Skip to main content
NCBI home page

This is a preprint.

It has not yet been peer reviewed by a journal.

The National Library of Medicine is running a pilot to include preprints that result from research funded by NIH in PMC and PubMed.

bioRxiv logoLink to bioRxiv
[Preprint]. 2021 Jan 11:2021.01.09.426058. [Version 1] doi: 10.1101/2021.01.09.426058

Intranasal ChAdOx1 nCoV-19/AZD1222 vaccination reduces shedding of SARS-CoV-2 D614G in rhesus macaques

Neeltje van Doremalen 1, Jyothi N Purushotham 1,2, Jonathan E Schulz 1, Myndi G Holbrook 1, Trenton Bushmaker 1, Aaron Carmody 3, Julia R Port 1, Claude K Yinda 1, Atsushi Okumura 1, Greg Saturday 4, Fatima Amanat 5,6, Florian Krammer 5, Patrick W Hanley 4, Brian J Smith 4, Jamie Lovaglio 4, Sarah L Anzick 3, Kent Barbian 3, Craig Martens 3, Sarah Gilbert 2, Teresa Lambe 2, Vincent J Munster 1
PMCID: PMC7808328  PMID: 33447831

Abstract

Intramuscular vaccination with ChAdOx1 nCoV-19/AZD1222 protected rhesus macaques against pneumonia but did not reduce shedding of SARS-CoV-2. Here we investigate whether intranasally administered ChAdOx1 nCoV-19 reduces shedding, using a SARS-CoV-2 virus with the D614G mutation in the spike protein. Viral load in swabs obtained from intranasally vaccinated hamsters was significantly decreased compared to controls and no viral RNA or infectious virus was found in lung tissue, both in a direct challenge and a transmission model. Intranasal vaccination of rhesus macaques resulted in reduced shedding and a reduction in viral load in bronchoalveolar lavage and lower respiratory tract tissue. In conclusion, intranasal vaccination reduced shedding in two different SARS-CoV-2 animal models, justifying further investigation as a potential vaccination route for COVID-19 vaccines.

The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) pandemic initiated the rapid development of vaccines based on a wide variety of platforms. Just 11 months later after the release of the first genome sequence, 13 vaccines are in phase III clinical trials and results of phase 3 clinical trial data for three different vaccines have been released1–3. These data suggest that vaccines based on the spike (S) protein of SARS-CoV-2, which generate a neutralizing antibody response, can reach an efficacy of up to 95%. Furthermore, several vaccines developed by Astrazeneca/Oxford, Bharat Biotech, CanSinoBIO, the Gamaleya Research Institute, Moderna/VRC, Pfizer/BioNTech, Sinopharm, Sinovac, and the Vector Institute have now been approved, fully or for emergency use.

In humans, most SARS-CoV-2 infections will present as asymptomatic or mild upper respiratory tract infection but are still accompanied by shedding of virus4. Depending on the study, shedding in asymptomatic infections was of shorter duration, but often to similar viral loads initially4. Asymptomatic as well as pre-symptomatic shedding has been associated with SARS-CoV-2 transmission5–7.

In preclinical non-human primate (NHP) challenge experiments, several vaccines were successful at preventing disease and reducing or preventing virus replication in the lower respiratory tract. However, subgenomic and genomic viral RNA was detected in nasal samples of all NHP experiments, dependent on vaccine dose8–13. Subgenomic viral RNA is indicative of replicating virus in the upper respiratory tract. It is currently unclear whether the detection of shedding in NHPs translate directly to humans.

It is possible that vaccination will result in attenuation or prevention of disease, but infection of the upper respiratory tract will occur even after vaccination possibly resulting in transmission. Currently, the majority of COVID-19 vaccines in development utilize an intramuscular (IM) injection, which predominantly produces a systemic IgG response and a poor mucosal response14. For a vaccine to elicit mucosal immunity, antigens will need to be encountered locally at the initial site of replication: the upper respiratory tract (URT). Here, we evaluate the potential of using COVID-19 vaccine candidate ChAdOx1 nCoV-19 as an intranasal (IN) vaccine in the hamster and rhesus macaque models.

Results

To evaluate the efficacy of an IN vaccination with ChAdOx1 nCoV-19, three groups of 10 Syrian hamsters15 were vaccinated with a single dose; group 1 received ChAdOx1 nCoV-19 via the IN route, group 2 received the same dose of vaccine via the IM route, and group 3 received control vaccine ChAdOx1 GFP via the IM route. Binding antibodies against SARS-CoV-2 S protein in peripheral blood were measured at −1 days post infection (DPI). Vaccination via either route resulted in high IgG titers (25,600–204,800) with no significant difference between vaccination routes (Figure 1A). Likewise, high neutralizing antibodies titers were detectable at −1 DPI. Intriguingly, neutralizing antibody titers were significantly higher in animals that received an IN vaccination (Figure 1B). For IN inoculation of Syrian hamsters 28 days post vaccination, we used isolate SARS-CoV-2/human/USA/RML-7/2020 which contains the D614G mutation in the S protein. Animals who received ChAdOx1 GFP started losing weight at 3 DPI and did not regain weight until 8 DPI. None of the vaccinated animals lost weight throughout the course of the experiment (Figure 1C). Six animals per group were swabbed daily up to 7 DPI. Viral RNA was detected in swabs from all animals. A significantly reduced amount of viral RNA was detected in nasal swabs from IN-vaccinated animals compared to control animals on 1–3 and 6–7 DPI. However, a significant reduction of viral RNA detected in oropharyngeal swabs from IM-vaccinated animals compared to control animals was only detected at 7 DPI (Mixed-effect analysis, p-value <0.05). When the area under the curve (AUC) was calculated as a measurement of total amount of viral RNA shed, IN-vaccinated animals shed significantly less than control animals (Kruskall-Wallis test, p=0.0074). Although viral RNA is an important measurement, the most crucial measurement in swabs is infectious virus. We found a significant difference between infectious virus detected in oropharyngeal swabs of IN-vaccinated animals compared to controls daily (Mixed-effect analysis, p-value<0.05). Likewise, the amount of infectious virus shed over the course of the experiment was significantly lower in IN-vaccinated animals than controls (Figure 1D, Kruskall-Wallis test, p=0.002). In contrast, we did not find a significant difference in AUC for viral RNA and infectious virus when comparing control and IM-vaccinated animals (Figure 1E). At five DPI, four animals in each group were euthanized. Viral load and infectious virus titer were high in lung tissue of control animals. We were unable to detect viral RNA or infectious virus in lung tissue from IN-vaccinated animals. Two animals in the IM group were weakly positive for genomic RNA, but not for subgenomic RNA or infectious virus (Figure 1F).

Figure 1. SARS-CoV-2 challenge of Syrian hamsters vaccinated with ChAdOx1 nCoV-19.

Figure 1.

Open in a new tab

Hamsters were vaccinated via the IN route (red), IM route (blue) or with control vaccine ChAdOx1 GFP via the IM route (green). A. Binding antibody titers against SARS-CoV-2 S protein. B. Virus neutralizing antibody titers. C. Relative weight upon challenge with SARS-CoV-2. A-B. Shown is geometric mean and 95% confidence interval. # = p-value <0.05 between IN and control group; * = p-value <0.05 between vaccinated groups and control group. D. Viral load and viral titer in oropharyngeal swabs. Shown is geometric mean (symbols) and 95% confidence interval (shade). E. Area under the curve analysis of viral load and titer shedding in oropharyngeal swabs. F. Viral load and titer in lung tissue, isolated at 5 DPI. E-F. Dashed line = median; dotted line = quartiles. Statistical analyses done using mixed-effect analyses (C), two-way ANOVA (D), or Kruskal-Wallis test (E-F). * = p value <0.05; ** = p-value < 0.01; *** = p-value < 0.001.

Lung tissue obtained at 5 DPI was then evaluated for pathology. Lesions were found in the lungs of control animals throughout (40–70% of tissue). Interstitial pneumonia was present in all animals, as well as edema, type II pneumocyte hyperplasia, and perivascular leukocyte infiltration, similar to what has been observed previously15. In stark contrast, no lesions or pathology were observed in lung tissue of vaccinated animals (Figure 2 and Table S1). SARS-CoV-2 N antigen in lung tissue was only found in control animals (20–70% of lung tissue was immunoreactive), but not for vaccinated animals (Figure 2 and Table S1).

Figure 2. Pulmonary effects of direct intranasal challenge with SARS-CoV-2 in Syrian hamsters.

Figure 2.

Open in a new tab

A-C. H&E; D-F. IHC. A/B. No pathology. C. Moderate to marked interstitial pneumonia. D/E. No immunoreactivity. F. Numerous immunoreactive bronchiolar epithelial cells and Type I&II pneumocytes. Bar = 50μm.

Since direct IN inoculation of Syrian hamsters is an artificial route of virus challenge, and Syrian hamsters transmit SARS-CoV-2 readily16, we repeated the above experiment within a direct contact horizontal transmission setting. Briefly, unvaccinated hamsters were IN challenged with SARS-CoV-2 (donor animals). After 24 hours, vaccinated animals were introduced into the cage, then four hours later, donor animals were removed (Figure 3A). As in the previous experiment, vaccination of hamsters with ChAdOx1 nCoV-19 resulted in high binding and neutralizing antibodies. Neutralizing antibodies were significantly higher in IN-vaccinated animals (Figure 3B–C). Control animals started losing weight at 4 days post exposure (DPE) and started recovering weight at 8 DPE. None of the vaccinated animals lost weight throughout the experiment, and a significant difference in weight was observed starting at 4 and 5 DPE for IN and IM-vaccinated animals compared to controls, respectively (Figure 3D). Ten animals per group were swabbed daily. Shedding of viral RNA, but not infectious virus, in controls was lower than in the previous experiment (multiple unpaired two-tailed Student’s t-test, 2–4 and 7 DPI, p-value <0.0001). A significantly reduced amount of shedding, both for viral RNA and infectious virus, was again detected in IN-vaccinated animals. However, as in the previous experiment, limited significant differences in shedding were detected in IM-vaccinated animals compared to controls (Figure 3E, mixed effects, p<0.05). The total amount of shedding, illustrated as AUC, was significantly different for IN-vaccinated animals compared to controls in both viral RNA and infectious virus (Kruskal-Wallis, p-value <0.0001), but not for IM-vaccinated animals (Figure 3F). Four animals per group were euthanized at 5 DPE and lung tissue was harvested. Again, no viral RNA or infectious virus was detected in lung tissue obtained from IN-vaccinated animals. However, viral RNA could be detected in lung tissue from three (gRNA) and two (sgRNA) IM-vaccinated animals, whereas infectious virus was detected in lung tissue of one IM animal (Figure 3G).

Figure 3. SARS-CoV-2 transmission to Syrian hamsters vaccinated with ChAdOx1 nCoV-19.

Figure 3.

Open in a new tab

Hamsters were vaccinated via the IN route (red), IM route (blue) or with control vaccine ChAdOx1 GFP via the IM route (green). A. Experimental schedule. Hamsters received a single vaccination 28 days prior to exposure. Donor animals were challenged at −1 DPE, and hamsters were co-housed for 4 hours. one day later. B. Binding antibody titers against SARS-CoV-2 S protein. C. Virus neutralizing antibody titers. D. Relative weight upon challenge with SARS-CoV-2. Shown is geometric mean and 95% confidence interval. # = p-value <0.05 between IN and control group; * = p-value <0.05 between vaccinated groups and control group. E. Viral load and viral titer in oropharyngeal swabs. Shown is geometric mean (symbols) and 95% confidence interval (shade). F. Area under the curve analysis of viral load and titer shedding in oropharyngeal swabs. G. Viral load and titer in lung tissue, isolated at 5 DPE. F-G. Dashed line = median; dotted line = quartiles. Statistical analyses done using mixed-effect analyses (C), two-way ANOVA (D), or Kruskal-Wallis test (E-F). * = p value <0.05; ** = p-value < 0.01; *** = p-value < 0.001.

Virus obtained from oropharyngeal swabs was sequenced at 2 and 5 DPE. Sequences obtained at 2 DPE from four different animals contained SNPs in the S protein. Two SNPs encoded a non-synonymous mutation; Asp839Glu and Lys1255Gln. Three swabs were obtained from IN-vaccinated animals, one swab was obtained from an IM-vaccinated animal (Table 1).

Table 1.

SNPs in SARS-CoV-2 sequences obtained from hamster swabs.

Nucleotide change Amino acid change Group Number of reads Mutation/total (%) Day
A23911T Ala783Ala IN 243/260 (93.5) 2 DPE
T24079G Asp839Glu IN 250/391 (63.9) 2 DPE
A24253C Pro897Pro IN 425/641 (66.3) 2 DPE
A25325C Lys1255Gln IM 273/768 (35.5) 2 DPE
Open in a new tab

Lung tissue of control animals obtained at 5 DPE had the same appearance as those obtained in the previous experiment. Lesions were observed in 40–50% of tissue, and interstitial pneumonia, edema, type II pneumocyte hyperplasia, and perivascular leukocyte infiltration were observed in all animals. As previously, no lesions or pathology were observed in lung tissue of IN-vaccinated animals. However, lesions were observed in the IM-vaccinated animals (5–20%, 3 out of 4 animals), accompanied with mild interstitial pneumonia (3 out of 4 animals), type II pneumocyte hyperplasia (2 out of 4 animals), and perivascular leukocyte infiltration (1 out of 4 animals). Edema was not observed (Figure 4 and Table S2). SARS-CoV-2 N antigen in lung tissue was found to be present in control animals (30–60% of lung tissue was immunoreactive) and to a lesser extent in IM-vaccinated animals (5% of lung tissue, 3 out of 4 animals), but not for IN-vaccinated animals (Figure 4 and Table S2).

Figure 4. Pulmonary effects of transmission of SARS-CoV-2 in Syrian hamsters.

Figure 4.

Open in a new tab

A-C. H&E; D-F. IHC. A. No pathology. B. Mild interstitial pneumonia. C. Moderate to marked interstitial pneumonia. D. No immunoreactivity. E. Scattered immunoreactive bronchiolar epithelial cells and Type I&II pneumocytes. F. Numerous immunoreactive bronchiolar epithelial cells and Type I&II pneumocytes. Bar = 50μm

The results obtained in hamster studies prompted us to investigate an IN vaccination in rhesus macaques17. Four non-human primates were vaccinated with a prime-boost regimen of ChAdOx1 nCoV-19 using the same dose as previously described8, utilizing an IN mucosal atomization device, which produced a spray of aerosols that were deposited in the nasal cavity. Four control animals were vaccinated with ChAdOx1 GFP. Blood, nasosorption swabs and bronchoalveolar lavage (BAL) samples were collected throughout the experiment. As expected, a higher fraction of IgA to total Ig was detected in nasosorption samples compared to BAL and serum samples (Figure S1). S and RBD-specific IgG antibodies were detected in serum and nasosorption samples after prime vaccination, but not in BAL, at seven days post prime vaccination (−49 DPI). Higher IgG titers were found in all samples obtained after a second vaccination at −28 DPI (Figure 5A–C). S and RBD-specific IgA antibodies were detected in serum upon prime vaccination but did not increase upon boost vaccination (Figure 5D). In contrast, SARS-CoV-2 specific IgA antibodies were only weakly detected in nasosorption samples upon prime vaccination but further increased upon boost vaccination (Figure 5E). No SARS-CoV-2 specific IgA antibodies were detected in BAL at −49 DPI but were detected seven days post boost vaccination (−21 DPI, Figure 5F). Circulating neutralizing antibodies were readily detected in vaccinated animals, to levels similar to convalescent serum obtained from human survivors (varying from asymptomatic to severe) and from NHPs which received a prime or prime-boost IM vaccinated with ChAdOx1 nCoV-198 (Kruskal-Wallis test, Figure 5G). Furthermore, multiple antigen-specific antibody Fc effector functions were detected; circulating antibodies in vaccinated animals promoted phagocytosis, complement deposition and NK cell activation (Figure 5H). Finally, levels of binding antibodies at 0 DPI against wildtype RBD and N501Y RBD were compared. N501Y is found in the RBD of two new variants of SARS-CoV-2: VOC2020–12/01 (B.1.1.7) and 501Y.V2 (B.1.351). No differences were found in the level of binding antibodies between wildtype and 501Y mutant RBD (Figure 5I).

Figure 5. Humoral response to IN vaccination with ChAdOx1 nCoV-19 in rhesus macaques.

Figure 5.

Open in a new tab

Truncated violin plot of SARS-CoV-2-specific IgG antibodies measured in serum (A), nasosorption samples (B), and BAL (C). Truncated violin plot of SARS-CoV-2-specific IgA antibodies measured in serum (D), nasosorption samples (E), and BAL (F). G. Truncated violin plot of neutralizing antibodies in serum. H. Effector functions of antibodies in serum. Black line = median; dotted line = quartiles; blue = vaccinated animals; purple = control animals (only 0 DPI values shown); black = human convalescent sera; red = NIBSC serum control 20/130. I. Anti-RBD-specific IgG titers at 0 DPI. Orange = RBD wildtype; aqua = RBD N501Y.

Animals were challenged via the intratracheal and IN route using 106 TCID50 of SARS-CoV-2 (SARS-CoV-2/human/USA/RML-7/2020). Nasal swabs were investigated for the presence of genomic RNA, subgenomic RNA and infectious virus. In control animals, both types of viral RNA were readily detected in nasal swabs. Genomic RNA was detected in all 4 animals (11 out of 16 swabs total), whereas subgenomic RNA was detected in 3 out of 4 animals (4 out of 16 swabs total). Infectious virus was detected in 3 out of 4 animals (5 out of 16 swabs total). Viral RNA was detected in nasal swabs obtained from vaccinated animals, but viral load was lower and fewer swabs were positive. Genomic RNA was detected in 3 out of 4 animals (5 out of 16 swabs total), whereas subgenomic RNA and infectious virus was only detected in 1 out of 4 animals (1 swab each) (Figure 6A). Total amount shed was depicted using AUC analysis. Although a downwards trend was observed in nasal swabs from vaccinated animals, this difference was not statistically significant (Mann-Whitney test, Figure 6B). Genomic and subgenomic RNA in BAL was detected in all four control animals (11 and 8 out of 12 samples, respectively). Infectious virus in BAL was detected in 2 out of 4 animals (3 out of 8 samples). Genomic RNA was detected in 4 out of 4 vaccinated animals, but only at early time points (5 out of 12 samples). Subgenomic RNA was only found in one animal and was very low (1 out of 12 samples). The differences in number of positive samples between vaccinated and control animals were significant using a Fisher’s exact test (genomic RNA p-value = 0.0272; subgenomic RNA p-value = 0.0094). No infectious virus could be detected in BAL samples from vaccinated animals (0 out of 12 samples) (Figure 6C). AUC analyses again showed a downwards trend in BAL from vaccinated animals, which was significant for sgRNA (Figure 6D, p=0.0286, Mann-Whitney test). Animals were euthanized at 7 DPI and viral RNA in nasal turbinates and lung tissue was analyzed. Viral load in lung was significantly lower for vaccinated animals than for control animals (Mann-Whitney test, p-value <0.0001 and 0.001 for genomic and subgenomic RNA, respectively), but no difference in viral load in nasal turbinates was detected (Figure 6E–F). No differences in hematology and radiographs were observed between groups.

Figure 6. SARS-CoV-2 detection in samples obtained from rhesus macaques upon virus challenge.

Figure 6.

Open in a new tab

gRNA, sgRNA and infectious virus in nasal swabs (A) and BAL (C) was determined. Dotted line = individual animals; solid line = geometric mean; shaded area = 95% confidence interval. Area under the curve (AUC) was calculated as an indication of the total amount of virus shed in nasal swabs (B) and BAL (D) and displayed as a truncated violin plot. Solid line = median; dotted line = quartiles. * = p-value <0.05 as determined via two-tailed Mann-Whitney test. Amount of gRNA and sgRNA in nasal turbinate (E) and lung tissue (F). Blue = vaccinated animals; purple = control animals; solid line = median; dotted line = quartiles. *** = p-value <0.001; **** = p-value <0.0001, as determined via two-tailed Mann-Whitney test.

We subsequently sought to define the impact of the vaccine-specific humoral response on nasal shedding and viral load after challenge. Principal component (PC) analysis of the pre-challenge, multivariate antibody profile revealed the distinct segregation of vaccinated animals from controls, driven by local and systemic antibodies with diverse functions (Figures 7A and 7B). Intergroup variation, largely encapsulated by PC2, was primarily mediated by differences in virus-specific IgA or IgG antibody levels in BAL and nasosorption samples. Notably, minimal levels of nasosorption IgG and relatively low levels of nasosorption IgA were detected in the only animal exhibiting significant nasal shedding after challenge (NHP1). This animal also had low serum IgG and virus neutralizing titers. Meanwhile, levels of BAL IgA and IgG were lowest in NHP2 and very high in NHP4; genomic and subgenomic RNA levels in BAL and lung tissue were highest and lowest in these animals, respectively. PC analysis of post-challenge viral load (AUC) in nasal swabs, BAL, and lung tissue, again, yielded dramatic clustering according to vaccination status (Figure 7C). Variation between control animals seemed to reflect site-specific differences in virus replication, between the upper and lower respiratory tract (Figure 7D).

Figure 7. Influence of the vaccine-induced humoral response on viral RNA levels post challenge.

Figure 7.

Open in a new tab

Principal component analysis (PCA) plot of the multivariate antibody (A) and AUC virology (C) profile across all animals (numbered dots). Ellipses indicate group distribution as 95% confidence levels. Mapped arrow projections indicate the influence of individual variables on the PCs; the antibody plot depicts only the top seven contributors. The complete antibody (B) and virology (D) variable loading plots for PCs 1 and 2 with a dotted line to indicate average expected contribution. Heatmap visualization of the correlations between antibody measures and viral RNA (AUC) levels (E) for the IN-vaccinated animals; R values were generated using two-sided Spearman rank correlation tests.

To examine these relationships further, we generated a correlation matrix integrating the pre- and post-challenge data from the IN-vaccinated animals (Figure S2). The Spearman rank correlation coefficients computed for individual antibody-virology variable pairings were assessed; however, the low number of animals precluded observations of statistical significance (Figure 7E). Nevertheless, clear trends emerged. While higher levels of serum (neutralizing and Fc effector-function-inducing) antibodies and nasosorption antibodies correlated with reduced nasal shedding, viral RNA in the BAL and lung tissue exclusively displayed strong negative correlations with BAL IgG and IgA levels. Of note, subgenomic RNA levels generally appeared to correlate more strongly with antibody levels than genomic RNA levels across sampling sites.

Discussion

Here we show that IN vaccination of hamsters and NHPs with ChAdOx1 nCoV-19 results in a robust mucosal and humoral immune response. In comparison to hamsters vaccinated via the IM route, a reduction in shedding is found in IN vaccinated animals, combined with full protection of the respiratory tract (no viral RNA). In NHPs, we observed a reduction in nasal shedding and viral load in BAL, as well as protection of the lower respiratory tract.

Since the release of the first full-length genome of SARS-CoV-218, thousands of complete genomes have been released. Multiple clades have been identified, as well as mutations throughout the genome of SARS-CoV-2. The most prevalent of the novel mutations is likely D614G in the S protein, which is present in the majority of circulating SARS-CoV-2 viruses19. All vaccines in clinical trials are based on the initial SARS-CoV-2 sequences18, and mutations in the S protein may result in immune evasion20. Here, a heterologous challenge was implemented in all experiments; we utilized isolate SARS-CoV-2/human/USA/RML-7/2020, which was isolated from a nasopharyngeal swab in July 2020 and belongs to clade 20A. This virus has one coding change in the S protein compared to the vaccine antigen; D614G. Both hamster and NHP studies described here demonstrate clearly that the ChAdOx1 nCoV-19 vaccine protects against SARS-CoV-2 containing the D614G mutation. It is likely that this translates to other vaccine platforms as well. Recently, new variants of SARS-CoV-2, named VOC2020–12/01 (B.1.1.7) and 501Y.V2 (B.1.351) were detected21 and VOC2020–12/01 (B.1.1.7) was potentially linked to increased transmission22. Both strains contain the N501Y mutation in RBD. We detected no difference in the ability of antibodies elicited by ChAdOx1 nCoV-19 vaccination to bind to N501Y mutant RBD.

Our previous and others’ studies investigating efficacy of COVID-19 vaccines in NHPs showed complete or near complete protection of the lower respiratory tract, but nasal shedding was still observed8–13. In natural infection with respiratory pathogens, a systemic immune response, dominated by IgG, as well as a mucosal immune response, dominated by secretory IgA (sIgA), is induced14,23. Although abundant literature exists on systemic immune responses to natural SARS-CoV-2 infection, literature on mucosal immunity is currently limited. In mucosal fluids from COVID-19 patients, S and RBD-specific IgA, IgG, and IgM were readily detected24–26. It is hypothesized that sIgA mainly protects the upper respiratory tract, whereas systemic IgG protects the lower respiratory tract14,27,28.

Upon IN vaccination of rhesus macaques with ChAdOx1 nCoV-19, we were able to detect SARS-CoV-2 specific IgG and IgA in serum. More importantly, SARS-CoV-2 specific IgG and IgA was also detected in nasosorption samples and BAL. No nasosorption samples were collected in our previous study8, but BAL collected at 3 and 5 DPI did not contain high levels of SARS-CoV-2 specific antibodies. Thus, IN vaccination resulted in systemic immunity comparable to that induced in vaccinated animals who received an IM vaccination with ChAdOx1 nCoV-19, but also elicited SARS-CoV-2-specific mucosal immunity as demonstrated by IgA detection in nasosorption and BAL samples.

Mucosal vaccination resulted in a reduction in shedding. In NHPs, subgenomic and infectious virus shedding was only detected in one vaccinated animal. This animal exhibited low levels of IgG and IgA antibodies in nasosorption samples coupled with low VN and sera IgG titers, suggesting that a robust humoral response in the nasal mucosa and in circulation is necessary to efficiently control nasal shedding.

Vaccination of small animal models with an adeno-vectored vaccine against SARS-CoV-2 has been reported by others, including two studies which investigated IN vaccination29–31. Bricker et al. showed a reduction in nasal shedding, near complete protection of upper respiratory tract and partial lower respiratory tract protection in hamsters29, whereas Hassan et al. did not investigate nasal shedding but found near complete protection of upper and lower respiratory tract tissue in mice30. Tostanoski et al. investigated IM vaccination in hamsters and found near complete protection in lung tissue dependent on vaccine candidate31. This agrees with our findings; we find a reduction in nasal shedding in IN-vaccinated animals, but not IM-vaccinated animals. We also find full protection of the lower respiratory tract in IN-vaccinated animals. Since IN vaccination of mice30 and NHPs elicited SARS-CoV-2-specific IgA in BAL and nasosorption samples (NHP only), we hypothesize that the same occurred in hamsters and combined with the higher neutralizing titers resulted in a reduction in nasal shedding.

In our second hamster study we moved away from IN inoculation and investigated vaccine efficacy in a transmission model. Transmission of SARS-CoV-2 was efficient, resulting in 100% transmission to control sentinel animals after just 4 hours of exposure to infected animals. Again, IN vaccination resulted in a reduction in shedding in sentinel hamsters compared to control animals. Although protection of the lower respiratory tract was complete in IN-vaccinated animals, only partial control was seen in IM-vaccinated animals, in contrast to the direct challenge experiment. It is possible that the difference between IN and IM-vaccinated animals is caused by virus seeding of the lungs from the URT; higher viral nasal shedding in IM-vaccinated animals compared to IN-vaccinated animals is likely reflective of a relative increase in virus deposition in the lung from the URT in IM compared to IN-vaccinated animals. That does not explain however why such a discrepancy between vaccination groups was not observed in the direct challenge study. Another hypothesis would be a difference in the initial site of virus deposition. Direct contact transmission likely represents a wide variety of exposure routes for the sentinel animals, including fomites and aerosols. A previous study in our laboratory showed the deposition of fluorescently labeled virus in the lungs of hamsters upon IN inoculation32. However, whereas that study used an inoculation volume of 80 μl, in the current study an inoculation volume of 40 μl was utilized, and it is possible that virus deposition directly into the lungs via IN inoculation with 40 μl is limited, whereas in direct transmission virus particles might be inhaled directly into the lung. Indeed, we recently showed that infection via aerosols, but not via direct IN inoculation, resulted in a high virus load in lung tissue at 1 DPI33. It should be noted that infectious virus titers in the lungs of control animals in the transmission experiment compared to the direct challenge experiment were ~5x higher, supporting this supposition. Mercado et al. previously showed the importance of different effector functions of antibodies in protection against SARS-CoV-2 in rhesus macaques13. We adapted their assays and show that upon IN vaccination with ChAdOx1 nCoV-19, a variety of antibody-dependent Fc effector functions are elicited, including monocyte cellular phagocytosis, complement deposition, and natural killer cell activation. Although the importance of neutralizing antibodies against SARSCoV-2 has been convincingly demonstrated in rhesus macaques34, the importance of other effector functions remains unknown. ChAdOx1 nCoV-19 has been shown to induce anti-S neutralizing antibody titers, as well as antibody-dependent neutrophil/monocyte phagocytosis, complement activation and natural killer cell activation35. A selective delay or defect in IgG development has been linked to severe and fatal outcome in human patients36. A recent study in mice demonstrated that in vitro neutralization did not uniformly correlate with in vivo protection, and that binding to Fc receptors was of importance, suggesting that antibody effector functions play a pivotal role in protection against SARS-CoV-237. Preliminary PC and correlation analyses suggested that while both vaccine-induced circulating antibodies—with neutralizing and non-neutralizing functionality—and upper respiratory antibodies play a role in reducing nasal shedding, virus replication in the airway and lung tissue is primarily controlled by antibodies localized to the lower respiratory tract. However, given that low animal numbers prevented us from establishing correlations of statistical significance, further studies will be required to more clearly define the relative impact of each component of the multifunctional humoral response on measures of protection.

The data presented supports the investigation of IN delivery of COVID-19 vaccines. With the roll-out of COVID-19 vaccines worldwide, it will be crucial to investigate whether the vaccines provide sterilizing immunity, or whether vaccinated people are still susceptible to infection of the URT and onward transmission of the virus. The data presented here demonstrates SARS-CoV-2-specific mucosal immunity is possible after IN vaccination, and results in a reduction in nasal shedding. It is now pertinent to investigate whether this finding translates to the clinic.

Supplementary Material

1

Acknowledgments:

We thank O. Abiona, B. Bailes, R. Cole, K. Corbett, K. Cordova, L. Crawford, H. Feldmann, B. S. Gallogly, Graham, L. Heaney, M. Jones, R. LaCasse, M. Marsh, K. Menk, A. Mora, Rose Perry, R. Rivera, L. Shupert, B. Smith, A. Weidow, and M. Woods for their assistance during this study.

Funding: This work was supported by the Intramural Research Program of the National Institute of Allergy and Infectious Diseases (NIAID), National Institutes of Health (NIH) (1ZIAAI001179-01) and the Department of Health and Social Care using UK Aid funding managed by the NIHR. Work in the Krammer laboratory was supported by the NIAID Centers of Excellence for Influenza Research and Surveillance (CEIRS) contract HHSN272201400008C and the Collaborative Influenza Vaccine Innovation Centers (CIVIC) contract 75N93019C00051.

Footnotes

Competing interests: S.C.G. is a board member of Vaccitech and named as an inventor on a patent covering the use of ChAdOx1-vector-based vaccines and a patent application covering a SARS-CoV-2 (nCoV-19) vaccine (UK patent application no. 2003670.3). T.L. is named as an inventor on a patent application covering a SARS-CoV-2 (nCoV-19) vaccine (UK patent application no. 2003670.3). The University of Oxford and Vaccitech, having joint rights in the vaccine, entered into a partnership with AstraZeneca in April 2020 for further development, large-scale manufacture and global supply of the vaccine. Equitable access to the vaccine is a key component of the partnership. Neither Oxford University nor Vaccitech will receive any royalties during the pandemic period or from any sales of the vaccine in developing countries. All other authors declare no competing interests. Mount Sinai has licensed SARS-CoV-2 serological assays to commercial entities and has filed for patent protection for serological assays as well as SARS-CoV-2 vaccines. FA and FK are listed as inventors on the pending patent applications.;

Data and materials availability: All data is available in the main text or the supplementary materials.

REFERENCES

  • 1.Voysey M. et al. Safety and efficacy of the ChAdOx1 nCoV-19 vaccine (AZD1222) against SARS-CoV-2: an interim analysis of four randomised controlled trials in Brazil, South Africa, and the UK. The Lancet S0140673620326611 (2020) doi: 10.1016/S0140-6736(20)32661-1 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Polack F. P. et al. Safety and Efficacy of the BNT162b2 mRNA Covid-19 Vaccine. N. Engl. J. Med. NEJMoa2034577 (2020) doi: 10.1056/NEJMoa2034577 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Baden L. R. et al. Efficacy and Safety of the mRNA-1273 SARS-CoV-2 Vaccine. N. Engl. J. Med. NEJMoa2035389 (2020) doi: 10.1056/NEJMoa2035389 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Cevik M. et al. SARS-CoV-2, SARS-CoV, and MERS-CoV viral load dynamics, duration of viral shedding, and infectiousness: a systematic review and meta-analysis. Lancet Microbe S2666524720301725 (2020) doi: 10.1016/S2666-5247(20)30172-5 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Bae S. H. et al. Asymptomatic Transmission of SARS-CoV-2 on Evacuation Flight. Emerg. Infect. Dis. 26, 2705–2708 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Cevik M., Kuppalli K., Kindrachuk J. & Peiris M. Virology, transmission, and pathogenesis of SARS-CoV-2. BMJ m3862 (2020) doi: 10.1136/bmj.m3862 [DOI] [PubMed] [Google Scholar]
  • 7.Madewell Z. J., Yang Y., Longini I. M., Halloran M. E. & Dean N. E. Household Transmission of SARS-CoV-2: A Systematic Review and Meta-analysis. JAMA Netw. Open 3, e2031756 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.van Doremalen N. et al. ChAdOx1 nCoV-19 vaccine prevents SARS-CoV-2 pneumonia in rhesus macaques. Nature 586, 578–582 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Corbett K. S. et al. Evaluation of the mRNA-1273 Vaccine against SARS-CoV-2 in Nonhuman Primates. N. Engl. J. Med. 383, 1544–1555 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Guebre-Xabier M. et al. NVX-CoV2373 vaccine protects cynomolgus macaque upper and lower airways against SARS-CoV-2 challenge. Vaccine 38, 7892–7896 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Wang H. et al. Development of an Inactivated Vaccine Candidate, BBIBP-CorV, with Potent Protection against SARS-CoV-2. Cell 182, 713–721.e9 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Gao Q. et al. Development of an inactivated vaccine candidate for SARS-CoV-2. Science 369, 77–81 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Mercado N. B. et al. Single-shot Ad26 vaccine protects against SARS-CoV-2 in rhesus macaques. Nature 586, 583–588 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Krammer F. SARS-CoV-2 vaccines in development. Nature 586, 516–527 (2020). [DOI] [PubMed] [Google Scholar]
  • 15.Rosenke K. et al. Defining the Syrian hamster as a highly susceptible preclinical model for SARS-CoV-2 infection. Emerg. Microbes Infect. 1–36 (2020) doi: 10.1080/22221751.2020.1858177 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Sia S. F. et al. Pathogenesis and transmission of SARS-CoV-2 in golden hamsters. Nature 583, 834–838 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Munster V. J. et al. Respiratory disease in rhesus macaques inoculated with SARS-CoV-2. Nature 585, 268–272 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Zhang Y.-Z. Novel 2019 coronavirus genome. https://virological.org/t/novel-2019-coronavirus-genome/319.
  • 19.Volz E. et al. Evaluating the Effects of SARS-CoV-2 Spike Mutation D614G on Transmissibility and Pathogenicity. Cell S0092867420315373 (2020) doi: 10.1016/j.cell.2020.11.020 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Starr T. N. et al. Prospective mapping of viral mutations that escape antibodies used to treat COVID-19. http://biorxiv.org/lookup/doi/10.1101/2020.11.30.405472 (2020) doi: 10.1101/2020.11.30.405472 [DOI] [PMC free article] [PubMed]
  • 21.Chand M. et al. Investigation of novel SARS-COV-2 variant of Concern 202012/01.
  • 22.Davies N. G. et al. Estimated transmissibility and severity of novel SARS-CoV-2 Variant of Concern 202012/01 in England. http://medrxiv.org/lookup/doi/10.1101/2020.12.24.20248822 (2020) doi: 10.1101/2020.12.24.20248822 [DOI]
  • 23.Russell M. W., Moldoveanu Z., Ogra P. L. & Mestecky J. Mucosal Immunity in COVID-19: A Neglected but Critical Aspect of SARS-CoV-2 Infection. Front. Immunol. 11, 611337 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Isho B. et al. Persistence of serum and saliva antibody responses to SARS-CoV-2 spike antigens in COVID-19 patients. Sci. Immunol. 5, eabe5511 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Cervia C. et al. Systemic and mucosal antibody secretion specific to SARS-CoV-2 during mild versus severe COVID-19. bioRxiv 2020.05.21.108308 (2020) doi: 10.1101/2020.05.21.108308 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Varadhachary A., Chatterjee D., Garza J., Garr R. P., Foley C., Letkeman A. F., Dean J., Haug D., Breeze J., Traylor R., Malek A., Nath R., & Linbeck L. Salivary anti-SARS-CoV-2 IgA as an accessible biomarker of mucosal immunity against COVID-19. medRxiv (2020) doi: 10.1101/2020.08.07.20170258 [DOI] [Google Scholar]
  • 27.Reynolds H. Y. Immunoglobulin G and Its Function in the Human Respiratory Tract. Mayo Clin. Proc. 63, 161–174 (1988). [DOI] [PubMed] [Google Scholar]
  • 28.Kubagawa H. et al. Analysis of paraprotein transport into the saliva by using anti-idiotype antibodies. J. Immunol. Baltim. Md 1950 138, 435–439 (1987). [PubMed] [Google Scholar]
  • 29.Bricker T. L. et al. A single intranasal or intramuscular immunization with chimpanzee adenovirus vectored SARS-CoV-2 vaccine protects against pneumonia in hamsters. http://biorxiv.org/lookup/doi/10.1101/2020.12.02.408823 (2020) doi: 10.1101/2020.12.02.408823 [DOI] [PMC free article] [PubMed]
  • 30.Hassan A. O. et al. A Single-Dose Intranasal ChAd Vaccine Protects Upper and Lower Respiratory Tracts against SARS-CoV-2. Cell 183, 169–184.e13 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Tostanoski L. H. et al. Ad26 vaccine protects against SARS-CoV-2 severe clinical disease in hamsters. Nat. Med. 26, 1694–1700 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.de Wit E. et al. Foodborne Transmission of Nipah Virus in Syrian Hamsters. PLoS Pathog. 10, e1004001 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Port J. R. et al. SARS-CoV-2 disease severity and transmission efficiency is increased for airborne but not fomite exposure in Syrian hamsters. http://biorxiv.org/lookup/doi/10.1101/2020.12.28.424565 (2020) doi: 10.1101/2020.12.28.424565 [DOI] [PMC free article] [PubMed]
  • 34.McMahan K. et al. Correlates of protection against SARS-CoV-2 in rhesus macaques. Nature (2020) doi: 10.1038/s41586-020-03041-6 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Barrett J. R. et al. Phase 1/2 trial of SARS-CoV-2 vaccine ChAdOx1 nCoV-19 with a booster dose induces multifunctional antibody responses. Nat. Med. (2020) doi: 10.1038/s41591-020-01179-4 [DOI] [PubMed] [Google Scholar]
  • 36.Zohar T. et al. Compromised Humoral Functional Evolution Tracks with SARS-CoV-2 Mortality. Cell 183, 1508–1519.e12 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Schäfer A. et al. Antibody potency, effector function, and combinations in protection and therapy for SARS-CoV-2 infection in vivo. J. Exp. Med. 218, e20201993 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Dicks M. D. J. et al. A Novel Chimpanzee Adenovirus Vector with Low Human Seroprevalence: Improved Systems for Vector Derivation and Comparative Immunogenicity. PLoS ONE 7, e40385 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Bewig B. & Schmidt W. E. Accelerated Titering of Adenoviruses. BioTechniques 28, 870–873 (2000). [DOI] [PubMed] [Google Scholar]
  • 40.Maizel J. V., White D. O. & Scharff M. D. The polypeptides of adenovirus. Virology 36, 115–125 (1968). [DOI] [PubMed] [Google Scholar]
  • 41.Thwaites R. S. et al. Nasosorption as a Minimally Invasive Sampling Procedure: Mucosal Viral Load and Inflammation in Primary RSV Bronchiolitis. J. Infect. Dis. 215, 1240–1244 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.Singletary M. L. et al. Modification of a common BAL technique to enhance sample diagnostic value. J. Am. Assoc. Lab. Anim. Sci. JAALAS 47, 47–51 (2008). [PMC free article] [PubMed] [Google Scholar]
  • 43.Corman V. M. et al. Detection of 2019 novel coronavirus (2019-nCoV) by real-time RT-PCR. Eurosurveillance 25, (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Wölfel R. et al. Virological assessment of hospitalized patients with COVID-2019. Nature 581, 465–469 (2020). [DOI] [PubMed] [Google Scholar]
  • 45.Avanzato V. A. et al. Case Study: Prolonged Infectious SARS-CoV-2 Shedding from an Asymptomatic Immunocompromised Individual with Cancer. Cell S0092867420314562 (2020) doi: 10.1016/j.cell.2020.10.049 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Stadlbauer D. et al. SARS‐CoV‐2 Seroconversion in Humans: A Detailed Protocol for a Serological Assay, Antigen Production, and Test Setup. Curr. Protoc. Microbiol. 57, (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Wrapp D. et al. Cryo-EM structure of the 2019-nCoV spike in the prefusion conformation. Science 367, 1260–1263 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Amanat F. et al. A serological assay to detect SARS-CoV-2 seroconversion in humans. Nat. Med. 26, 1033–1036 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Briese T. et al. Virome Capture Sequencing Enables Sensitive Viral Diagnosis and Comprehensive Virome Analysis. mBio 6, e01491–15 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50.Martin M. Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet.journal 17, 10 (2011). [Google Scholar]
  • 51.Langmead B. & Salzberg S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357–359 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.McKenna A. et al. The Genome Analysis Toolkit: A MapReduce framework for analyzing next-generation DNA sequencing data. Genome Res. 20, 1297–1303 (2010). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53.Li H. A statistical framework for SNP calling, mutation discovery, association mapping and population genetical parameter estimation from sequencing data. Bioinformatics 27, 2987–2993 (2011). [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

1
NIHPP2021.01.09.426058-supplement-1.pdf (641.4KB, pdf)

Articles from bioRxiv are provided here courtesy of Cold Spring Harbor Laboratory Preprints

RESOURCES