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. 2025 Aug 4;11(4):00940-2024. doi: 10.1183/23120541.00940-2024

SARS-CoV-2 spike treatment and transfection impairs airway epithelial repair

Tony Guo 1,✉, Gurpreet Singhera 1,2, Jasmine Memar Vaghri 1, Wan Yi Liang 1, Janice M Leung 1,2, Del Dorscheid 1,2
PMCID: PMC12320110  PMID: 40761650

Abstract

Background

The airway epithelium serves as a physical and immune barrier against inhaled insults. This tissue is susceptible to severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) infection and, following injury, the airway epithelium undergoes repair to restore barrier function. Although components of SARS-CoV-2, such as the spike glycoprotein essential in viral entry, have been shown to alter biological functions in various tissues, it is unclear how SARS-CoV-2 can impact airway epithelial functions, such as wound repair.

Methods

In this study, 16HBE14o- epithelial monolayer cultures were either treated with recombinant SARS-CoV-2 spike glycoprotein S1 subunit at 4 μg·mL−1 or transfected with a plasmid expressing full-length spike glycoprotein. Secreted inflammatory mediators, markers of proliferation and cell cycle arrest, culture proliferation, and wound closure measurements following mechanical injury were assessed.

Results

Spike treatment and transfection altered measures of culture proliferation and markers of proliferation and cell cycle arrest. Secreted interleukin-6 but not interleukin-8 were significantly higher with spike S1 treatment, while both were significantly elevated with spike transfection. Wound closure was inhibited by both spike treatment and transfection, with significant reductions compared to control.

Conclusions

SARS-CoV-2 spike S1 treatment and transfection can alter measures of proliferation and inflammation as well as impair wound closure of 16HBE14o- airway epithelial cells. These results highlight how components of SARS-CoV-2 can impair functions of the airway epithelium independent of viral replication.

Shareable abstract

SARS-CoV-2 spike glycoprotein can hinder cellular proliferation, trigger inflammation and disrupt repair in airway epithelial cells in vitro, potentially contributing to airway damage and dysfunction in COVID-19, even in the absence of active infection https://bit.ly/3Cr8H2C

Introduction

The airway epithelium, which lines the respiratory tract, acts as both a physical and immune barrier. It plays a crucial role in immune responses related to severe COVID-19, which may result from dysregulated repair and reduced barrier function [1]. Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), the agent responsible for COVID-19, can infect the airway epithelium [2]. SARS-CoV-2 enters susceptible host cells by binding to the cell receptor angiotensin-converting enzyme 2 (ACE2) with its spike glycoprotein, located on the surface of the viral membrane and comprises S1 and S2 subunits [3]. Binding of the virus to ACE2 triggers conformational changes in the S1 subunit, exposing a cleavage site [4]. The proteolytic cleavage of the viral spike protein is mediated by transmembrane serine protease 2 (TMPRSS2) on the cell surface or by cathepsin B and L in the endosomes if the virus has been endocytosed [5]. This primes the virus for membrane fusion, which is mediated by the S2 subunit. Furin, a protease localised to the Golgi, cleaves the spike protein, which leads to enhanced infectivity of the virus [6]. ACE2, TMPRSS2 and other host viral entry factors determine the viral tropism of SARS-CoV-2 [7].

Infection by viruses, such as SARS-CoV-2, can cause cell damage, death and shedding. Cell death by SARS-CoV-2 can occur through apoptosis, necroptosis and promotion of autophagy resulting in cytopathic effects and disruption of epithelial integrity [8–11]. This damage requires airway epithelial repair to restore barrier function and defence [9]. This repair process is usually coordinated, involving cell spreading at the margins to close the wound, cell migration and proliferation to recover lost cell populations, and differentiation to restore normal cellular function [12]. However, recent observations suggest that the SARS-CoV-2 spike protein, which is responsible for receptor binding and entry into susceptible cells, can trigger cell signalling to alter various biological processes independent of viral infection [13].

In this study, we used two models to investigate SARS-CoV-2 interaction, namely spike S1 subunit treatment and transient expression of full-length spike protein through transfection. These approaches allowed us to avoid the use of live authentic virus due to its pathogenicity. The spike S1 subunit, which contains the receptor binding domain [14], was used to mimic the binding mechanisms observed between SARS-CoV-2 and the airway epithelium, replicating events following virion entry. Transfection, on the other hand, simulated the viral replication phase, focusing on the synthesis of virus components prior to virion assembly. Both methods aim to characterise interactions without considering active viral replication. To examine these interactions, we measured markers of cellular proliferation, cell cycle regulation and inflammatory mediators, such as interleukin (IL)-6 and IL-8, which play key roles in wound repair, proliferation and migration [15–18]. Furthermore, these two cytokines have been implicated in COVID-19 pathogenesis [1, 19–21]. Additionally, wound repair assays were used to characterise the interaction between the spike protein and the airway epithelium. The goal of this project is to investigate whether the SARS-CoV-2 spike protein alters the repair of the wounded airway epithelium. Exploring how this important component of SARS-CoV-2 interacts with the epithelium may improve our understanding of COVID-19 pathogenesis within the lungs.

Material and methods

More detailed methods can be found in the supplementary materials.

Cell culture

16HBE14o- (16HBE) airway epithelial cells were a generous gift from Dr. Dieter Gruenert, University of California, San Francisco. Cells were grown in complete DMEM, containing 10% fetal bovine serum, 10 U·mL−1 of penicillin and 10 μg·mL−1 streptomycin. Recombinant spike S1 protein (Sino Biological, Beijing, China) was diluted to a final concentration of 4 μg·mL−1 in complete DMEM before cell treatment and incubation at 37°C for the specified time periods. This concentration was chosen as an average of values reported in the literature, which ranged from 0.8 to 10 μg·mL−1 in studies assessing exposure effects on airway epithelial cells, Vero E6 cells and vascular endothelial cells [22–26]. 16HBE cells were seeded and transfected with pCMV14-3X-Flag-SARS-CoV-2 S plasmid using Lipofectamine 3000 (Thermo Fisher Scientific), according to the manufacturer's instructions. An empty pCMV14-3X-FLAG backbone plasmid vector was used as the negative control.

Immunofluorescence

Methanol-fixed and permeabilised 16HBE cultures were probed with rabbit polyclonal anti-ACE2 antibody (ab15348, 1:500 dilution, Abcam, Cambridge, UK) and mouse monoclonal anti-β-tubulin antibody (T8328, 1:5000 dilution, Sigma-Aldrich, St. Louis, MO) overnight at 4°C. The next day, goat anti-rabbit IgG (H+L) conjugated with Alexa Fluor 488 (A28175, 1:200, Thermo Fisher Scientific) and goat anti-mouse IgG (H+L) conjugated with Alexa Fluor 594 (A11032, 1:200, Thermo Fisher Scientific) were used as secondary antibodies, followed by Hoechst 33342 (Thermo Fisher Scientific) for nuclear counterstain. Slides were imaged using the Zeiss LSM880 inverted confocal microscope (Baden-Württemberg, Germany) at ×100 magnification.

To examine the transient expression of spike protein following transfection, spike-transfected 16HBE cells were fixed, permeabilised and blocked at 48 h post-transfection as described above. Mouse monoclonal anti-FLAG tag antibody (F3165, 1:1000, Sigma-Aldrich) was added and incubated with the cultures overnight at 4°C. Goat anti-rabbit IgG (H+L) antibody, Alexa Fluor 488 conjugated (A28175, 1:200), was added and nuclei were counterstained with 4′,6-diamidino-2-phenylindole (DAPI). Slides were mounted and imaging was performed as described above.

Western blotting

Lysates of spike S1-treated, spike-transfected and control 16HBE were separated using a 4–20% SDS polyacrylamide gel electrophoresis gradient gel. Following transfer, the blots were blocked using 5% skim milk in TBS with 0.1% Tween 20. The spike protein was detected using mouse monoclonal anti-FLAG tag antibody (F3165, 1:1000). Blots were also probed for Ki67 and p21Cip1. Ki67, which is expressed predominantly during the cell cycle synthesis phase [27], serves as the marker for proliferation, and was detected with anti-Ki67 rabbit polyclonal antibody (ab16667, Abcam, 1:500). p21Cip1 is a cyclin-dependent kinase inhibitor responsive to genotoxic stress [28] and serves as a marker of cell cycle arrest. It was detected with anti-p21Cip1 rabbit polyclonal antibody (A1483, Abclonal, 1:1000). To assess baseline host viral entry factor expression, the following antibodies were used: anti-ACE2 (ab15348, 1:1000, Abcam), anti-TMPRSS2 (ab109131, 1:1000, Abcam) and anti-furin (ab3467, 1:1000, Abcam). Secondary anti-rabbit horseradish peroxidase (HRP, AS014, 1:2000, Abclonal Technologies) was incubated with the blots for 1 h at room temperature. β-Actin was probed as a loading control (sc-47778, 1:2000, Santa Cruz, Dallas, TX). Protein bands were detected using enhanced chemiluminescence.

Proliferation assay

16HBE cells were seeded at a 20% confluency and 16 h post-seeding, the cells were treated with spike S1 at 4 μg·mL−1 diluted in complete DMEM or transfected with spike-expressing plasmid as outlined above. The cultures were imaged at 0, 24, 48, 96 and 120 h post-treatment or post-transfection. Captured images (each with eight independent replicates) were counted by two blinded observers for further analysis.

Flow cytometry for cell cycle analysis

16HBE cells were seeded onto 24-well cell culture plates at a density of 7.5×104 cells per well. 16 h post-seeding, these cells were treated with spike S1 at 4 μg·mL−1 diluted in complete DMEM or transfected with spike-expressing plasmid as outlined above. Cells were collected at 24 or 48 h post-treatment via trypsin detachment, washed with PBS, resuspended into 300 μL of 70% ethanol for fixation and permeabilisation, and stored at 4°C for at least 24 h. Subsequently, cells were washed with PBS, resuspended in 2 μg·mL−1 RNase (EN0531, Thermo Fisher Scientific) diluted in distilled water for 30 min at room temperature, stained with propidium iodide (PI) (#P4170, Sigma-Aldrich) diluted in PBS at a final concentration of 50 μg·mL−1 and incubated in the dark for 30 min at room temperature. Cells were then suspended into 300 μL PBS and PI fluorescence was detected at 488 nm using Gallios flow cytometer (Beckman Coulter, Brea, CA). Approximately 6000 cells exhibiting PI fluorescence were analysed per sample. Gating was optimised using Kaluza Analysis (Beckman Coulter) to define the cell population and cell cycle phases were assessed using the Floreada analysis tool (https://floreada.io).

Wound repair assay and quantification of wound closure

Spike-treated or transfected 16HBE cultures were mechanically injured by generating crosshatch wounds and gently washed with PBS to wash off any detached cells. Crosshatch wounds were imaged through light microscopy using the EVOS M5000 imaging system (Thermo Fisher Scientific) at 0, 3, 12 and 24 h post-wounding. The percentage of wound closure was calculated as the change in wound area between a specified time-point and the 0 h time-point, to normalise to the original wound size. This analysis was performed by two blinded observers using the ImageJ image analysis software (National Institutes of Health).

Immunoassays on inflammatory markers

Supernatants were collected at 24 and 48 h post-treatment or transfection of 16HBE cultures, respectively, centrifuged at 500 g for 10 min and stored at −80°C until use. These supernatants were thawed on ice and the expression of IL-6 and IL-8 were quantified using ELISA kits, catalogue numbers CHC1263 and CHC1303, respectively, from Thermo Fisher Scientific.

Statistical analysis

Data were represented as means±standard error of the mean (sem). Data were analysed using Prism statistical analysis software (version 9.3.1, GraphPad, La Jolla, CA). One-way ANOVA with Tukey's multiple comparison test was used unless specified otherwise.

Results

16HBE cells express ACE2, TMPRSS2 and furin – host viral entry receptors for SARS-CoV-2 entry

To ascertain the suitability of the 16HBE cell line for modelling the interaction between SARS-CoV-2 and the airway epithelium, we characterised the expression of ACE2, TMPRSS2 and furin. ACE2 immunofluorescence showed punctate and globular staining within the cytoplasm, which was co-localised with the cytoskeletal signal of β-tubulin. Diffuse ACE2 signal was also observed within the nuclei of 16HBE monolayers (figure 1a). Evidence of ACE2 expression was also seen through Western blotting, where it is resolved as a 120 kDa band, corresponding to its mature, glycosylated form (figure 1b). Furthermore, TMPRSS2 and furin were also expressed in 16HBE, resolved as 90 and 55 kDa bands, respectively. Given the observation that 16HBE airway epithelial cells express host cell viral entry factors, these characteristics render 16HBE a suitable cell line for modelling the interaction with SARS-CoV-2.

FIGURE 1.

FIGURE 1

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16HBE airway epithelial cell line expresses angiotensin-converting enzyme 2 (ACE2), transmembrane serine protease 2 (TMPRSS2) and furin. a) 16HBE monolayers were fixed, permeabilised and probed against ACE2, while the cytoskeleton was labelled using a mouse monoclonal anti-β-tubulin antibody for the immunofluorescence assay. Nuclei were counterstained and the samples were imaged using confocal microscopy. ACE2 expression (green) in 16HBE monolayers was observed as punctate and globular staining within the cytoplasm (β-tubulin, red) and diffuse staining within the nuclei (4′,6-diamidino-2-phenylindole (DAPI), blue). Scale bar: 25 μm. b) Western blot analysis of ACE2, TMPRSS2 and furin in 16HBE monolayers indicated the presence of all three host viral entry factors. Lanes represent two biological replicates.

Spike S1 treatment and spike transfection alter the expression of markers associated with senescence and proliferation

For transient expression of the SARS-CoV-2 spike protein intracellularly, 16HBE monolayers were transfected with pCMV14-3X-Flag-SARS-CoV-2 S plasmid. Spike protein expression was validated via probing against the attached artificial tag, 3X FLAG peptide, via Western blotting (figure 2a) and immunofluorescence (figure 2b).

FIGURE 2.

FIGURE 2

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Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) spike expressed in 16HBE cultures transfected with pCMV14-3XFLAG-SARS-CoV-2 S. a) Cell lysates of 16HBE cultures transfected with spike-expressing plasmid (lane I) or transfected with an empty backbone plasmid vector (lane II) analysed via Western blotting against the anti-FLAG tag. The spike transfection in lane I reveals three bands, as follows: 1) at ∼360 kDa corresponding to the dimeric spike protein, 2) at ∼180 kDa corresponding to the full-length monomeric spike protein and 3) at ∼100 kDa corresponding to the cleaved spike protein, likely the S2 subunit. b) Spike protein expression in spike-transfected 16HBE monolayers imaged through immunofluorescence staining. Cultures were fixed, permeabilised and probed for the FLAG tag, nuclear counterstained and imaged using confocal microscopy. Scale bar: 50 μm.

Western blotting revealed three bands corresponding to dimeric, full-length and cleaved spike protein. No signal was observed with transfection of the empty plasmid. Immunofluorescence revealed that around ∼20% of cells were expressing spike protein 48 h post-transfection.

After validation, we further examined the impact of spike extracellular treatment or spike transfection on key cellular repair processes.

With spike S1 treatment, we observed a significant reduction in Ki67 expression at all time-points tested (figure 3a and b) and an overall corresponding increase in p21Cip1 expression, compared to control, which was significant at all time-points except at 48 h (figure 3a and c). Whereas the reduced expression was maintained in all time-points for Ki67, p21Cip1 expression increased and peaked before reducing to a level that was insignificant compared to control at 48 h post-treatment. Moreover, to characterise whether proliferation is functionally impaired with spike treatment, treated cultures were monitored for growth over time (figure 3d). Significant differences in cell counts were observed only at 120 h post-treatment. Furthermore, the secretion of pro-inflammatory cytokine, IL-6 and chemokine, IL-8, was quantified using ELISA (figure 3e). While the increase in IL-8 secretion under spike treatment was not significant, a significant elevation was observed in IL-6 secretion. DNA cell cycle analysis shows a significant decrease in the proportion of cells in G1 (42.1±0.7% versus 38.5±0.2%, p=0.02), a significant increase with the S phase population (27.8±0.9% versus 32.1±0.6%, p=0.007) and no significant difference in G2 (29.4±0.6% versus 30.0±0.2%, p>0.99) with spike S1 treatment compared to control (figure 3f). Representative histograms can be found in supplementary figure 1A. Activation of caspase-3, a key terminal death protease [29], was also assessed via Western blotting with no cleaved, activated caspase-3 visualised (supplementary figure 2A).

FIGURE 3.

FIGURE 3

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Spike S1 treatment alters Ki67 and p21Cip1 protein expression, proliferation efficiency and inflammatory mediator release. a) Representative images of S1-treated 16HBE monolayer cultures lysed at various time-points. Protein expression was characterised using Western blotting against Ki67 and p21Cip1. b) Densitometry analysis of Ki67 expression normalised to β-actin with one-way ANOVA and Dunnett's post-test used for statistical analysis. c) Densitometry analysis of p21Cip1 expression normalised to β-actin with one-way ANOVA and Dunnett's post-test used for statistical analysis. d) The proliferation efficiency of the culture was assessed by cell counting and is summarised. Two-way ANOVA with Šídák's multiple comparisons test was used for statistical testing. e) Release of interleukin (IL)-6 and IL-8 from S1-treated or control 16HBE cultures were quantified using ELISA. An unpaired t-test was used. f) DNA cell cycle analysis with propidium iodide staining was performed. 6000 cells gated as positive were analysed and summarised here with the cell population segregated based on the phase of the cell cycle. Two-way ANOVA with Šídák's multiple comparisons test was used. #: The proportion of cells in the G1 phase was lower (p=0.02), while that in the S phase was higher (p=0.007) following S1 treatment compared to the control. For all experiments, only pairwise comparisons p<0.05 are shown. *: p<0.05. **: p<0.01. ***: p<0.001.

With spike-expressing plasmid transfection, we observed that spike expression increased and peaked at 48 h post-transfection before decreasing at 72 h, exemplifying the transiency of its expression (figure 4a). At the time-points tested, whereas no significant change in normalised p21Cip1 expression was seen with spike transfection (figure 4c), normalised Ki67 expression was significantly reduced (p<0.0001) compared to control (figure 4b). This decrease in Ki67 expression was found to be reciprocal to the increase in spike expression, with the trough of Ki67 expression and peak of spike expression occurring simultaneously at 48 h post-transfection (figure 4b). A proliferation assay demonstrated significantly lower cell counts at all time-points post-transfection compared to control (figure 4d). Moreover, IL-6 and IL-8 secretion were significantly greater at 48 h post-transfection compared to control (figure 4e). DNA analysis shows no significant difference in G1 (41.8±0.5% versus 43.2±2.3%, p=0.97), S phase (32.3±2.4% versus 32.0±1.1%, p>0.99) or G2 (25.9±1.9% versus 24.8±3.3%, p=0.98) between the empty plasmid control and spike transfection (figure 4f). Representative histograms can be found in supplementary figure 1B. Western blotting was used to assess full-length and cleaved caspase-3, with no cleaved caspase-3 detected following spike transfection (supplementary figure 2B).

FIGURE 4.

FIGURE 4

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Transfection effects of the pCMV14-3X-FLAG-SARS-CoV-2 S spike-expressing plasmid or the pCMV14-3X-FLAG empty plasmid vector (control). a) Protein expression in spike-transfected 16HBE characterised via Western blotting against FLAG (a peptide tagged to the spike protein), Ki67 (a marker of proliferation) and p21Cip1 (a marker of cell cycle arrest). b) Relative comparison of densitometry between spike protein and Ki67 expression across various time-points. One-way ANOVA and Dunnett's post-test were used for statistical analysis. c) Densitometry analysis of p21cip1 expression normalised to β-actin. One-way ANOVA and Dunnett's post-test were used for statistical analysis. d) Culture proliferation was measured and cell counts are reported here. Two-way ANOVA with Šídák's multiple comparisons test was used. e) Secreted interleukin (IL)-6 and IL-8 of spike-transfected and control 16HBE cultures quantified using ELISA. An unpaired t-test was used for comparison. f) DNA cell cycle analysis with propidium iodide staining performed via flow cytometry. 6000 cells gated as positive were analysed and are summarised here with the cell population segregated based on their cell cycle phase. Two-way ANOVA with Šídák's multiple comparisons test was used. For all experiments, only pairwise comparisons p<0.05 are shown. *: p<0.05. **: p<0.01. ****: p<0.0001.

Spike S1 treatment and spike transfection reduce the repair rate of mechanically wounded 16HBE cultures

Next, we investigated whether spike treatment or transfection can impair repair following mechanical wounding of 16HBE monolayer cultures. Spike S1 treatment reduced the rate and extent of wound closure in 16HBE monolayers (figure 5a). By 48 h post-wounding, the control cultures were completely closed, whereas the S1-treated cultures still had open wounds. At all time-points, S1-treated cultures significantly reduced the percentage of wound closure compared to the control (figure 5b). At 48 h post-wounding, the percentage of wound closure was 99.5±3.9% with the control versus 76.6±1.5% with S1 treatment (p<0.0001). Wounds of transfected cultures remained significantly larger 48 h post-wounding compared to those in treated cultures (figure 5c). At 48 h post-wounding, the percentage of wound closure was 93.7±3.1% with the control versus 33.4±8.8% with spike transfection (p<0.0001). Wound closure was significantly lower with spike transfection compared to spike S1 treatment (p<0.0001). The change in wound closure with treatment or transfection between time-points was also characterised. There was a significant increase in wound closure between 12 and 24 h (p<0.0001) but not between 24 and 48 h post-treatment (p=0.25). Across these two pairs of time-points, there was no significant change in wound closure with spike transfection (12 versus 24 h, p=0.38, 24 versus 48 h, p=0.99).

FIGURE 5.

FIGURE 5

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Wound closure in 16HBE monolayer cultures reduced with spike treatment or transfection compared to empty plasmid transfection (control). A crosshatch wound model was used in monolayer cultures of 16HBE a) Representative images of wounded spike S1-treated and untreated control monolayers at ×40 magnification. The blue areas represent the wounds. b) Summary of the percentage (%) wound closure of S1-treated monolayers as a measurement of repair normalised to the initial size of the wound. Bars represent the mean±sem. Statistical comparisons were done with two-way ANOVA with Šídák's multiple comparisons test. c) Representative images of 12 wounded spike-transfected and empty plasmid-transfected monolayers (control) at ×40 magnification. The blue areas represent the wounds. d) Summary of % wound closure of spike-transfected monolayers with two-way ANOVA with Šídák's multiple comparisons test. ****: p<0.0001.

Discussion

Airway epithelial proliferation plays an essential role in airway repair [30]. Disruptions in airway epithelial repair can lead to increased paracellular permeability, enabling the passage of inhaled pathogens through the barrier and subsequent sensitisation by sub-epithelial immune cells [31]. Using the 16HBE submerged monolayer airway epithelial model, the findings presented in this study shed light on the interactions between the SARS-CoV-2 spike protein and airway epithelial repair processes.

We observed that key host viral entry factors, including ACE2, TMPRSS2 and furin, are expressed in 16HBE airway epithelial cells. These factors are pivotal for SARS-CoV-2 entry into host cells and their presence in 16HBE cells underscores the suitability of this cell line for modelling the interaction between the virus and the airway epithelium. Spike S1 treatment led to alterations in markers associated with cellular proliferation and/or cell cycle regulation. Specifically, a reduction in Ki67 expression, a marker of proliferation, was observed, coupled with an increase in p21Cip1 expression, which is suggestive of cell cycle arrest. p21Cip1 typically induces G1 phase arrest by inhibiting the activity of cyclin–cyclin-dependent kinase (CDK) complexes, particularly cyclin E/CDK2 and cyclin D/CDK4/6, which are crucial for G1 to S phase transition [32] and cyclin B1-CDK1, important in G2 to mitosis progression [33, 34]. As a result, this leads to the relative proportions of cells in G1 and G2 to increase. However, cell cycle analysis has revealed a significant decrease in the proportions of cells in G1 and an increase in the S phase. p21Cip1 may be phosphorylated, altering its function and allowing the bypass of G1 arrest, thereby promoting S-phase entry [35]. Further investigations into p21Cip1 dynamics and the effects of spike treatment, along with other potential underlying mechanisms, are needed to fully understand these observations. With spike transfection, Ki67 expression significantly decreased, whereas p21Cip1 expression and the distribution of cells in G1/S/G2 phases were unchanged. Ki67 expression varies throughout the cell cycle, accumulating during the S, G2 and M phases, while being continuously degraded in the G1 and G0 phases [36, 37]. In the G1 and G0 phases, its expression is heterogeneous and depends on the duration a cell remains in the G0 phase [38]. This suggests that spike transfection reduces the number of actively proliferating cells but does not induce cell cycle arrest at any particular phase. Functional assays have revealed a significant decrease in cell counts following spike treatment or transfection, indicating impaired cellular proliferation over time. The secretion of pro-inflammatory cytokines IL-6 and IL-8 was elevated, suggesting an inflammatory response induced by the spike protein. Additionally, both spike S1 treatment and spike transfection impaired wound closure in mechanically injured 16HBE cultures, potentially exacerbating barrier dysfunction and facilitating deeper invasion of virions beyond the epithelial barrier. In the lungs, activation of the epithelium by S1 or spike protein exposure following transfection, as indicated by increased IL-6 and IL-8 expression, can lead to the secretion of additional epithelial-specific cytokines that recruit immune cells, further compromising both immune and barrier functions of the epithelium [1]. Although studies investigating spike-mediated disruptions in repair mechanisms are limited, the SARS-CoV-2 spike protein has been observed to disrupt barrier integrity in endothelial linings, leading to vascular leakage [39–41]. Overall, these findings show that spike protein can interfere with cellular proliferation and repair.

A potential key factor in these effects may be the interaction between the spike protein and ACE2, the canonical receptor for SARS-CoV-2. Studies have characterised that spike expression on the host cell membrane can interact and bind with ACE2 receptors present on adjacent cells. Following cleavage by cellular proteases, the spike protein undergoes a conformational change that induces membrane fusion of the two cells, forming a syncytia [42]. It is hypothesised that spike-induced syncytia formation can promote cell–cell viral dissemination, replacing viral dissemination into the extracellular space and leading to immune cell evasion [43]. Lee et al. [44] observed that spike-induced syncytia formation can promote cellular senescence and inflammatory cytokine release, corresponding to the conditions observed in our study. However, whether spike treatment alone can induce syncytia formation remains to be investigated.

ACE2 primarily serves as a regulator of the renin–angiotensin–aldosterone system by cleaving angiotensin II, a potent vasoconstrictor, into angiotensin 1–7 [45]. Direct binding of the spike protein to ACE2 induces inflammation in the endothelium, significantly increasing IL-6 and monocyte chemoattractant protein-1 gene expression [46]. In a three-dimensional differentiated airway epithelial culture, spike treatment led to enrichment of pathways associated with interferon signalling and antiviral responses. In addition, an enrichment of p53 pathways was observed, which is associated with senescence within the airway epithelium [22]. However, ACE2 can also regulate other signalling pathways. ACE2 can bind integrins to increase cellular adhesion and affect integrin signalling that regulates migration, cytoskeletal organisation and proliferation [47, 48]. Spike protein binding to ACE2 can induce shedding and production of soluble ACE2 [49]. Soluble ACE2 can significantly reduce the phosphorylation of focal adhesion kinase and inhibit subsequent signalling through the NF-κB pathway [48]. This pathway may contribute to the reduction in proliferation observed in this study. However, our results do not provide an obvious explanation of the specific mechanisms underlying the spike-mediated effects of proliferation in the airway epithelium, prompting the need for further investigations.

This study has some limitations and areas for future investigation. We did not control for the contribution of migration in repair. Spike treatment of BEAS-2B bronchial epithelial cells was found to inhibit cell migration, which reduced repair [50]. As cell spreading and migration occur during the first 12–24 h and cellular proliferation begins 15–24 h post-injury [30], it is likely that the spike-mediated inhibition in repair observed in this study reflects the impact of the spike protein on cellular migration and proliferation. We observed that markers of proliferation and cell counts were reduced with spike treatment and transfection, suggesting the need for investigations into the contributions of cell death. Although cleaved caspase-3 expression was not observed with spike S1 treatment or spike transfection, further investigation using additional markers for apoptosis and caspase-independent pathways, such as by assessing annexin V and mitochondrial membrane potential [51, 52], is warranted. Additionally, necrosis, which has been observed in alveolar epithelial cells and contributes to lung inflammation and injury [53, 54], should also be investigated in future studies.

Overall, these findings advance our understanding of COVID-19 pathogenesis by highlighting the impact of the SARS-CoV-2 spike protein on airway epithelial repair processes, independent of viral replication. Additionally, these insights may be crucial for understanding how COVID-19 interacts with other chronic lung diseases, which are also characterised by chronic inflammation, impaired epithelial repair and reduced proliferative capacity [55–57]. Further research is needed to explore the specific mechanisms involved and identify potential therapeutic targets to mitigate the harmful effects of viral protein interactions with the airway epithelium, ultimately aiding the development of novel treatment strategies for COVID-19 and other chronic lung diseases.

Acknowledgments

We would like to thank members of the Dorscheid laboratory for their thoughtful contributions.

Footnotes

Provenance: Submitted article, peer reviewed.

Author contributions: Conceptualisation and design of the study: T. Guo, G. Singhera, J.M. Leung and D. Dorscheid; performing experiments: T. Guo, J. Memar Vaghri and W.Y. Liang; analysis of data: T. Guo; writing: T. Guo; review and editing: T. Guo, J. Memar Vaghri, W.Y. Liang, G. Singhera, J.M. Leung and D. Dorscheid; supervision: J.M. Leung and D. Dorscheid; funding acquisition: J.M. Leung and D. Dorscheid. All authors have read and agreed to the published version of the manuscript.

Conflict of interest All authors declare no competing interests. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data, in the writing of the manuscript, or in the decision to publish the results.

Support statement: T. Guo is supported by an award jointly funded by Asthma Canada, the Canadian Allergy, Asthma, and Immunology Foundation, and the Canadian Institutes of Health Research (CIHR) Institute of Circulatory and Respiratory Health. T. Guo is also supported by the Canadian Graduate Scholarships – Master's Program funded by the CIHR. J.M. Leung is a Tier 2 Canada Research Chair in Translational Airway Biology. D. Dorscheid acknowledges the CIHR, Michael Smith Health Research BC and the BC Lung Foundation. This study was funded by the Providence Healthcare Research Institute and the St Paul’s Foundation. Funding information for this article has been deposited with the Open Funder Registry.

Supplementary material

Please note: supplementary material is not edited by the Editorial Office, and is uploaded as it has been supplied by the author.

Figure S1

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DOI: 10.1183/23120541.00940-2024.Supp1
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Figure S2

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DOI: 10.1183/23120541.00940-2024.Supp1
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Data availability

Raw and analysed data obtained for this study are available from the corresponding author upon reasonable request.

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Supplementary Materials

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Figure S1

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DOI: 10.1183/23120541.00940-2024.Supp1
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00940-2024.SUPPLEMENT

Figure S2

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DOI: 10.1183/23120541.00940-2024.Supp1
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00940-2024.SUPPLEMENT2

Data Availability Statement

Raw and analysed data obtained for this study are available from the corresponding author upon reasonable request.

Articles from ERJ Open Research are provided here courtesy of European Respiratory Society

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