key: cord-1012599-kvlv8vtz
authors: Cross, Robert W; Agans, Krystle N; Prasad, Abhishek N; Borisevich, Viktoriya; Woolsey, Courtney; Deer, Daniel J; Dobias, Natalie S; Geisbert, Joan B; Fenton, Karla A; Geisbert, Thomas W
title: Intranasal exposure of African green monkeys to SARS-CoV-2 results in acute phase pneumonia with shedding and lung injury still present in the early convalescence phase
date: 2020-08-13
journal: Res Sq
DOI: 10.21203/rs.3.rs-50023/v2
sha: 7b6096b464b697e7ec925eff92bea6924ab8faec
doc_id: 1012599
cord_uid: kvlv8vtz

We recently reported the development of the first African green monkey (AGM) model for COVID-19 based on a combined liquid intranasal (i.n.) and intratracheal (i.t.) exposure to severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2). Here, we followed up on this work by assessing an i.n. particle only route of exposure using the LMA mucosal atomization device (MAD). Six AGMs were infected with SARS-CoV-2; three animals were euthanized near the peak stage of virus replication (day 5) and three animals were euthanized during the early convalescence period (day 34). All six AGMs supported robust SARS-CoV-2 replication and developed respiratory disease. Evidence of coagulation dysfunction as noted by a transient increases in aPTT and circulating levels of fibrinogen was observed in all AGMs. The level of SARS-CoV-2 replication and lung pathology was not quite as pronounced as previously reported with AGMs exposed by the combined i.n. and i.t. routes; however, SARS-CoV-2 RNA was detected in nasal swabs of some animals as late as day 15 and rectal swabs as late as day 28 after virus challenge. Of particular importance to this study, all three AGMs that were followed until the early convalescence stage of COVID-19 showed substantial lung pathology at necropsy as evidenced by multifocal chronic interstitial pneumonia and increased collagen deposition in alveolar walls despite the absence of detectable SARS-CoV-2 in any of the lungs of these animals. These findings are consistent with human COVID-19 further demonstrating that the AGM faithfully reproduces the human condition.

The unprecedented pandemic of COVID-19 caused by severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) has had devastating effects on public health and the global economy. Considerable resources have been allocated by Governments, philanthropic organizations, and private companies in an attempt to expedite the development of vaccines and treatments to combat COVID-19. With the rapid development of 24 preventative vaccines in clinical evaluation [1] , and nearly 200 more in the pipeline [2] , coupled with the availability of nearly 300 candidate antivirals and disease modulators [2] it is impossible to investigate the safety and e cacy of all of these various interventions in humans. Both small animal models and nonhuman primates (NHP) may prove valuable in triaging the most promising medical countermeasures prior to use in humans. Hamsters and ferrets are currently being used as immunocompetent small animal models of COVID-19 [3] [4] [5] while several NHP models have been quickly developed [6] [7] [8] [9] [10] [11] [12] . Among the nonhuman primate models evaluated the African green monkey (AGM) appears to best recapitulate the most salient features of human COVID-19 [10] [11] [12] .

We recently reported the development of the rst AGM model for COVID-19 and showed that backchallenge of animals with SARS-CoV-2 ve weeks after initial exposure resulted in protection from reinfection [10] . In this study the AGMs were exposed to SARS-CoV-2 by a combination of the intranasal (i.n.) and intratracheal (i.t.) routes with the virus delivered in liquid media. As a natural extension of this initial work we sought to assess the pathogenesis of SARS-CoV-2 in AGMs exposed by the i.n. route only using the LMA Mucosal Atomization Device (MAD). Previous studies with another respiratory virus, Nipah virus, showed that there were no major differences in disease pathogenesis when virus was delivered to AGMs by a combined liquid-based i.n. and i.t. delivery [13] or by the LMA MAD system [14] . The LMA MAD was developed for the e cient and safe delivery of test particles and is currently employed to administer US FDA approved drugs for i.n. delivery. The LMA MAD delivers atomized particles that range in size from 30 to 100 μm, which is highly consistent with the size of droplets exhaled by humans due to coughing [15] . In addition, in our previous work as the AGMs were back challenged with SARS-CoV-2 it was impossible to assess tissue pathology during convalescence after primary challenge. Here, we focused on assessing the pathogenesis of SARS-CoV-2 infection in AGMs when administered as 30 to 100 μm particles and on evaluating virus shedding and lung pathology during early convalescence.

The virus (SARS-CoV-2/INMI1-Isolate/2020/Italy) was isolated on January 30, 2020 from the sputum of the rst clinical case in Italy, a tourist visiting from the Hubei province of China that developed respiratory illness while traveling [16] . The virus was initially passaged twice (P2) on Vero E6 cells; the supernatant and cell lysate were collected and clari ed following a freeze/thaw cycle. This isolate is certi ed mycoplasma and Foot-and-Mouth Disease virus free. The complete sequence was submitted to GenBank (MT066156) and is available on the GISAID website (BetaCoV/Italy/INMI1-isl/2020: EPI_ISL_410545) upon registration. For in vivo challenge, the P2 virus was propagated on Vero E6 cells and the supernatant was collected and clari ed by centrifugation making the virus used in this study a P3 stock.

Animal challenge SARS-CoV-2 seronegative AGMs (Chlorocebus aethiops) (6 females) (St Kitts origin, Worldwide Primates, Inc.) were randomized into two cohorts where one group (n=3) was scheduled for euthanasia at 5 dpi and the other at 34 dpi. Animals were anesthetized with ketamine and inoculated with a target dose of 3.0 x 10 6 PFU of SARS-CoV-2 (SARS-CoV-2/INMI1-Isolate/2020/Italy) using the LMA MAD, with the dose being equally divided between each nostril. All animals were longitudinally monitored for clinical signs of illness including temperature (measured by surgically implanted DST micro-T small implantable thermo loggers (Star-Oddi, Gardabaer, Iceland)), respiration quality, and clinical pathology. All measurements requiring physical manipulation of the animals were performed under sedation by ketamine. Mucosal swabs were obtained using sterile swabs inserted into the mucosal cavity, gently rotated to maximize contact with the mucosal surface, and deposited into 2.0 mL screw-top tubes containing sterile MEM media supplemented to 2% with FBS.

On speci ed procedure days (days 0, 2, 3, 4, 5, 7, 12, 15, 21, 28, 34), 100 μl of blood was added to 600 μl of AVL viral lysis buffer (Qiagen) for virus inactivation and RNA extraction. Following removal from the high containment laboratory, RNA was isolated from blood and swabs using the QIAamp viral RNA kit (Qiagen).

Detection of SARS-CoV-2 load RNA was isolated from blood and mucosal swabs and assessed using the CDC SARS-CoV-2 N2 assay primers/probe for reverse transcriptase quantitative PCR (RT-qPCR) [17] . SARS-CoV-2 RNA was detected using One-step probe RT-qPCR kits (Qiagen) run on the CFX96 detection system (Bio-Rad), with the following cycle conditions: 50°C for 10 minutes, 95°C for 10 seconds, and 45 cycles of 95°C for 10 seconds and 55°C for 30 seconds. Threshold cycle (C T ) values representing SARS-CoV-2 genomes were analyzed with CFX Manager Software, and data are presented as GEq. To generate the GEq standard curve, RNA was extracted from supernatant derived from Vero E6 cells infected with SARS-CoV-2/INMI1-Isolate/2020/Italy was extracted and the number of genomes was calculated using Avogadro's number and the molecular weight of the SARS-CoV-2 genome.

Infectious virus was quantitated by plaque assay on Vero E6 cells (ATCC CRL-1586) from all blood plasma and mucosal swabs, and bronchoalveolar lavage (BAL) samples. Brie y, increasing 10-fold dilutions of the samples were adsorbed to Vero E6 cell monolayers in duplicate wells (200 μl). Cells were overlaid with EMEM medium plus 1.25% Avicel, incubated for 2 days, and plaques were counted after staining with 1% crystal violet in formalin. The limit of detection for this assay is 25 PFU/ml.

Total white blood cell counts, white blood cell differentials, red blood cell counts, platelet counts, hematocrit values, total hemoglobin concentrations, mean cell volumes, mean corpuscular volumes, and mean corpuscular hemoglobin concentrations were analyzed from blood collected in tubes containing EDTA using a Vetscan HM5 hematologic analyzer (Abaxis). Serum samples were tested for concentrations of albumin, amylase, alanine aminotransferase (ALT), aspartate aminotransferase (AST), alkaline phosphatase (ALP), blood urea nitrogen (BUN), calcium, creatinine (CRE), C-reactive protein (CRP), gamma-glutamyltransferase (GGT), glucose, total protein, and uric acid by using a Piccolo pointof-care analyzer and Biochemistry Panel Plus analyzer discs (Abaxis). Partial pressures of CO 2 and O 2 were obtained using an iSTAT Alinity hematological analyzer (Abbott).

Neutralization titers were calculated by determining the dilution of serum that reduced 50% of plaques (PRNT 50 ). A standard 100 PFU amount of SARS-CoV-2 was incubated with two-fold serial dilutions of serum samples for one hour. The virus-serum mixture was then used to inoculate Vero E6 cells for 60 minutes. Cells were overlaid with EMEM medium plus 1.25% Avicel, incubated for 2 days, and plaques were counted after staining with 1% crystal violet in formalin.

ELISA SARS-CoV-2-speci c IgG antibodies to nucleoprotein were measured in sera by ELISA at the indicated time points. Nucleoprotein ELISA kits were kindly provided by Zalgen Labs, LLC. Sera were initially diluted 1:100 and then two-fold through 1:25,600 in 4 in (1 x PBS with 0.02% Tween-20). After a one-hour incubation, plates were washed six times with wash buffer (1 x PBS with 0.2% Tween-20) and incubated for an hour with a 1:5000 dilution of horseradish peroxidase conjugated anti-primate IgG antibody (Fitzgerald Industries International; Cat: 43R-IG020HRP). Tetramethylbenzidine was used to develop the reaction; the reaction was stopped with methane-sulfonic acid and plates were read at a wavelength of 450 nm. Absorbance values were normalized by blank-subtracting values from wells incubated with sera from a SARS-CoV-2-naïve animal at the corresponding serum dilution. End-point titers were de ned as the reciprocal of the last adjusted serum dilution with a value ≥ 0.20.

Necropsy was performed on all subjects euthanized at 5 dpi and 34 dpi. Tissue samples of all major organs were collected for histopathologic and immunohistochemical (IHC) examination and were immersion-xed in 10% neutral buffered formalin for > 7 days. Specimens were processed and embedded in para n and sectioned at 5 μm thickness. For IHC, speci c anti-SARS immunoreactivity was detected using an anti-SARS nucleocapsid protein rabbit primary antibody at a 1:800 dilution for 60 minutes (Novusbio). The tissue sections were processed for IHC using the ThermoFisher Scienti c Lab Vision Autostainer 360 (ThermoFisher Scienti c). Secondary antibody used was biotinylated goat 

We challenged six healthy, adult AGMs with a target dose of 3.0 x 10 6 PFU of SARS-CoV-2 (SARS-CoV-2/INMI1-Isolate/2020/Italy) via intranasal inoculation with the LMA MAD (actual delivered dose of 2.8 x 10 6 PFU). Three animals were euthanized at 5 days post-infection (dpi) which is thought to be the approximate time point of peak disease in AGMs [10] , while the remaining three animals were euthanized at 34 dpi during early convalescence. Blood and mucosal swabs were sampled from all animals on days 0, 2, 3, 4, 5, and additionally on days 7, 9, 12, 15, 21, 28, and 34 for AGM-4, AGM-5, and AGM-6. BAL uid collection was performed on days -8, 3, and 5 for all animals, as well as 7 dpi for AGM-4, AGM-5 and AGM-6. Consistent with our previous report describing the development of the combined intranasal and intratracheal SARS-CoV-2 challenge model in AGMs [10] , we did not observe overt signs of clinical illness in any AGMs in this study, other than decreased appetite or brief (single day) anorexia (Supp Table 1 ). Temperature was longitudinally monitored in 15 minute increments for the entire study duration using surgically implanted temperature loggers; several animals (AGM-4, AGM-6) experienced brief (< 2 hours) periods of mildly elevated temperatures at 3 dpi, and two animals (AGM-2, AGM-3) exhibited an abnormal temperature cycling pattern at 3 dpi (Supp Figure 1) .

As in our previous report, transient shifts in leukocyte populations, predominately manifested as lymphocytopenia (5/6 animals), thrombocytopenia (3/6 animals), and granulocytosis (de ned by neutrophilia, eosinophilia, and/or basophilia) (6/6 animals) were observed, while markers for renal (BUN, CRE) and hepatic function (ALT, AST, ALP, GGT) remained unchanged for the most part, with the exception of mild (≤ 2-fold) increases in ALT (2/6 animals), and mild to moderate (1 to 16-fold) increases in CRP, a marker of acute systemic in ammation (5/6 animals) (Supp Table 1 ), although statistical signi cance was not reached for most parameters at most time points (Figure 1 ). In addition, hypercapnia (de ned here as ≥ 4 mmHg increase in dissolved CO 2 ) was observed in 3/6 animals (Supp Table 1 ), which as we observed previously [10] , appeared to follow a biphasic pattern ( Figure 1A , data shown as fold-change from baseline]).

All animals exhibited normal prothrombin times (PT) as compared to their individual baseline values; however, mild to moderate prolongation of the activated partial thromboplastin time (aPTT) was also observed in all animals through the acute phase of disease, most prominently in AGM-1 and AGM-2, indicating possible disorder of the intrinsic coagulation pathway ( Figure 1H , I); this was mirrored by increased levels of circulating brinogen ( Figure 1J ). We previously showed that the pathways connected to IL-6 production are activated during SARS-CoV-2 infection of AGMs [10] , indicating possible mechanisms of coagulopathy in the current study.

All animals seroconverted, with weakly neutralizing titers (as quanti ed by PRNT 50 ) being detected as early as 5 dpi and gradually increasing in potency by 34 dpi, with terminal neutralizing antibody titers ranging from ~1:16-1:128 (Figure 2A-E) . We next quanti ed SARS-CoV-2 nucleoprotein speci c IgG by ELISA ( Figure 2F ). Seroconversion was not detected until day 15 in two animals (AGM-4 & AGM-5). Interestingly, not until 34 dpi was a modest level (1:800) of seroconversion detected in the third animal.

Quanti cation of viral load in blood, mucosal swabs, and lungs Viral RNA (vRNA) was puri ed from whole blood, oral, nasal and rectal mucosa, and BAL uid from all collection days, as well as from lung tissue harvested at necropsy. As we previously reported [10] , we were unable to detect SARS-CoV-2 vRNA in whole blood by RT-qPCR, nor were we able to recover infectious virus in the plasma fraction by plaque assay, con rming a lack of either cell-associated or freelycirculating virus in the peripheral blood. SARS-CoV-2 vRNA and infectious virus was detected in the nasal mucosa from all animals as early as 2 dpi, with vRNA persisting in a single animal up to 15 dpi ( Figure  3A , B). Likewise, vRNA was detected in oral swabs from all animals beginning 2-3 dpi before falling below the limit of detection by 7 dpi, while low quantities of infectious virus (1-2 log 10 PFU/mL) were only isolated from three animals (AGM-4, AGM-5, and AGM-6) ( Figure 3C, D) . Remarkably, vRNA was transiently shed from the lower gastrointestinal tract up to 28 dpi (AGM-4 and AGM-6), although infectious virus could only be recovered from the rectal swab of a single animal (AGM-3) 4-5 dpi ( Figure  3E , F). vRNA was detected in BAL uid from 4/6 animals 3 dpi and up to 7 dpi in all three animals held past 5 dpi, while infectious virus was recovered from 3/6 animals ( Figure 3G, H) . Detectable quantities of vRNA were absent from lungs harvested during necropsy of AGMs euthanized 34 dpi, while 6-9 log 10 GEq/g were detected from all three animals euthanized at 5 dpi ( Figure 3I ).

Necropsy was performed on all animals following euthanasia, and lungs were collected for gross examination and histopathological analysis. Consistent with our previous study utilizing a combined i.n. and i.t. inoculation route [10] , all AGMs displayed varying degrees of pulmonary consolidation with hyperemia and hemorrhage, characterized by depressed and patchy dark red to light pink regions (Fig. 4,  arrows) . In all AGMs, the most severe lesions were located in the dorsal aspects of the lower lung lobes. A board-certi ed veterinary pathologist approximated lesion severity for each lung lobe (Supp Table 2 ). All AGMs at 5 dpi also had segmentally accid and gas distention of small intestines. There were no other signi cant gross lesions.

Histologically, all three AGMs euthanized at 5 dpi developed mild multifocal neutrophilic bronchointerstitial pneumonia ( Figure 5A-E, O) . Histologic features include acute in ammation centered within the airways of terminal bronchioles with occasional ooding of adjacent alveolar spaces with neutrophils, macrophages, brin, edema, hemorrhage, mucous and rarely multinucleated giant cells (5A, B). In lesser-affected regions alveolar septate were expanded with mixed in ammatory cells and alveolar spaces contain increased numbers of alveolar macrophages with scattered red blood cells. Ulcerative tracheobronchitis was also present in all three AGMs and characterized by multifocal epithelial erosion associated with underlying hemorrhage, brin accumulation and in ltrating acute in ammation. Polymerized brin, highlighted by IHC, colocalized with acute in ammation within the bronchial lumen, alveolar spaces, alveolar walls and ulcerated regions of the trachea and bronchus ( Figure 5C ). Fibrin was also present within medium and small caliber vessels but was not associated with an obvious adherent thrombus. Trichrome stain of representative lung sections identi ed modest collagen deposition within multifocal regions of alveolar septae ( Figure 5D ). IHC for SARS-CoV-2 antigen was positive in all three AGMs associated with pulmonary lesions. Positive IHC labeling was noted diffusely within the cytoplasm of respiratory epithelium of the bronchus ( Figure 5O ) and less in type I and type II pneumocytes ( Figure  5E ).

Histologically, all three AGMs euthanized at 34 dpi developed moderate multifocal chronic interstitial pneumonia Figure (5F-J) . Histologic features include expansion of alveolar septae with macrophages, lymphocytes, and very rarely neutrophils ( Figure 5F, G) . Wispy, pale eosinophilic, acellular material also multifocally expanded the alveolar walls and stained as immature collagen with trichrome staining ( Figure 5I ). Polymerized brin was present within medium and small caliber vessels but was not associated with an obvious adherent thrombus ( Figure 5H ). No immunolabeling for SARS-CoV-2 was noted with IHC in any of the examined tissue sections from this 34 dpi cohort ( Figure 5J ).

We previously reported the development of the AGM as a promising animal model of human COVID-19 [10] . Other studies have subsequently reported similar ndings [11, 12] . The focus of the current study was to assess a more natural route of human exposure, speci cally an exposure mimicking an infection resulting from mucosal exposure to infectious droplets expelled from close quarter exposure to a sneeze, cough, or even speech in order to begin characterization of lung pathology in the early convalescence phase of COVID-19. The disease resulting from the i.n. MAD challenge was largely re ective of that observed with the combination of the i.t. and i.n. routes except it appeared to be somewhat milder in terms of length of any fever, less severe signs of pneumonia as evidenced by reduced alveolar ooding, and a lower prevalence of SARS-CoV-2 infection. [10] . However, the MAD-infected AGMs still developed virus-induced pneumonia and viral shedding was detected into the early convalescence period. While it appears that inclusion of direct i.t. instillation of SARS-CoV-2 as an exposure route may result in a more severe disease in AGMs, it is also possible that animal to animal variability may have contributed to the modest difference between the studies. SARS-CoV-2 infection of humans results in a wide spectrum of disease ranging from asymptomatic to severe fatal disease so it is not unexpected that there could be variability among AGMs as well. While the current study employed female AGMs because of animal availability at the time the work was initiated gender did not affect the outcome when compared to similar studies [10] [11] [12] .

Coagulation dysfunction is a consistent observation in human COVID-19 and has been associated with disease severity [18] [19] [20] [21] [22] . Here, we performed a limited analysis of blood clotting times (PT and aPTT) and circulating brinogen levels to begin to characterize the coagulopathy in SARS-CoV-2-infected AGMs. Transient increases in aPTT and in circulating brinogen levels were observed during the acute phase of infection. Increases in PT and/or aPTT have been linked to severe human COVID-19 cases in some but not all studies [18] [19] [20] [21] [22] . However, nearly all severe COVID-19 cases have been associated with high levels of brinogen [20] [21] [22] .

Our ndings regarding lung injury in the three AGMs that were euthanized at 34 dpi during early convalescence are consistent with the limited human COVID-19 studies that have been reported so far. For example, a recent study of fty-seven COVID-19 patients in China was completed during the early convalescence phase, approximately 30 days after discharge [23] . The study included 40 non-severe cases and 17 severe cases. Thirty-one patients (54.3%) had abnormal CT ndings while abnormalities were detected in the pulmonary function tests in 43 (75.4%) of the patients. In a second human study, 21 patients recovering from COVID-19 (without severe respiratory distress during the disease course), had lung abnormalities visible on chest CT at 10 days after initial onset of symptoms [24] . While other studies suggest that some of the abnormalities may be resolved over time [25, 26] more research needs to be conducted in this area.

Regarding histopathology, human data is particularly sparse. One small study performed thoracoscopies with blebs resection and pleurectomies on performed on the 16th and 23rd days from symptoms onset of two patients [27] . Despite well-known pulmonary damages induced during the acute phase of COVID-19, the late-phase gross and histological changes include nonspeci c chronic reparative lesions, similarly to what we have described in the AGMs at 34 dpi. Grossly in the human study, there was non-speci c diffuse pulmonary congestion, edema and hemorrhagic necrosis. Histologically, the main lesions were focused on alveolar damage with mildly thickened alveolar interstitial tissues with brosis and mononuclear cellular in ltration (lymphocytes, plasma cells and multinucleate giant cells). Intravascular hemorrhagic thrombosis was also noted in these specimens.

In summary, we have expanded on our previous development of the combined i.n. and i.t. inoculation model of SARS-CoV-2 in AGMs. Importantly, while AGMs challenged with SARS-CoV-2 via the LMA MAD exhibited apparently milder clinical illness and disease, hallmark features from our previous study were still apparent, notably the development of viral pneumonia during the acute phase. The AGM COVID-19 model should be useful in future studies to assess disease and develop interventions that improve recovery. Declarations infectivity assays. KNA optimized and performed the PCR. NSD optimized and performed the immunohistochemistry. CW performed ELISAs. KAF performed necropsies and analysis of the gross pathology, histopathology, and immunohistochemistry. All authors analyzed the clinical pathology, virology, and immunology data. RWC, ANP, KAF, and TWG, wrote the paper. All authors had access to all of the data and approved the nal version of the manuscript.

This study was supported by funds from the Department of Microbiology and Immunology, University of Texas Medical Branch at Galveston, Galveston, TX to TWG. Operations support of the Galveston National Laboratory was supported by NIAID/NIH grant UC7AI094660.

The data supporting the conclusions of this article are included within the article.

All animal studies were approved by the University of Texas Medical Branch (UTMB) Institutional Animal Care and Use Committee and adhere to the NIH Guide for the Care and Use of Laboratory Animals.

Not applicable. Statistical signi cance was determined in Graphpad Prism 8.4.3 by mixed-effects analysis with the Geisser-Greenhouse correction without the assumption of sphericity, with multiple comparisons made using Dunnett's post-hoc test and all comparisons made to baseline values (0 dpi). Asterisks denote signi cance: * = p ≤ 0.05, ** = p ≤ 0.01, *** = p ≤ 0.001. Two-tailed p-values were computed for all comparisons. Gross lung pathology in AGMs infected with SARS-CoV-2. Dorsal view of lungs from AGM-1 (A), AGM-2 (B) and AGM-3(C) euthanized at 5 dpi with SARS-CoV-2 exhibiting mild to moderate locally extensive pulmonary consolidation with hyperemia and hemorrhage. Dorsal view of lungs from AGM-4 (D), AGM-5 (E) and AGM-6 (F) euthanized at 34 dpi with SARS-CoV-2 exhibiting mild to marked locally extensive pulmonary consolidation with hyperemia and hemorrhage. Dorsal view of control lungs with no signi cant lesions from SARS-CoV-2 negative AGM (G). Figure 5 

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The authors would like to thank the UTMB Animal Resource Center for veterinary support for surgery to implant temperature data loggers and husbandry support of laboratory animals and Dr. Kevin Melody for assistance with animal studies. The virus used in this publication was kindly provided by the European Virus Archive goes Global (EVAg) project that has received funding from the European Union's Horizon 2020 research and innovation program under grant agreement No 653316.Author contributions RWC and TWG conceived and designed the study. DJD, JBG, and TWG performed the SARS-CoV-2 challenge experiments. RWC, DJD, CW, JBG, and TWG performed animal procedures and clinical observations. KNA and VB performed the clinical pathology assays. VB performed the SARS-CoV-2

The authors declare no competing interests.Comparative pulmonary histologic lesions in AGMs infected with SARS-CoV-2. Representative tissues of AGM from 5 dpi (A-E & O) and 34 dpi (F-J). SARS-CoV-2 naïve tissues from an AGM (K-N). H&E staining at