Cutting Edge: Nucleocapsid Vaccine Elicits Spike-Independent SARS-CoV-2 Protective Immunity Free

Vaccine candidates targeting SARS-CoV-2 S protein, which is essential for cell entry, were designed based on viral sequences reported in January 2020. Studies of the highly efficacious Pfizer/BioNTech and Moderna mRNA vaccines show a strong correlation between the establishment of neutralizing Abs (NAb) and protection from disease (1, 2). As the virus continues to spread globally, variants have emerged that partly evade neutralization by vaccine-elicited S Abs (3), which may require continued vaccine adaptation and boosting.

T cells contribute to immunity against respiratory pathogens, including serological variants of influenza virus (4). T cells may target conserved epitopes that have an unfavorable fitness barrier to mutation (5). Unlike NAb, T cell immunity is not limited to surface Ags and immunodominant epitopes vary considerably between individuals because of recognition in the context of genetically diverse MHC molecules. Accordingly, T cell immunity may be less vulnerable to immune selection pressure and viral escape mutations. We immunized mice and hamsters against SARS-CoV-2-N to address whether protection could be conferred independently of spike-specific responses.

Male Syrian hamsters (Charles River Laboratories) were maintained in a BSL-2 containment facility under specific pathogen–free conditions at the University of Minnesota in accordance with the Institutional Animal Care and Use Committees guidelines.

Eighteen- to 19-wk-old hamsters were anesthetized with isoflurane and immunized using a single i.v. dose of 1.8 × 10¹¹ viral particles of either a replication-deficient (E1- and E3-deleted) human adenovirus type 5 (Ad5) vectors expressing SARS- CoV-2-N (Ad5-N) or a control Ad-5 vector (Ad5-NULL). Ad5 vectors were constructed as described (6). Hamsters were moved to BSL-3 containment for SARS-CoV-2 challenge studies, anesthetized with isoflurane and challenged intranasally (i.n.) with a total of 6.75 × 10⁵ PFU in a 100 ul volume of either the 2019-nCoV/USA_WA1/2020 (WA) strain or the B.1.1.7 variant SARS-CoV- 2/human/USA/CA_CDC_5574/2020 (provided by World Reference Center for Emerging Viruses and Arboviruses at the University of Texas Medical Branch). Animals were weighed daily and terminated 13 d after the challenge. Hamsters challenged for determining viral titers were euthanized 3 d after challenge.

Six- to 10-wk-old female C57BL/6 or hemizygous K18.hACE2 (K18) mice (stock no. 034860, Jackson Laboratory) were maintained in specific pathogen–free conditions at the University of Minnesota in accordance with the Institutional Animal Care and Use Committees guidelines (7). Littermates were immunized i.v. with 5 × 10¹⁰ viral particles of Ad5-N vector or Ad5-NULL.

Immunized K18 mice were challenged with SARS-CoV-2 in the BSL-3 containment facility. For depletion experiments, mice received i.p. injections of 300 μg of both CD4 (GK1.5) and CD8b (Lyt 3.2) or isotype controls (all from InVivoMab) on days 2, 3, 5, and 6 before challenge. Animals were anesthetized with ketamine/xylazine, laid supine, and challenged i.n. with 300 PFU of WA SARS-CoV-2 in a 30 μl volume. Infected animals were monitored daily for health, weight loss, and morbidity and were deemed terminal if they reached 75% of starting weight. All weight loss experiments were terminated 13 d postinfection. Mice assigned for flow cytometry analysis and the concurrent determination of viral titers were euthanized 4 d after challenge.

To discriminate extravascular cells from intravascular cells, mice were injected i.v. with BV605-conjugated anti-CD8a Ab, sacrificed after 3 min, and tissues were harvested (8). Mouse lungs were mechanically minced into small pieces and further dissociated in PBS using gentleMACS tubes. For SARS-CoV-2 challenge studies, dissociated tissue was split equally into two parts, one for quantification of SARS-CoV-2 viral titers and the other for isolation of lymphocytes for flow cytometric analyses. Isolation of cells for flow cytometric analysis was performed as previously described (9). Cells were stained with Abs to CD8a (53-6.7), CD62L (MEL-14), Ly6C (HK1.4), CD44 (IM7), CD69 (1H.2F3), and CD103 (M290) (all from BD Biosciences, Tonbo Biosciences, BioLegend, or Affymetrix eBiosciences) and H2-D^b/N_219–227 MHC class I tetramers (made in-house) and ghost dye 780 (Tonbo Biosciences). For monomer preparation, the N_219–227 peptide sequence was LALLLLDRL. Stained samples were acquired on LSRII or LSR Fortessa flow cytometers (BD Biosciences) and analyzed with FlowJo software (BD Biosciences). Cells were gated on singlets, live lymphocytes, CD8 T cells, and/or tetramer-specific cells as indicated.

Hamsters and mice were euthanized 3 and 4 d after SARS-CoV-2 infection, respectively. Lungs were harvested from hamsters, weighed, and homogenized in PBS using gentleMACS M tubes and a gentleMACS dissociator on the protein setting. Mouse lungs were weighed and processed as described above before being further homogenized in PBS using gentleMACS M tubes (9). Tubes were centrifuged to pellet debris. The supernatants were collected, aliquoted, and stored at –80°C.

Vero E6 cells (American Type Culture Collection) were maintained in DMEM with 10% FBS and 1% penicillin-streptomycin. Infections were carried out in an infection medium (DMEM with 1% penicillin-streptomycin, 1% nonessential amino acids, 10mM HEPES, and 2% FBS). The plaque assay was adapted from Mendoza et al. (10). Ten-fold dilutions of the homogenized lung samples were adsorbed to Vero E6 cells. The cells were incubated for 1 h at 37°C in a 5% CO₂ incubator, rocking every 10 min. Cells were washed with PBS and overlaid with a carboxymethylcellulose (CMC) overlay media (MEM, 1.5% CMC) and incubated at 37°C in a 5% CO₂ incubator for 3 d. The overlay media was removed, cells were washed with PBS, and fixed for 1 h with 4% paraformaldehyde at room temperature. The fixative was discarded, and the cells were stained with crystal violet (0.5%) for 5–15 min. Cells were washed with distilled water 2–4 times, blotted dry, and plaques were enumerated.

Outbred Syrian hamsters are permissive to SARS-CoV-2 infection. We vaccinated hamsters i.v. with Ad5-N, which expresses the N sequence derived from WA, or a control Ad5 vector lacking an SARS-CoV2 sequence, Ad5-NULL. Seven- to eight-week later, animals were challenged i.n. with 6.75 × 10⁴ PFU WA SARS-CoV-2. Vaccination reduced weight loss (Fig. 1A, 1B). Vaccinated hamsters were also challenged with WA or a variant strain B.1.1.7, which contains two amino acid substitutions in N that do not occur within the N_219–227 immunodominant epitope that we previously defined in C57BL/6 mice (6). Vaccination elicited a significant 30-fold (WA) and 12-fold (B.1.1.7) reduction in median lung viral titer 3 days following challenge, although the latter was not significant. We next immunized transgenic K18 mice that are highly susceptible to SARS-CoV-2 (11). Thirty days following i.v. vaccination with Ad5-N or Ad5-NULL, K18 mice were challenged with 300 PFU SARS-CoV-2. Ad5-N–vaccinated mice experienced significantly reduced weight loss (Fig. 1D, 1E) and mortality, with 75% surviving challenge versus 0% in the Ad5-NULL–vaccinated group (Fig. 1F).

Vaccination established SARS-CoV-2-N_219–227–specific memory CD8 T cells that persisted in lung, airways, and spleen from 40 to 86 d, and increased in the lung draining mediastinal lymph node (Fig. 2A). We examined whether T cells in vaccinated mice participated in the response 4 d after WA SARS-CoV-2 challenge using a combined Ab-mediated CD8b and CD4 depletion strategy (Fig 2B, 2C). Vaccinated mice showed a ∼3.8-log 10 reduction in lung viral load (Fig. 2D). Combined CD8b and CD4 depletion prior to challenge partially abrogated protection, although T cell depletion was not absolute, Fig. 2B) and residual N_219–227–specific CD8 T cells were still observed in vaccinated mice (Fig. 2C).

i.n. challenge increased granzyme B⁺ N_219–227–specific CD8 T cells in the pulmonary mucosa (Fig. 2E). Total N_219–227–specific CD8 T cells substantially increased in both lung parenchyma [defined by the absence of intravascular anti-CD8a staining (IVneg), as previously described (8)] and draining mediastinal lymph node, moderately decreased in spleen, and substantially decreased in the lung vasculature [defined by the presence of intravascular anti-CD8a staining (IVpos)] (Fig. 2F). These data indicate that vaccine-elicited memory CD8 T cells underwent rapid reactivation and migration to the site of viral challenge, and that T cells may have contributed to viral control.

The rapidity and success of SARS-CoV-2 vaccine development and deployment is unprecedented. Most strategies, including all vaccines licensed for emergency use in the U.S.A., immunize only against the viral spike. Spike is a logical choice because it contains the receptor binding domain that is the main target of NAb. Nevertheless, SARS-CoV-2 is likely to become endemic. Viral variants have emerged that reduce vaccine-elicited NAb efficacy (12), suggesting partial escape, and SARS-CoV-2 may continue to evolve with or without selection pressure from increased global immunity. Whereas vaccination strategies often rely on NAbs for efficacy, engaging other arms of the immune system, such as cellular immune memory, have been shown to offer protection against viral infections and/or reduce the threshold of NAbs needed for protection (13, 14). This study provides evidence in rodents that immunological memory to additional Ags could provide protection, which may involve memory T cells and other nonneutralizing effector mechanisms. It appears likely that viral evolution will necessitate vaccine evolution and booster immunizations that address emergent variants. This study supports the rationale for including additional viral Ags into future vaccine candidates to broaden epitope diversity and protection mechanisms while limiting opportunities for viral escape.

We thank the Biosafety Level 3 Program at the University of Minnesota. SARS-CoV-2 viruses were obtained through the World Reference Center for Emerging Viruses and Arboviruses at the University of Texas Medical Branch (Galveston, TX).

The authors have no financial conflicts of interest.