virus were more lethargic (movement less frequently) and poorly feeding more than the icPDCoV infected group, suggesting that the spike protein could play a role in virus infection. However, cloacal and tracheal swabs from experimentally infected turkey poults were negative for viral RNA throughout the study. At the cutoff value of 5.17 log 10 GE/mL, no samples tested higher than control samples. The results indicate that both cloacal and tracheal swabs were negative for all viruses tested although mild clinical signs were noted. All data points (cts) were first converted to GE/mL, and then, the percentage of releases was calculated, according to the equation previously described. GE, genomic equivalent. One-way ANOVA with Tukey's multiple comparisons post hoc test, p ≤ 0.05, ns; not statistically significant.

Clinical Manifestations in SPF Turkey Poults and RT-qPCR of Viral RNA in Tracheal and Fecal Samples
Fecal consistency scores at 1, 3, and 5 PDI were significantly higher in groups treated with icPDCoV-SHKU17 virus or icPDCoV-RBDISU virus compared to control group ( Figure  8A). Moreover, uninfected sentinel birds that commingled with icPDCoV-SHKU17or icPD-CoV-RBDISUinfected group had higher fecal score at 2 DPI compared to control infection ( Figure 8B). Consistent with the results from the fecal consistency score, at 1 DPI, the groups infected with icPDCoV-SHKU17 or icPDCoV-RBDISU virus were more lethargic (movement less frequently) and poorly feeding more than the icPDCoV infected group, suggesting that the spike protein could play a role in virus infection. However, cloacal and tracheal swabs from experimentally infected turkey poults were negative for viral RNA throughout the study. At the cutoff value of 5.17 log10 GE/mL, no samples tested higher than control samples. The results indicate that both cloacal and tracheal swabs were negative for all viruses tested although mild clinical signs were noted. Figure 8. Fecal scores of turkey poults at 0-to 5-DPI. Fecal consistency was scored as follows: 0, normal; 1, pasty; 2, semiliquid; 3, watery), and the fecal score >2 was considered as diarrhea. Red circles, green squares, blue triangles, and black triangles represent (A) icPDCoV-inoculated, icPD-CoV-SHKU17-inoculated, icPDCoV-RBDISU-inoculated, and control groups (n = 10 biologically independent samples). (B) sentinel poults that commingled at 1 DPI (n = 4 biologically independent samples), respectively. Different lower-case letters (a; icPDCoV, b; icPDCoV-SHKU17, c; icPDCoV-RBDISU, d; mock) indicate significant differences between the given group at each time point. Significant diarrhea for each test virus was only observed at day 1 post infection but for groups icPDCoV-SHKU17 and icPDCoV-RBDISU diarrhea was also observed on days 3 and 5 in inoculated birds and day 2 in sentinel birds. Two-way ANOVA with Bonferroni post hoc test, p ≤ 0.05. ns, not statistically significant. Figure 8. Fecal scores of turkey poults at 0-to 5-DPI. Fecal consistency was scored as follows: 0, normal; 1, pasty; 2, semiliquid; 3, watery, and the fecal score >2 was considered as diarrhea. Red circles, green squares, blue triangles, and black triangles represent (A) icPDCoV-inoculated, icPDCoV-S HKU17 -inoculated, icPDCoV-RBD ISU -inoculated, and control groups (n = 10 biologically independent samples). (B) sentinel poults that commingled at 1 DPI (n = 4 biologically independent samples), respectively. Different lower-case letters (a; icPDCoV, b; icPDCoV-S HKU17 , c; icPDCoV-RBD ISU , d; mock) indicate significant differences between the given group at each time point. Significant diarrhea for each test virus was only observed at day 1 post infection but for groups icPDCoV-S HKU17 and icPDCoV-RBD ISU diarrhea was also observed on days 3 and 5 in inoculated birds and day 2 in sentinel birds. Two-way ANOVA with Bonferroni post hoc test, p ≤ 0.05. ns, not statistically significant.

Gross and Immunobiological Examination of PDCoV Pathology and NP Immunoreactivity in Inoculated Poults
Although turkey poults from the virus-inoculated groups showed mild signs of intestinal congestion, and intestinal gas accumulation, and those from icPDCoV-S HKU17 group showed a fecal score of 2, briefly, after standard IHC criteria and mAb incubation, IHC staining showed no cellular signal for positive PDCoV NP in the lung or small intestinal tissue sections that were included in this study (Figure 9), which is consistent with our observed in vitro results, that DF-1 cells are not permissible to the virus infections at infectious MOI of 0.1, 1, 5, and 10. These data demonstrate that while icPDCoV and chimeric viruses seemed to cause mild gross and histopathological complications in turkey poults, we were unable to detect significant viral replication.

Gross and Immunobiological Examination of PDCoV Pathology and NP Immunoreactiv in Inoculated poults
Although turkey poults from the virus-inoculated groups showed mild signs o testinal congestion, and intestinal gas accumulation, and those from icPDCoV-S group showed a fecal score of 2, briefly, after standard IHC criteria and mAb incuba IHC staining showed no cellular signal for positive PDCoV NP in the lung or small in tinal tissue sections that were included in this study (Figure 9), which is consistent our observed in vitro results, that DF-1 cells are not permissible to the virus infection infectious MOI of 0.1, 1, 5, and 10. These data demonstrate that while icPDCoV and meric viruses seemed to cause mild gross and histopathological complications in tu poults, we were unable to detect significant viral replication.

Absence of Viral RNA Titer Increases in 11-Day Old ECEs Indicate Lack of Viral Replication
We performed an RT-qPCR assay using M gene specific primers for the detectio viral RNAs from total RNA extracted from allantoic fluid (n= 29), thoracic-(n= 29), abdominal-(n= 29) tissues. A total of 87 samples from three groups were tested. E

Absence of Viral RNA Titer Increases in 11-Day Old ECEs Indicate Lack of Viral Replication
We performed an RT-qPCR assay using M gene specific primers for the detection of viral RNAs from total RNA extracted from allantoic fluid (n = 29), thoracic-(n = 29), and abdominal-(n = 29) tissues. A total of 87 samples from three groups were tested. ECEs were inoculated by the allantoic cavity with heat-inactivated (n = 3 for each virus) or viable (n = 5 for each virus) icPDCoV-, icPDCoV-S HKU17 -, icPDCoV-RBD ISU viruses, or mock-inoculated (n = 5). Our data indicated that the icPDCoV-, icPDCoV-S HKU17 -or icPDCoV-RBD ISU RNAs were detected in allantoic fluids at 3 DPI, and the viral RNA titers were significantly higher in allantoic fluids than in the control group ( Figure 10A). However, viral RNA from the heat inactivated controls remained at the same level as the live virus, suggesting that no significant replication occurred. Similarly, levels of the viral RNAs quantitated from thoracic-and abdominal tissues were not statistically different between infected and mock-inoculated groups and also did not deviate from the heat inactivated viral RNA titers ( Figure 10B,C), suggesting that icPDCoV and chimeric viruses did not replicate in the 11-day-old ECEs. Moreover, no gross lesions or pathological changes or embryo deaths were observed or recorded in the present study. These findings are consistent with a recent report of PDCoV being unable to propagate in 8-day-old ECEs [38]. were inoculated by the allantoic cavity with heat-inactivated (n= 3 for each virus) or viable (n= 5 for each virus) icPDCoV-, icPDCoV-SHKU17-, icPDCoV-RBDISU viruses, or mock-inoculated (n = 5). Our data indicated that the icPDCoV-, icPDCoV-SHKU17or icPDCoV-RBDISU RNAs were detected in allantoic fluids at 3 DPI, and the viral RNA titers were significantly higher in allantoic fluids than in the control group ( Figure 10A). However, viral RNA from the heat inactivated controls remained at the same level as the live virus, suggesting that no significant replication occurred. Similarly, levels of the viral RNAs quantitated from thoracic-and abdominal tissues were not statistically different between infected and mock-inoculated groups and also did not deviate from the heat inactivated viral RNA titers ( Figure 10B, C), suggesting that icPDCoV and chimeric viruses did not replicate in the 11-day-old ECEs. Moreover, no gross lesions or pathological changes or embryo deaths were observed or recorded in the present study. These findings are consistent with a recent report of PDCoV being unable to propagate in 8-day-old ECEs [38].

Differential Virus Replication in 5-Day-Old ECEs
After inoculating 5-day-old ECEs with serially diluted icPDCoV-, icPDCoV-SHKU17-or icPDCoV-RBDISU (100 µL/ECE), we observed 100% (5/5) ECE death in icPDCoV-inoculated ECEs at the 10 −2 through 10 −5 dilutions, whereas icPDCoV-RBDISU had limited ECE death rates of 50% (3/6) at 10 −2 dilution and 40% (2/5) at the 10 −4 dilution only, and icPDCoV-SHKU17 was unable to produce ECE death at any infectious titer by 5 DPI (Table 1). The allantoic fluid samples (at no dilution and 1:10 dilution) were tested for infectious virus in LLC-PK1 cells. Selected samples were tested by DCoV-specific RT-PCR. We found that the infectivity assay was as sensitive as the RT-PCR. These results indicate that wild type PDCoV was the most capable at entering and replicating within 5-day-old ECEs, suggesting that the RBD or S from sparrow CoVs was less able to support PDCoV entry into chicken cells or a potential synergism within and between S protein and other viral factors is needed for optimal virus replication.

Differential Virus Replication in 5-Day-Old ECEs
After inoculating 5-day-old ECEs with serially diluted icPDCoV-, icPDCoV-S HKU17 -or icPDCoV-RBD ISU (100 µL/ECE), we observed 100% (5/5) ECE death in icPDCoV-inoculated ECEs at the 10 −2 through 10 −5 dilutions, whereas icPDCoV-RBD ISU had limited ECE death rates of 50% (3/6) at 10 −2 dilution and 40% (2/5) at the 10 −4 dilution only, and icPDCoV-S HKU17 was unable to produce ECE death at any infectious titer by 5 DPI (Table 1). The allantoic fluid samples (at no dilution and 1:10 dilution) were tested for infectious virus in LLC-PK1 cells. Selected samples were tested by DCoV-specific RT-PCR. We found that the infectivity assay was as sensitive as the RT-PCR. These results indicate that wild type PDCoV was the most capable at entering and replicating within 5-day-old ECEs, suggesting that the RBD or S from sparrow CoVs was less able to support PDCoV entry into chicken cells or a potential synergism within and between S protein and other viral factors is needed for optimal virus replication.

Discussion
Emerging infectious diseases are frequently diagnosed in poultry species. Thes3 emerging infectious diseases can arise due to the cross-species transmission of infectious agents, and these crossover infections may pose a serious threat to public and animal health.  [19]. Similarly, highly pathogenic avian influenza (HPAI) A (H5N1) viruses that originated in poultry, have raised concerns for transmission to humans and have spread worldwide [39,40]. Additionally, a novel avian-origin influenza A (H7N9) emerged in 2013 in southeast China, spread among humans, and caused deaths in one third of the patients due to severe lower respiratory infection [41,42].
The S protein of coronaviruses plays a critical role in the recognition of host cellular receptors and mediation of membrane fusion [31]. Genetic recombination and mutations occur most frequently in the S protein [13]. The emergence of multiple new variants of concern of SARS-CoV-2 are attributed to multiple mutations in the S protein. Additionally, some of the S protein mutations can lead to enhanced viral infectivity or transmissibility, as evidenced by the D614G mutation in the S protein of SARS-CoV-2 [43]. Mutations within CoVs arise in three ways: (1) Intrinsically, due to the errors that occur during viral replication [44]; (2) genomic variability, which arises when two viral lineages infect the same host [45]; and (3) due to host RNA-editing systems, which are considered part of the natural viral defense mechanism [46]. It has been stated that the majority of mutations are neutral [47]. However, some mutations may increase the virulence and/or transmission and others are deleterious to the virus [43,48]. Advantageous and neutral mutations have higher frequencies [47,48].
In the present study, we used chimeric viruses that express spike (S) protein of SpCoV HKU17 or the RBD of SpDCoV ISU73347 on the genomic backbone of an infectious cDNA clone of virulent PDCoV OH-FD22 strain (icPDCoV). The first fundamental step in viral infection is the receptor interaction. We demonstrate that chicken DF-1 and confirm that swine LLC-PK1 cells-derived from chickens and pigs, respectively-are susceptible to icPDCoV-, icPDCoV-S HKU17 -and icPDCoV-RBD ISU virus infections, suggesting that chimeric viruses exhibit a broad species cell tropism, infecting cells derived from both chickens and swine. Our previous data demonstrated that PDCoV employs aminopeptidase N (APN) for cell entry, and PDCoV can infect cell lines derived from chickens and humans [19]. Therefore, chimeric viruses in the present study might employ APN as a receptor to enter DF-1 cells. However, investigators have recently found that upon infection of APN-knockout ST cells, the replication of icPDCoV-S HKU17 virus remained at the same level as its replication in wild type ST cells, but the replication of icPDCoV-RBD ISU virus increased significantly compared to the wild type ST cells [31]. Coupled with the ability of DCoVs to infect hosts from various species such as birds and mammals, this suggests that chimeric viruses might employ a broad range of receptors. Furthermore, it has been suggested that the use of APN or angiotensin-converting enzyme 2 (ACE2) as a receptor was independently selected, according to CoV evolution [19,49]. Although chimeric viruses replicated to significantly higher titers in DF-1 cells, their replication was not optimal since they only began to produce virus measurable via FFU at an MOI of 40, whereas chimeric viruses and icPDCoV showed replication in LLC-PK1 cell line at an MOI of 0.01. These findings indicate that one or more mechanistic pathways and cellular factors play an essential role in enabling the permissiveness since the cycle of viral replication is a complex process. Here, we suggest that two mechanistic pathways could have adverse consequences on permissiveness: first, we hypothesize that a strong upregulation of interferon responses, in particular type I interferons, may play a role in preventing new virus progeny infection if present within the supernatant applied to naïve cells in the FFU assay. A direct correlation between interferon responses and viral load has recently been found [50]. Second, we hypothesize that DF-1 cells may not be permissible due to reasons related to the receptors such as APN or other specific receptors. For instance, several studies demonstrated that SARS-CoV could not replicate in Madin-Darby Canine Kidney (MDCK) cells [51,52]. However, the replication of SARS-CoV was promoted and observed in engineered cells that express transmembrane serine protease 2 (TMPRSS2) that promotes change of the spike to allow cell entry [53]. The mechanisms for in vitro inhibition of the replication of chimeric and icPDCoV viruses need to be further investigated in the future.
In the inoculated turkey poults, immunohistochemistry analyses showed no signal against the N protein of PDCoV in small intestinal and lung tissues. The IHC data are consistent with the findings [31] that pigs showed no diarrhea and clinical signs, no villous atrophy, no intestinal lesions, and showed no viral antigens in small intestinal tissues after being orally/oro-nasally inoculated with the icPDCoV-S HKU17 -, and icPDCoV-RBD ISU virus. Consistent with our findings, Liang et al. [34] recently observed that PDCoV inoculated 4-day-old SPF chickens exhibited no clinical signs, and their body temperatures were the same as control group. More recently an in vitro study found that PDCoV replication was inhibited by bile acids chenodeoxycholic acid (CDCA) and lithocholic acid (LCA) due to their antiviral activity by stimulating interferons, λ3 and ISG15 [54]. Furthermore, none of the cloacal and tracheal swabs of inoculated birds and sentinel birds were clearly positive using RT-qPCR. Our results are further consistent with a previous report in which piglets did not shed viral RNA in feces following icPDCoV-S HKU17 -and icPDCoV-RBD ISU virus infections, nor did commingled pigs, indicating no pig-to-pig transmission [31]. The derivation of a single homogenous icPDCoV sequence based upon virus passaged eight times in tissue culture may have led to genomic mutations rendering the icPDCoV virus less able to infect and cause pathology in vivo than the virus inoculum derived from intestinal contents of gnotobiotic pigs. Additional studies comparing infectivity of icPDCoV and intestinal content-derived PDCoV along with virus sequence comparisons are needed to determine if specific mutations are responsible for a reduction of infectivity observed for the icPDCoV derived virus in inoculated turkey poults.
Differences between our studies and the Boley et al. results may be attributed to the use of the icPDCoV-derived clone. As mentioned above, the icPDCoV-derived virus may produce a more homogeneous starting virus (potentially clonal) versus the large intestinal contents utilized in the Boley et al. study. The viral material in that study was derived from LLC-PK1 tissue culture passage 20 virus which was also passaged through gnotobiotic pigs potentially allowing mutations and quasi-species that might have enhanced pathology in poultry. Additional differences exist between previous reported diarrhea, cloacal and tracheal viral shedding and enteric positive cells, and our present observations [25]. There are several hypotheses that account for these potential differences: First, the poults included in the present study were younger than the age of the poults in the previous study [25]. Age may play an important role in infectivity. Age is a noted factor in CoV pathology including with COVID-19 [55,56]. From this and other published studies there appears to be a window of 2-6 days in ovo, followed by days 11-24 post hatch in which poults are more susceptible to detectable infection [25,38,57]. Further infection studies with varied ages are needed to assess the ages in which chicks and poults are most susceptible to PDCoV infection. One potential age-related complication could be the quantity and localization of APN or the coreceptor. For example, the expression levels of α2,3SA-gal and α2,6SA-gal receptors in respiratory and intestinal tissues important for avian influenza infection were related to the age and species of poultry [58]. Additionally, we cannot rule out potential effects of maternal immunity in our study. Although the turkey poults in our study were from SPF flocks with no known prior exposure to PDCoV, they have not been tested to confirm they are seronegative for PDCoV. Thus, passively acquired immunity in hatching birds, which continues in the first 10 days of the life [59], could influence the susceptibility and permissiveness of these poults to viral infections. There is still a need to identify and determine of the tissue tropism of PDCoV in poultry. There is also a need to characterize the specific receptors and cellular proteases utilized by PDCoV and SpCoV. In our previous findings, we reported that PDCoV utilizes APN as an entry receptor because, in APN knockout cell lines, the susceptibility was drastically decreased and expression of APN in non-permissive cells allowed infectivity [19]. Recently, Niu et al. have found that, in APN-knockout ST cells inoculated with sparrow/swine chimeric viruses, the percentage of infectivity increased for icPDCoV-RBD ISU ; however, the infectivity was the same as in wildtype ST cells for icPDCoV-SHKU17 infection [31]. Furthermore, in the same study, they found limited replication of recombinant chimeric viruses in the respiratory tract of inoculated piglets, whereas the viruses lost their tropism for the pig intestine.
Liang et al. reported that PDCoV could be propagated in 11-day-old ECEs, but also indicated that the levels of viral RNA were low, linking the reason behind their observation to incomplete viral adaptation [34]. For this study we initially hypothesized that sparrow CoV spike proteins may bestow a binding enhancement in avian species, therefore we attempted to infect 11-day-old ECEs. We were unable to detect significant replication in 11-day-old ECEs. 11-day-old ECEs inoculated with either heat-inactivated or viable icPDCoV-, icPDCoV-S HKU17 -, icPDCoV-RBD ISU viruses, or mock via the allantoic cavity produced no embryo death or increased viral RNA loads in ECEs (live vs. inactivated viruses), thus no evidence of viral replication was detected. These findings are consistent with the recent demonstration that > 8-day-old ECEs were not susceptible to PDCoV, suggesting that once the cells differentiate and become mature in ovo, they are not susceptible to PDCoV infection [38]. Upon repeating infectivity experiments, we were able to see embryo death in 5-day-old ECEs that were inoculated with icPDCoV, and to a lesser extent with icPDCoV-RBD ISU but not icPDCoV-S HKU17 . Further research is needed to determine whether replacement of the full S protein from ISU SpCoV would reduce replication levels to that of HKU17 SpCoV spike or whether S RBD insertions are simply more viable than full S replacements. Rather than sparrow CoV-derived RBD or spike protein bestowing enhanced binding of PDCoV to ECEs, this data indicates that an S-dependent adaptation in PDCoV has allowed for more efficient cross-species spread than ancestral sparrow S proteins.

Conclusions
Our study presents the first qualitative and quantitative data for in vitro replication kinetics in chicken DF-1 cells, and ECEs of chimeric icPDCoV-S HKU17 -, icPDCoV-RBD ISU viruses. Our results indicated that the infectious clone-derived wildtype or chimeric viruses do not replicate in 11-day-old ECEs but icPDCoV and icPDCoV-RBD ISU do replicate in 5-day-old ECEs with icPDCoV-S HKU17 unable to replicate efficiently even at the highest starting inoculum. Although the viruses replicated in chicken DF-1 cells, they required high starting MOIs and failed to maintain efficient replication. The findings of this study are important to better understand cross-species threat of delta-coronaviruses based on icPDCoV and the chimeric viruses bearing swine vs. avian RBD or S, respectively. Future work should be directed toward understanding genetic differences between infectious clone-derived PDCoV and chimeras versus porcine intestinal content-derived virus, along with the mechanistic pathways and cell and host factors that decrease infection in DF-1 cells and restrict infection of differentiated ECEs by the chimeric viruses. In addition, understanding the receptors utilized by chimeric viruses for cellular entry will be an important next step. This could aid in the design of novel antiviral protection against delta-CoVs.