In 2019 a new coronavirus (SARS-CoV-2) was identified as the cause of an epidemic outbreak of an acute respiratory syndrome in Wuhan, China. SARS-CoV-2 used the same cell entry receptor—angiotensin converting enzyme II (ACE2)—as did SARS-CoV-1 (Zhou et al., 2020). The SARS-CoV-2 Spike protein mediates cell entry and is a single-pass transmembrane proteins that forms homotrimers. It has a large N-terminal ectodomain exposed to the exterior, a transmembrane helix, and a short C-terminal tail located within the virus. Each Spike monomer contains two regions termed S1 and S2. In the assembled trimer the S1 regions contain the receptor-binding domain while the S2 regions form a flexible stalk, which mediates membrane fusion between the viral envelope and the host cell membrane (Fan et al., 2020; Ke et al., 2020; Wrapp et al., 2020). The S1 region is divided into an N-terminal domain and a C-terminal domain, the latter interacts with the target cell ACE2 (Li et al., 2005; Benton et al., 2020; Lan et al., 2020; Ou et al., 2020; Shang et al., 2020a; Shang et al., 2020b; Yan et al., 2020). Like other coronavirus Spike proteins, SARS-CoV-2 Spike protein is heavily N-linked glycosylated (Walls et al., 2016; Lenza et al., 2020) and blocking N- and O-glycans dramatically reduced viral entry (Yang et al., 2020). The SARS-CoV-2 Spike proteins are activated by host cell proteases that cleave the protein at the S1-S2 boundary and subsequently at the S2' site (Hoffmann et al., 2020a, Hoffmann et al., 2020b, Takeda, 2022). Because Spike proteins are located on the surface of the virus, they are a major antigen targeted by the host immune system (Errico et al., 2022).

During natural infection host immune cells encounter Spike proteins via several different avenues. First, by direct contact with Spike protein bearing viral particles released from infected cells. Second, although the SARS-CoV-2 virions assemble in intracellular compartments of infected cells, unincorporated Spike proteins can reach the plasma membrane. Infected cells expressing Spike proteins may bind to cellular receptors present on resident or recruited immune cells. Third, extracellular vesicles (EVs) released by virally infected cells can contain Spike proteins. Mass spectrometry and nanoscale flow cytometry demonstrated SARS-CoV-2 Spike protein incorporation into EVs (Troyer et al., 2021). Fourth, Spike proteins are preprocessed during viral assembly utilizing the furin protease cleavage site between the S1 and S2 subunits (Takeda, 2022). When the Spike protein adopts a fusion conformation, the ACE2 receptor-binding domain separates from the membrane-bound S2 subunit. Soluble S1 subunits shed from infected cells or from the virions in vivo may bind to other cells via ACE2 or other binding partners. Intriguingly, 60% of the plasma samples from patients with post-acute sequelae of coronavirus disease 2019 had detectable levels of Spike protein using an ultrasensitive antigen capture assay (Swank et al., 2022). Prolonged exposure to Spike protein has been suggested to be responsible for Long-COVID syndrome (Theoharides, 2022). Humans also encounter SARS-CoV-2 Spike protein via vaccination (Thanh Le et al., 2020). Serious adverse events following vaccination are rare but include vaccine-induced immune thrombocytopenia and thrombosis; myocarditis and pericarditis; and a variety of autoimmune illnesses (Lamprinou et al., 2023). Due to waning effectiveness humans require repeated immunizations to maintain immunity and protection against potentially severe disease raising some concerns about the impact of repeated exposure to the SARS-CoV-2 Spike proteins.

To better understand the localization and trafficking of the SARS-CoV-2 Spike protein following administration and perhaps during natural infection, we prepared a recombinant SARS-CoV-2 Spike ectodomain stabilized in a prefusion conformation (Hsieh et al., 2020). This variant (S-2P) contained two consecutive proline substitutions in the S2 subunit. This double-proline substitution (SARS-CoV-2 S-2P) has allowed the rapid determination of high-resolution cryo-EM structures. Fluorescently labeled SARS-CoV-2 Spike ectodomain, a D614G variant (Yurkovetskiy et al., 2020; Zhang et al., 2020; Volz et al., 2021), a high mannose (Yang et al., 2020), and de-glycosylated version were injected into mice to characterize uptake and identify human mononuclear cells that bound these proteins, and to perform functional studies. In some instances, we used viral like particles (VLPs) expressing the full-length Spike protein. Our results identified the in vivo cellular tropism of the SARS-CoV-2 Spike proteins, delineated their human mononuclear cell targets, provided insights into their functional effects, thereby, helping afford a basis for understanding their impact on humans.

Preparation of SARS-CoV-2 Spike proteins and VLPs expressing them

We purified a stabilized exodomain of the original SARS-CoV-2 Spike protein (Hsieh et al., 2020) using media conditioned by transfected CHO-S or HEK293F cells. Because of higher yields most experiments used CHO-S-derived protein. The strategy for producing the recombinant protein is shown (Figure 1—figure supplement 1). Based on its mobility on size exclusion chromatography, the Spike proteins spontaneously formed trimers. Kifunensine-treated cultures were used to generate a high mannose version, and PNGase F-treated protein generated an N-glycan deficient version, and they will be referred to as such. The PNGase F treatment resulted in two distinct chromatography peaks that exhibited slightly different mobilities on SDS-PAGE. As discussed below we predominately used the preparation from fractions 9–11. The D614G mutant of the original Wuhan SARS-CoV-2 Spike protein was also purified. Each of the recombinant proteins was subjected to two rounds of Triton X-114 extraction to remove any residual lipopolysaccharide (LPS) (Aida and Pabst, 1990). The N-termini of the recombinant proteins were labeled with Alexa Fluor 488. In some experiments we used VLPs that expressed the original Wuhan Spike protein. We produced the VLPs using HEK293T cells along with a plasmid encoding the human immunodeficiency GAG protein fused to green fluorescent protein (GFP) and a full-length SARS-CoV-2 Spike protein expression vector. Mice received the Spike protein preparations or the SARS-CoV-2 VLPs by intranasal instillation, while only the Spike proteins were injected intravenously or subcutaneously.

Instillation of SARS-CoV-2 Spike proteins and Spike protein expressing VLPs affects the lung cellular composition and architecture

At various time points after Spike protein or VLP nasal instillation lungs from mice were processed for confocal microscopy. A Siglec-F antibody identified alveolar macrophages (AMs) while CD169 (Siglec-1) immunostaining delineated a subset of large airway-associated macrophages (Hetzel et al., 2021). Interstitial and inflammatory macrophages do not express Siglec-F. At 3 hr post instillation, low-magnification images of mouse lung sections revealed Spike protein (Trimer) uptake by Siglec-F-positive cells. Due to the uneven distribution of the instilled Spike protein, the Siglec-F-positive cells in the lower portion of the image lack signal (Figure 1A, Figure 1—figure supplement 2A). The Siglec-F-positive macrophages located near the bronchial epithelium and in the nearby alveoli initially accumulated the labeled protein (Figure 1B). Even at the 30 min time point AMs had already accumulated it (Figure 1C). Most of the macrophages that acquired the Spike proteins were Siglec-F positive, lacked CD169 (Figure 1D), but were wheat germ agglutinin positive, a mucin marker (Figure 1E). Repeating the experiment but substituting the Spike protein with VLPs expressing the SARS-CoV-2 Spike protein revealed a similar pattern of uptake by AMs, but with greater neutrophil recruitment and granulation compared to delta Env VLPs (Figure 1F, Figure 1—figure supplement 2B and C). Immunostaining with LYVE-1 and CD31 to identify the lymphatic and blood vessel endothelium, respectively (Figure 1G), or with Podoplanin, another lymphatic endothelial cell marker (Figure 1H, Figure 1—figure supplement 2D) showed the rapid accumulation of the Spike protein bearing VLPs in small and large lymphatic vessels. Within 3 hr of administration of either the VLPs (Figure 1I) or the Spike protein (data not shown), areas of alveolar collapse and lung damage were evident.

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Original image of the lung section in Figure 1B upper left.

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Original image of the lung section in Figure 1G.

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Original image of the lung section in Figure 1I.

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To assess the cellular response to Spike protein instillation and to delineate the cells that had acquired Spike protein in vivo, we collected mouse lungs 18 hr post exposure to different SARS-CoV-2 Spike protein preparations. Saline instillation served as a control. Lung fragments were digested, and then pushed through a 40 µm cell strainer. The cells collected were analyzed using flow cytometry following the established gating strategy (Yu et al., 2016). The results showed that the SARS-CoV-2 Spike protein increased the number of neutrophils, monocytes, and dendritic cells in the lung, but did not affect lymphocyte or eosinophil numbers (Figure 2A). Due to the uneven distribution of the instilled protein and because of their rapid turnover, the neutrophil numbers likely underestimate the local neutrophil recruitment. As the imaging experiments indicated AMs most avidly acquired the Spike protein, some interstitial macrophages also retained it, while only a low percentage of the lung neutrophils and dendritic cells were positive following isolation (Figure 2B). Instillation of the glycan-deficient Spike protein reduced the monocyte cellular infiltrate and decreased the % of AMs that retained the protein. The high mannose Spike protein slightly increased the numbers of neutrophil and macrophages in the lung despite a lower uptake by alveolar and interstitial macrophages. Finally, the D614G mutation reduced the cellular infiltrate compared to the unmutated protein and had a slightly different cell binding profile as a greater percentage of monocytes, eosinophils, and dendritic cells retained it (Figure 2A and B). We also tested whether the addition of human ACE2 (hACE2) affected the cellular uptake of the Spike protein following intranasal administration by comparing wild-type and the hACE2 transgenic mice. While we saw little difference by imaging, we did note some minor changes in the cellular uptake pattern, most notably an increase uptake by AM and a decrease uptake by interstitial macrophages (Figure 2—figure supplement 1).

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Altered lung vascular permeability, neutrophil recruitment, and lung damage 3 hr post instillation of the Spike protein

The increase in lung leukocytes following the Spike protein nasal instillation suggested that it may have altered the vascular permeability of the lung. To assess whether vasculature permeability changes had occurred, we intravenously injected Evans blue dye, which in the absence of a permeability defect remains confined to the vasculature (Matthew et al., 2002). Prior to injecting the Evans blue, we instilled in the nasal cavity the SARS-CoV-2 Spike protein, the S1 subunit of the HCoV-HKU1 Spike protein, or a saline control. Inspection of the lungs from the SARS-CoV-2 spike protein-treated mice revealed a strong increase in Evan blue staining while the lungs from the S1 subunit HCoV-HKU1-treated mice had a minimal increase. Quantifying the Evans blue dye in collected lungs and liver confirmed the increase in vasculature permeability in the lungs of the SARS-CoV-2 Spike protein-treated mice (Figure 3A). Next, we compared the S1 subunit of SARS-CoV-2 Spike protein to the HCoV-HKU1 S1 subunit. The S1 subunit preparations were purified from transfected HEK 293 cells. The results of administrating the S1 subunit of HCoV-HKU1 Spike protein did not differ from the saline control while the S1 subunit from SARS-CoV-2 like the SARS-CoV-2 trimer increased the lung vascular permeability to Evans blue (Figure 3—figure supplement 1A). We also tested whether the presence of hACE2 affected the response by using K18-hACE2 transgenic mice as the recipients. The presence of the K18-hACE2 transgene slightly increased the lung vasculature permeability upon SARS-CoV-2 trimer instillation (Figure 3—figure supplement 1B). Consistent with the increase in lung vascular permeability 3 hr post SARS-CoV-2 Spike protein instillation, Siglec-F-positive lung macrophages bearing the Spike protein were surrounded by neutrophils and neutrophil fragments (Figure 3B and C).

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Next, we investigated the potential impact of modifying the glycans displayed by the spike proteins on lung vasculature homeostasis and neutrophil recruitment. Imaging fixed lung tissue from mice instilled with saline, SARS-CoV-2 Spike protein, glycan-deficient protein, high mannose protein, and the HCoV-HKU1 S1 subunit recombinant protein revealed enhanced neutrophil recruitment with each SARS-CoV-2 Spike protein preparation. We analyzed the neutrophil count in a 200×200 μm² area at six different locations. The surveyed areas of the SARS-CoV-2 Spike protein instilled mice had approximately a 6.5-fold higher neutrophil density compared to the saline-treated control mice. The high mannose version of the Spike protein recruited fewer neutrophils at the 3 hr time point while the PNGase F-treated version did not differ from the untreated trimer (Figure 3D, Figure 3—figure supplement 1C).

Neutrophil recruitment and damage in the cremaster muscle following local protein injection, and in the liver following intravenous injection

An exteriorized cremaster muscle is commonly used to intravitally image mouse neutrophil transmigration and interstitial motility (Yan et al., 2021). Although this model lacks Siglec F-positive macrophages, it is worth monitoring the effect of the SARS-CoV-2 Spike protein on neutrophils recruited into an inflammatory site. Within 3 hr local IL-1β injection recruits bone marrow neutrophils and leads to their transmigration into the nearby tissue. Co-injecting fluorescently labeled Ly6G and Gp1bβ antibodies identified neutrophils and platelets, respectively. Local PBS injection and intermittent imaging over a 6 hr time frame revealed occasional blood neutrophil and numerous flowing platelets (Figure 4A, Video 1). Local intrascrotal injection of unlabeled SARS-CoV-2 Spike protein caused a modest recruitment of neutrophils at 4–6 hr post injection (Figure 4A, Figure 4—figure supplement 1, Video 1). To better assess the long-term effect of the SARS-CoV-2 Spike protein, we co-injected IL-1β and waited until the following day to image. Typically, at 24 hr post IL-1β the inflammatory response is resolving, and the interstitial neutrophil numbers are declining from their peak (Video 2). In contrast, each of the Spike protein preparations adversely affected neutrophil motility and morphology. Numerous neutrophils and neutrophil fragments localized along the blood vessel walls and were scattered within the interstitium (Figure 4B, Video 2). We did not note any significant difference between the different SARS-CoV-2 Spike protein preparations as each caused neutrophil fragmentation and a decline in mobile neutrophils (Figure 4C).

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We also assessed the impact of injecting the labeled SARS-CoV-2 protein into the blood on the liver sinusoids using intravital microscopy. Snapshots at 3 hr post infusion revealed Spike protein outlined liver sinusoid endothelial cells co-localized with the CD31 delineated sinusoid endothelial membranes. Kupffer cells identified by F4-80 immunostaining rapidly acquired large amounts of the infused Spike protein (Figure 4—figure supplement 2A, Video 3). The sinusoids, normally devoid of neutrophils, contained many Ly6G⁺ neutrophils. However, by 18 hr post infusion the amount of Spike protein outlining the sinusoids had declined as had the Kupffer cell-associated material (Figure 4—figure supplement 2B). The neutrophil infiltration had largely resolved suggesting that the inflammatory signals had declined. The origin of the white dots interspersed between the sinusoids is unknown, but they were not present at 3 hr post infusion despite identical imaging conditions. One possibility is that altered vascular permeability had allowed some labeled antibodies to leak into the liver parenchyma. To determine whether other endothelial beds also acquired intravenous administered Spike protein, we examined the spleen, heart ventricle, and Peyer’s patches. At 3 hr post infusion endothelial cells at these three sites had acquired the labeled Spike proteins, most prominently in Peyer’s patches (Figure 4—figure supplement 3A–C). Finally, we intravitally imaged the inguinal lymph node at 3 and 18 hr after local injection. Typically, locally injected material rapidly enters nearby afferent lymphatics for delivery to the lymph node where subcapsular sinus macrophages first encounter it (Park et al., 2015). If it bypasses these macrophages lymph-borne material flows into the lymph node medullary region, where medullary macrophages reside (Figure 4—figure supplement 3D). At both the 3 hr time point (data not shown) and at 18 hr subcapsular macrophages showed little interest in the Spike protein, but medullary macrophages avidly acquired it. In contrast to the liver, we did not detect neutrophil recruitment into the lymph node at either time point after tail base injection.

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Murine and human neutrophils NETosis following exposure to the SARS-CoV-2 Spike protein

Live cell imaging of lung sections following intranasal instillation of the Spike protein revealed ongoing neutrophil damage and likely NETosis (Figure 5A). Time-lapse images show several disrupted neutrophils near a Siglec-F-positive AM along with Spike protein bound to a dying neutrophil. This observation along with the cremaster muscle imaging suggested direct neutrophil toxicity. Furthermore, a previous study had shown that Spike protein induced neutrophil NETosis (Youn et al., 2021). To confirm that the Spike protein can trigger neutrophil NETosis and to compare different Spike protein preparations, we briefly exposed murine bone marrow neutrophils and assessed the % of cells undergoing cell death using a flow-based assay. Exposure to N-formylmethionyl-leucyl-phenylalanine (fMLP) served as a positive control. The lowest concentration of Spike protein (0.1 µg) tested increased the number of dying neutrophils (Figure 5B). Switching to human neutrophils purified from human peripheral blood, exposure to tumor necrosis factor-α (TNF-α), LPS, or phorbol myristate acetate (PMA) for 4 hr or overnight reduced their viability as expected. Addition of SARS-CoV-2 or the D614G mutant protein (1 µg/ml) decreased their viability at 4 hr and more dramatically upon overnight exposure. The high mannose versions of the Spike protein and D614G Spike had a slightly greater toxicity at 4 hr but had a similar impact in the overnight assay (Figure 5C). Finally, we imaged human neutrophils overlaid on SARS-CoV-2 Spike protein-treated A549 cells, a human lung epithelial cell line. Numerous dying neutrophils could be observed likely undergoing NETosis (Figure 5D, Video 4). These results confirm that the SARS-CoV-2 Spike protein can cause neutrophil damage potentially exacerbating the inflammatory response.

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Human peripheral blood monocytes, B cells, neutrophils, and dendritic cells bind the SARS-CoV-2 Spike protein

Next, we assessed human peripheral blood leukocyte binding using the fluorescently labeled Spike protein. Binding assays were performed on ice to avoid endocytosis using Hanks’ Balanced Salt Solution (HBSS) with added Ca²⁺ and Mn²⁺, or with EDTA to assess cation dependence. Representative CD4 T cell, B cell, monocyte, or neutrophil flow patterns using either the Spike protein or the high mannose version are shown (Figure 6A). The flow cytometry results demonstrated that cations enhanced leukocyte binding particularly so with the high mannose protein. The SARS-CoV-2 S1 protein bound better than did the HKU1 S1 protein, most evident with NK cells, neutrophils, monocytes, and dendritic cells (Figure 6B). The SARS-CoV-2 Spike protein bound better than did the S1 protein with B cells, neutrophils, and monocytes exhibiting the best binding. The high mannose protein bound less well to the different leukocyte subsets. Surprisingly CD4 T cells poorly bound each of the proteins, irrespective of the presence or absence of cations. The D614G Spike protein bound better to neutrophils and monocytes than did the wild-type Spike protein (Figure 6B).

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Murine and human cells use Siglecs to help capture the SARS-CoV-2 Spike Protein

Since murine and human leukocytes lack significant ACE2 levels, the major entry receptor for SARS-CoV-1 and -2, they likely use other receptors to capture the Spike proteins. The strong co-localization with Siglec-F expressing murine AMs prompted an examination of the role of Siglec-F and other Siglecs in capturing the Spike proteins. Siglecs are transmembrane proteins that exhibit specificity for sialic acids attached to the terminal portions of cell surface glycoproteins. Several viruses take advantage of sialic acid-Siglec interactions for cell targeting, spreading, and trans-infection (Hammonds et al., 2017; Perez-Zsolt et al., 2019; Perez-Zsolt et al., 2021). Initially, we established a bead assay to assess the binding of SARS-CoV-2 Spike proteins. We coupled the S1 domain protein, the SARS-CoV-2-stabilized Spike protein, or the PNGase F-treated Spike protein and reacted the beads with fluorescently labeled antibody or different recombinant proteins. Flow cytometry analysis revealed strong binding of the Spike protein antibody, hACE2, but not murine ACE2 as expected (Figure 7A). Siglec-F also bound well, while the human Siglec-5 and Siglec-8 bound poorly despite being the structural and functional equivalents of Siglec-F, respectively (Connolly et al., 2002). Of note the PNGase F-treated fraction we used likely retained some N-linked glycans as it remained able to bind ACE2 although it lost Siglec-F binding (Figure 7A).

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Original image of a human Siglec-5-GFP transfected HEK293 cell in Figure 7C, 3D reconstruction.

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Despite the poor binding to the recombinant human Siglecs in the bead assay, we elected to test them in context of a human cell by expressing them in HEK293 cells. We chose HEK293 cells as they exhibited little Spike protein binding. We established ACE2, Siglec5, Siglec-8, Siglec-5/8, ACE2/Siglec-5, and ACE2/Siglec-8 expressing cell lines (Figure 7B). The ACE2 transfected cells behaved as anticipated binding the original Spike protein and even better the D614G version; and binding the S1 protein. The high mannose and glycan-deficient Spike proteins bound poorly. In contrast to the bead assay, both the Siglec-5 and the Siglec-8 expressing HEK293 cells bound the Spike protein, although less efficiently than did the ACE2 expressing cells. They bound the D614G version less well, and very weakly bound the S1 domain protein. The high mannose and glycan-deficient Spike proteins exhibited little binding to the Siglec expressing cells. Co-expression of Siglec-5 and -8 did not improve binding, although the ACE2/Siglec co-expressing cells bound the SARS-CoV-2 Spike protein slightly better than did ACE2-only cells. We also imaged the Siglec-5 expressing HEK293 with labeled protein (Figure 7C). The imaging revealed efficient uptake and Spike protein endocytosis. Co-cultured ACE2 and Siglec-5 expressing cells both captured the Spike protein while HEK293 cells not expressing either protein failed to bind or uptake it (Figure 7D, Video 5). To determine whether Siglec-5 or Siglec-8 could contribute to viral transduction, we produced lentiviral particles expressing the Spike protein from SARS-CoV-2, HKU1, or 229E and attempted to transduce the HEK293 ACE2 expressing cells. Successful transduction resulted in GFP expression, which we quantitated by flow cytometry and only occurred with the lentiviral particles with SARS-CoV-2 Spike protein incorporation (Figure 7E). Both a receptor binding domain antibody and recombinant ACE2 blocked transduction. Recombinant human Siglec-5 partially inhibited although it required relatively high concentrations, while human Siglec-1, human Siglec-2, and murine Siglec-F had no impact on the transduction frequency.

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Increased TNF-α and IL-6 production by human macrophages exposed to the SARS-CoV-2 Spike proteins

Dysregulated cytokine production contributes to the pathogenesis of severe COVID-19 infections (Vora et al., 2021; Sefik et al., 2022). To assess whether the Spike proteins can affect macrophage cytokine profiles, we cultured human monocytes using conditions that generate M0, M1, or M2 macrophages and treated them with different Spike protein preparations. We collected cell supernatants 2 days later and measured a panel of known macrophage-derived cytokines by ELISA (Figure 8). We found little induction of IL-1β in the cell supernatant indicating that the Spike proteins alone did not trigger the inflammasome activation in these macrophages. In contrast, we did observe significant increases in TNF-α and IL-6 secretion by the original SARS-CoV-2 Spike protein-treated macrophages irrespective of whether it was produced in CHO or HEK293F cells. The spike protein produced in HEK293F is predicted to contain more complex carbohydrates than that expressed in CHO cells. While both preparations increased IL-6 and TNF-α production, the spike protein produced by HEK293F cell elicited more TNF-α and less IL-6 compared with the CHO-produced spike protein. The spike protein produced in CHO cells treated with Kifunesine elicited a similar cytokine profile as did the spike protein produced in CHO cells in the absence of Kifunesine. The addition of the HKU1 S1 protein did not modify cytokine production by any of the macrophage subsets.

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This study analyzed the cell tropism of the SARS-CoV-2 Spike protein and some of its variants using fluorescently labeled proteins, flow cytometry, and mice to assess in vivo binding. The intranasal administration of SARS-CoV-2 Spike protein to mice led to its rapid uptake by Siglec-F-positive AMs and an increase in the number of neutrophils, monocytes, and dendritic cells in the lung 18 hr after instillation. Modifying the carbohydrate content of the Spike protein or using a Spike protein with a D614G mutation slightly altered the cell recruitment pattern. The high mannose Spike protein purified from the Kifunensine-treated CHO cells increased the number of lung macrophages at 18 hr despite a lower % of macrophages having retained it. As AMs express the mannose receptor (MR, CD206), a member of the C-type lectin (CLEC) family, perhaps the high mannose Spike protein is more rapidly endocytosed and degraded accounting for a lower percentage of cells retaining it. This receptor binds high mannose structures present on the surface of pathogens helping to neutralize them by phagocytic engulfment. Other cell types including immature dendritic cells, and endothelial cells in hepatic, splenic, lymphatic, and dermal microvasculature express the MR and would be expected to be targeted by the high mannose Spike protein. We confirmed Siglec-F as a capturing receptor, as it efficiently bound to the S1 domain and the SARS-CoV-2 Spike protein. Yet AMs employ additional receptors to capture the Spike protein as they still accumulated the PNGase F-treated Spike protein, which had lost binding to Siglec-F.

Arguing against a role for mSiglec-1, CD169⁺ lung macrophages and CD169⁺ subcapsular sinus macrophages failed to accumulate the Spike protein in vivo. The closest human paralog of mouse Siglec-F is hSiglec-8 (Aizawa et al., 2003). While expressed on human eosinophils and mast cells, human AMs apparently lack it. In contrast, human AMs do express Siglec-5 (Connolly et al., 2002). Along with its paired receptor, hSiglec-14, Siglec-5 can modulate innate immune responses (Tsai et al., 2020). When tested in a bead binding assay, in contrast to Siglec-F, neither hSiglec-5 or -8 bound the recombinant Spike protein, yet their expression in a cellular context allowed binding. Additionally, recombinant hSiglec-5 partially inhibited the transduction of Spike protein bearing VLPs into ACE2 expressing HEK293 cells. Evidently, the recombinant protein binding assay we employed did not fully capture the properties of hSiglec-5 or hSiglec-8 in a cellular context. A recent study of human lungs revealed scant alveolar ACE2 expression, highlighting the importance of alternative Spike protein receptors (Hönzke et al., 2022). Ex vivo infected human lungs and COVID-19 autopsy samples showed AMs positive for SARS-CoV-2 and single-cell transcriptomics revealed nonproductive virus uptake, and activation of inflammatory and anti-viral pathways. Another study which analyzed pulmonary cells from COVID-19 patients found that myeloid cell C-type lectin engagement induced a robust proinflammatory responses that correlated with COVID-19 severity (Lu et al., 2021). The robust Spike binding to AMs and their potential roles in COVID-19 lung infection warrants a further exploration of their Spike protein binding partners.

The intranasal administration of the Spike protein resulted in an increase in lung vascular permeability and local tissue damage. While the mechanisms are unknown, infectious agents that target AMs can trigger the secretion of factors that disrupt the integrity of the pulmonary microvascular cell barrier. For example, Porcine reproductive and respiratory syndrome virus-infected AMs trigger transcriptional changes in genes in co-cultured microvascular endothelial cells that reduce vascular integrity (Sun et al., 2022). Lung endothelial cells may also be directly targeted (Bernard et al., 2020). Spike protein/ACE2 interactions reduce endothelial cell ACE2 expression, which can alter vascular permeability. Yet, the low affinity of mouse ACE2 for the SARS-CoV-2 Spike protein presumably precludes this mechanism in this study. Also arguing against a role for ACE2 in our model, the responses to intranasal Spike protein in hACE2 transgenic mice resembled those of wild-type mice. The Spike protein RGD (arginine-glycine-aspartic acid) motif can also bind α_Vβ₃ integrins present on endothelial cells (Nader and Kerrigan, 2022). This affects VE-cadherin function and vascular integrity. However, we did not detect any Spike protein on the pulmonary blood vessels following intranasal administration although a low level of binding may have escaped visualization. In sum, the rapid and intense uptake of Spike protein by AMs supports a role for these cells in the localized neutrophil recruitment, nearby tissue damage, and increased vascular permeability that follows its intranasal administration.

The intravenous administration of the Spike protein led to its accumulation by Kupffer cells in the mouse liver. This uptake was accompanied by a transitory increase in liver sinusoid neutrophils. Human Kupffer cells express LSECtin (liver Siglec and lymph node sinusoidal endothelial cell C-type lectin, CLEC4G) a C-type lectin receptor encoded within the L-SIGN/DC-SIGN/CD23 gene cluster. LSECtin acts as a pathogen attachment factor for Ebola virus and the SARS coronaviruses suggesting that it contributed to the Kupffer cell uptake we noted (Domínguez-Soto et al., 2009). The liver sinusoid endothelial cells also avidly accumulated the Spike protein. These cells express another C-type lectin receptor L-SIGN (CD209L/CLEC4M), which has been shown to interact in a Ca²⁺-dependent manner with high mannose-type N-glycans on the SARS-CoV-2 Spike protein (Kondo et al., 2021). Based on our mouse studies, blood-borne SARS-CoV-2 Spike proteins, viral particles, or exosomes bearing the Spike protein are likely to accumulate in the liver during severe or prolonged COVID-19 infection.

Local injection of the Spike protein in the region of the cremaster muscle recruited low numbers of neutrophils, but co-administration with IL-1β caused severe neutrophil damage. The Spike protein also proved toxic to purified mouse and human neutrophils cultured ex vivo in its presence. Most human neutrophils bound the recombinant Spike protein. The presence of cations in the binding buffer substantially enhanced the binding suggesting that a cation-dependent receptor accounted for a significant portion of the binding. While the high mannose version bound neutrophils less well, in the NETosis assay it produced a similar level of neutrophil cell death. Neutrophils likely employ several different receptors to capture the Spike protein. Human neutrophils express several C-type lectin receptors including CLEC5A, which has been implicated in SARS-CoV-2-triggered neutrophil NETosis (Sung et al., 2022). They also express Siglec-5 (Connolly et al., 2002), which bound the Spike protein when expressed in HEK293 cells.

We also assessed the binding of various Spike protein preparations to human peripheral blood mononuclear cells (PBMC) and their impact on cytokine production by M0, M1, and M2 human monocyte-derived macrophages. Monocytes, neutrophils, and B cells bound the full-length trimer best, while T and NK cells exhibited little binding. All the cellar subsets bound more of the SARS-CoV-2 Spike protein in the presence of cations arguing for a major role for cation-dependent receptors. The high mannose Spike protein bound less well to all tested cell types and its binding exhibited the highest cation dependence. Among the eight cytokines measured in the supernatants conditioned by human monocyte-derived macrophages treated with Spike proteins, only TNF-α and IL-6 were elevated compared to untreated or HKU1 S1-treated cells. We found little or no change in IL-1β levels suggesting that the tested Spike protein preparations did not activate inflammasomes. Nor did we find any material difference between recombinant Spike protein prepared from HEK293F and CHO cells. Further studies are needed to identify the monocyte, neutrophil, and B cell receptors that account for the Spike protein binding and additional functional studies to assess the consequences of that binding.

Several conclusions can be drawn from this study. First, instilled SARS-CoV-2 Spike protein and VLPs bearing the Spike protein rapidly accumulated on mouse AMs suggesting a similar uptake by human AMs during an active SARS-CoV-2 infection. In mice, the nasal instillation of the Spike protein is accompanied by lung leukocyte infiltration. Second, murine AMs likely use Siglec-F as a capturing receptor although other receptors contribute. Third, following nasal instillation of the Spike protein, the integrity of the pulmonary vasculature weakens. Fourth, resident and recruited neutrophils suffer, either directly targeted by the Spike protein or secondarily due to the inflammatory response. A similar scenario would be expected during SARS-CoV-2 infection, following direct instillation of the Spike protein in the upper airway of humans, or following an intranasal vaccine that directs the synthesis of the SARS-CoV-2 Spike protein or fragments. Fifth, blood monocytes, B cells, neutrophils, and dendritic cells efficiently bind the Spike protein, while only a low percentage of CD8 T cells and even fewer CD4 T cells do. Spike protein binding to human monocyte-derived macrophages increases the TNF-α and IL-6 secretion. Sixth, altering the glycan composition of the Spike and the D614G mutation had surprisingly little impact on the initial inflammatory response. Seventh, viral particles and soluble Spike protein that spills into the blood during SARS-CoV-2 infection or following immunization will likely be cleared by liver Kupffer cells but will also target blood vessel endothelial cells in multiple organs. Finally, injection of the Spike protein as occurs following vaccination rapidly delivers it to the draining lymph node via afferent lymphatics. Surprisingly, lymph node medullary macrophages, which largely serve a degradative function, preferentially uptake the Spike protein. Designing a less toxic Spike protein immunogen that targets subcapsular sinus macrophages might better deliver the immunogen to B cells, thereby eliciting a more efficacious antibody response.

Mice

C57BL/6 and K18-ACE2 transgenic mice were obtained from Jackson Laboratory. All mice (female) used in this study were 8–12 weeks of age. Mice were housed under specific pathogen-free conditions. All the animal experiments and protocols used in the study were approved by the National Institute of Allergy and Infectious Diseases (NIAID) Animal Care and Use Committee (ACUC) at the National Institutes of Health.

Cells

To isolate mouse lung cells, lungs were carefully collected and gently teased apart using forceps into RPMI 1640 media containing 2 mM L-glutamine, antibiotics (100 IU/ml penicillin, 100 μg/ml streptomycin), 1 mM sodium pyruvate, and 50 μM 2-mercaptoethanol, pH 7.2. The tissue was then digested with Liberase Blendzyme 2 (0.2 mg/ml, Roche Applied Science) and DNase I (20 μg/ml) for 1 hr at 37°C. The proteases were inactivated by adding 10% fetal bovine serum and 2 mM EDTA and the cell disaggregated by passing them through a 40 μm nylon sieve (BD Bioscience). Single cells were then washed with 1% bovine serum albumin (BSA) in PBS and blocked with anti-Fcγ receptor (BD Biosciences). Human PBMC were purified from whole blood by density gradient centrifugation (FicollPaque, Miltenyi Biotec). Whole blood was collected from healthy donors through an NIH Department of Transfusion Medicine (DTM)-approved protocol (Institutional Review Board of the NIAID). Neutrophil and monocyte cell population were each obtained by negative selection (>97% purity, Stem Cell Technologies). To generate human monocyte-derived macrophages, purified monocytes were treated for 7 days with 50 ng/ml human recombinant M-CSF (PeproTech) in RPMI medium supplemented with 10% heat-inactivated fetal bovine serum, and 1 mM sodium pyruvate, 100U Penicillin-Streptomycin (Gibco, Thermo Fisher Scientific) in ultra-low attachment culture 100 mm dishes. On day 7 the mature macrophages were collected and verified to be more than 90% CD68⁺ by flow cytometry.

Reagents

See Key resources table.

Flow cytometry

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Single cells were re-suspended in 0.1% fatty acid-free BSA-HBSS with 100 μM CaCl₂ and 1 mM MnCl₂ unless otherwise specified. Buffer without divalent cations included 10 mM EDTA. The cells were stained with fluorochrome-conjugated antibodies against various cell surface markers or with different fluorochrome-conjugated proteins, which are listed in the resource and reagent tables in the supplement. LIVE/DEAD Fixable Aqua Dead Cell Stain Kit, LIVE/DEAD Fixable NearIR Dead Cell Stain Kit, or LIVE/DEAD Fixable Yellow Dead Cell Stain Kit (Thermo Fisher) were used in all experiments to exclude dead cells. Compensation was performed using AbC Total Antibody Compensation Bead Kit (Thermo Fisher) and ArCTM Amine Reactive Compensation Bead (Thermo Fisher) individually stained with each fluorochrome. Compensation matrices were calculated with FACSdiva software. Data acquisition including cell number count was done on a FACSCelesta SORP (BD) flow cytometer and analyzed with FlowJo 10.8.x software (Tree Star).

Human macrophage polarization and cytokine profiling

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To polarize human macrophages, monocyte-derived macrophages were plated in 48-well plates at 5×10⁴ cells per well in 500 µl RPMI medium and allowed to rest for 2 hr at 37°C before treating cytokines or LPS. The human M0 macrophages were either left untreated or treated for 48 hr to induce macrophage polarization: (i) for M1 polarization with 10 ng/ml LPS (E055:B55; Sigma-Aldrich) and 20 ng/ml IFNγ (PeproTech), (ii) for M2 polarization with 50 ng/ml human recombinant M-CSF, 20 ng/ml human recombinant IL-4, and 20 ng/ml human recombinant IL-13 (PeproTech). Human macrophages were then cultured alone or with 0.1 or 1 µg SARS-CoV-2 Spike proteins for 48 hr. The cultured supernatants were collected, and cytokine levels determined using the bead-based immunoassays LEGENDplex Human Macrophage/Microglia Panel (BioLegend). The assays were performed in 96-well plates following the manufacturer’s instructions. For measurements, a FACSCelesta SORP flow cytometer (BD Biosciences) was employed, and data were evaluated with the LEGENDplex Data Analysis software.

Production and purification of recombinant SARS-CoV-2 Spike proteins (see Figure 1—figure supplement 1)

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SARS-CoV-2 Spike expression vectors were purchased from Addgene (Key resources table). Endotoxin-free plasmids were prepared by Alta Biotech and provided at a concentration of 5 mg/ml. Transfections were carried out using CHO Freestyle cells (Invitrogen). 1.24 mg of plasmid DNA was transfected into 90 million cells, using a MaxCyte Electroporation Transfection System. A detailed protocol is available at https://maxcyte.com/atx. Culture supernatants were harvested on day 6 and clarified by centrifugation, followed by filtration through a 0.45 µm filter, followed by the addition of EDTA-free protease inhibitor cocktail tablets (Roche). Culture supernatants were then dialyzed overnight at 4°C in HBS (150 mM NaCl, 10 mM HEPES, pH 8.0) using 10 kDa MWCO Slide-A-Lyzer dialysis cassettes (Thermo Scientific). Supernatants were passed over a 10 ml StrepTrap HP column (Cytiva) at 1 ml/min using an ÄKTA pure 150 purifier (Cytiva), maintained at 4°C. Bound protein was eluted with elution buffer (2.5 mM desthiobiotin, 100 mM Tris-Cl, 150 mM NaCl, 1 mM EDTA, pH 8.0). Peak fractions were pooled and concentrated using 30 kDa MW CO Amicon centrifugal concentrators (Millipore). Trace endotoxins were removed by two sequential Triton X-114 extractions, followed by passage through an HiPPR detergent removal column (Thermo Fisher). Following the removal of endotoxin, the remaining contamination was assessed using the Pierce Chromogenic Endotoxin Quant Kit (Thermo Fisher), based on the amebocyte lysate assay. The endotoxin level in the purified recombinant protein preparation is below 1.0 EU/ml, which closely aligns with the levels specified by the company for recombinant proteins. Protein concentrations were determined by a BCA Protein assay (Thermo Fisher).

VLPs and lentivirus preparation

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SARS-CoV-2 Spike protein-incorporated NL4.3-GFP VLPs were produced by transfecting HEK293T cells with full-length SARS-CoV-2 Spike-S and HIV-1 NL4-3 Gag-iGFP ΔEnv (12455, NIH AIDS Reagent Program) at a ratio of 1:2.5 using a previously reported method (Park et al., 2015). EGFP control lentiviruses (no Spike protein) were produced by transfecting HEK293T cells with pCMV-dR8.2 dvpr (packaging) and pLentipuro3 TO V5-GW EGFP-Firefly Luciferase (reporter). Coronavirus Spike proteins were incorporated by co-transfection with SARS-CoV-2 Spike-S, HCoV-229E Spike, or HCoV-HKU1 Spike expressing plasmids. Eighty percent confluent HEK293T in six-well plates (2.5 ml/well) were transfected with 5 μg of plasmid diluted in Opti-MEM with a 1:4 (DNA/reagent) dilution of TransIT-293 Transfection Reagent. The media was harvested 64 hr later and centrifuged at 500×g for 10 min at 4°C. The supernatants were collected and mixed with Lenti-X Concentrator (Takara Bio USA, Inc) at a 1:3 ratio. The mixture was placed at 4°C overnight and centrifuged the following day at 1500×g for 45 min. Fluorescent VLPs were directly counted and measured by BD FACSCelesta flow cytometer. The forward scatter (FSC) detector (photodiode with 488/10 BP filter) and side scatter (SSC) detector (photomultiplier tube [PMT] with 488/10 BP filter) were tuned to detect voltages up to 530 for FSC and 220 for SSC. The NL4.3-GFP VLPs were distinguished from noise and non-VLP particles by GFP signals. NL4.3-GFP VLP Spike protein incorporation was tested using hACE2 expressing HEK293 cells (ACE2/HEK293). Ten thousand VLPs were incubated with 1×10⁴ ACE2/HEK293 cells in 1× HBSS (contains 1 mM Ca²⁺, 2 mM Mg²⁺, 1 mM Mn²⁺, and 0.5% fatty acid-free BSA) buffer at 4°C for 30 min. Binding of the VLPs to ACE2/HEK293 cells was measured with BD FACSCelesta. The 50% lentivirus transduction dose was determined using percentile (50%) of GFP-positive ACE2/HEK293 cells 3 days after transduction. The pelleted VLPs or lentiviruses were suspended in PBS and frozen in single use aliquots.

Lentivirus transduction assay

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96-well plates were seeded with 200 µl of ACE2/HEK293 cells (1.5×10⁴ cells/ml). After a 24 hr incubation, recombinant proteins or neutralizing antibody was added. Lentiviral particles capable of transducing 50% of the cells (CoV2-lenti, HKU1-lenti, and 229E-lenti) were added (~25 µl) 30 min later. Following a 3-day culture, the ACE2/HEK293 cells were harvested and GFP expression levels determined by BD FACSCelesta flow cytometry. Each group contained two to five replicates.

Measurement of vascular permeability

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Pulmonary vascular permeability was measured by i.v. administration of Evans blue dye (0.2 ml 0.5% in PBS) (Zhou et al., 2011). One and half hours after Spike protein nasal administration, Evans blue dye was injected intravenously. After another 1.5 hr the mice were perfused with PBS, and lungs and livers were harvested, and dye extracted in formamide overnight at 55°C. Dye concentrations were quantified by measuring absorbance at 610 nm with subtraction of reference absorbance at 450 nm. The content of Evans blue dye was determined by generating a standard curve from dye dilutions.

NETosis assays

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Mouse bone marrow-derived or human neutrophils (>97% purity) were re-suspended in RPMI 1640 media (1×10⁶/ml) and incubated at 37°C in 5% CO₂ for 30 min. To induce NETosis, the cells were exposed to fMLP (1 µM); LPS (10 ng/ml); TNF-α (20 ng/ml); PMA (30 nM), or four different SARS-CoV Spike protein preparations (0.1 or 1 µg/ml) for 4 hr to overnight at 37°. The cultures were terminated by adding 4% PFA for 15 min. Helix NP NIR (0.1 μM; BioLegend) and DAPI (0.3 nM; BioLegend) were added to detect NETs. Data acquisition (Helix NP NIR⁺ DAPI⁺ Cells) was done on FACSCelesta SORP (BD) flow cytometer and analyzed with FlowJo software (Tree Star). In vitro imaging of NETosis was performed using a modified protocol from a previous report (Hoppenbrouwers et al., 2017). The A549 cells were plated at 48 hr before imaging at 60% cell confluent. Fluorescently labeled SARS-CoV-2 Spike protein (Alexa Fluor 488; 0.5 μg/ml) was added to the A549 cells 24 hr before human neutrophil seeding. Human neutrophils were purified from whole blood by EasySep Direct Human Neutrophil Isolation Kit (Stem Cell Technologies). Purified human neutrophils were stained with Hoechst prior to seeding on SARS-CoV-2 Spike protein-pretreated A549 cells in propidium iodide (PI, 1 μg/ml) containing culture media. Time-lapse images were acquired with a Leica SP8-inverted five-channel confocal microscope (Leica Microsystems) equipped with 40× oil objective, 0.95 NA (immersion medium used distilled water). The temperature of air (5% CO₂) was maintained at 37.0 ± 0.5°C. Time interval between frames was set as 10 min for 5 hr acquisition. The loss of nucleus of human neutrophils was detected by the loss of Hoechst signal and extracellular DNA released by NETosis was labeled by PI signals.

Fluorescent nanobead binding assay of SARS-CoV-2 Spike protein

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FluoSpheres NeutrAvidin-Labeled Microspheres, 0.2 µm, yellow-green fluorescent (505/515), 1% solids (Cat# F8774, Thermo Fisher Scientific Inc) was used as a nanobead platform for SARS-CoV-2 Spike protein conjugation. Fluorescent nanobead was directly counted and measured by BD FACSCelesta flow cytometer. A flow cytometer equipped with FSC detector (Photodiode with 488/10 BP filter) and SSC detector (PMT with 488/10 BP filter) was tuned to detect voltages up to 550 for FSC and 230 for SSC. Million counts of nanobeads were conjugated with 0.5 μg of SARS-CoV-2 Spike protein by interaction of Neutravidin on beads and Strep Tag II on recombinant proteins. Nanobeads and SARS-CoV-2 Spike proteins in PBS were coupled at room temperature for 1 hr and washed with 1 ml of PBS. The coupled nanobeads were spun down with a benchtop centrifuge at a speed of 20,000 ×g, at 4°C for 20 min. The nanobeads’ pellet was suspended with PBS concentration at 2×10⁴/μl. SARS-CoV-2 Spike protein couplings to nanobeads were tested with Alexa Fluor 647-conjugated SARSCoV-2 Spike neutralizing antibody (Cat# 40591-MM45, SinoBiological). To test the binding ability of various recombinant proteins (hACE2, human Siglec-5, human Siglec-8, mouse ACE2, and mouse Siglec-F) were directly conjugated with Alexa Fluor 647. SARS-CoV-2 Spike protein-conjugated nanobeads (2×10⁴ counts) in 20 μl of 1× HBSS (contains 1 mM Ca²⁺, 2 mM Mg²⁺, 1 mM Mn²⁺, and 0.5% fatty acid-free BSA) buffer were incubated with 0.2 μg of fluorescent recombinant proteins at room temperature for 30 min. Fluorescent antibody or recombinant protein biding on fluorescent nanobeads was directly counted and measured by BD FACSCelesta flow cytometer.

Thick section immunohistochemistry and confocal microscopy

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Immunohistochemistry was performed using a modified method of a previously published protocol (Park et al., 2015; Park et al., 2018). Briefly, freshly isolated lungs were fixed in newly prepared 4% paraformaldehyde (Electron Microscopy Science) overnight at 4°C on an agitation stage. Fixed lungs were embedded in 4% low melting agarose (Thermo Fisher Scientific) in PBS and sectioned with a vibratome (Leica VT-1000 S) at a 50 μm thickness. Thick sections were blocked in PBS containing 10% fetal calf serum, 1 mg/ml anti-Fcγ receptor (BD Biosciences), and 0.1% Triton X-100 (Sigma) for 30 min at room temperature. Sections were stained overnight at 4°C on an agitation stage with Ly6G, Siglec-F, CD169, CD31, and F4/80, and labeled WGA. Stained thick sections were microscopically analyzed using a Leica SP8 confocal microscope equipped with an HC PL APO CS2 40× (NA, 1.30) oil objective (Leica Microsystem, Inc) and images were processed with Leica LAS AF software (Leica Microsystem, Inc) and Imaris software v.9.9.1 64× (Oxford Instruments plc). The intensities of fluorescent signals in regions of interests were measured by LSA AF Lite software (Leica Microsystem).

In vitro imaging of HEK293 cells

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HEK293 cells were transfected using TransIT-293 transfection reagent (Mirus Bio LLC). For transient expression of human Siglec-5-GFP, 80% confluent HEK293 in eight-well chamber slide (0.25 ml/well) were transfected by adding dropwise to each well 25 μl containing 0.25 μg of plasmid diluted in Opti-MEM with a 1:4 (DNA/reagent) dilution of TransIT-293 Transfection Reagent. To generate a stable cell line of human Siglec-5-GFP and hACE2-OFP, transfected HEK293 cells were sorted with FACS Aria II by GFP and OFP signals. Sorted human Siglec-5-GFP transfected HEK293 cells were further selected with G418 (0.8 mg/ml) containing growth media and maintained with G418 (0.5 mg/ml) media. Sorted hACE2OFP transfected HEK293 cells were further selected with hygromycin B (0.2 mg/ml) containing growth media and maintained with hygromycin B (0.1 mg/ml) media. Transiently transfected HEK293 cells in eight-well chamber slide were directly imaged with a Leica SP8-inverted five-channel confocal microscope (Leica Microsystems). HEK293 cells, HEK293 Siglec-5-GFP, and HEK293 ACE2-OFP cell lines were plated at a ratio of 1:1:1 (cell numbers) in eight-well chamber with normal media 18 hr prior to imaging. Imaging was performed with a confocal microscope equipped with 40× oil objective, 0.95 NA (immersion medium used distilled water). The temperature of air (5% CO₂) was maintained at 37.0 ± 0.5°C. Fluorescent SARS-CoV-2 Spike protein (Alexa Fluor 647) (1 μg/ml) was added into culture media. Images were acquired with Leica LAS AF software (Leica Microsystem, Inc) and processed with Imaris software v.9.9.1 64× (Oxford Instruments plc).

Time-lapse imaging of lung sections with confocal microscopy

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Lung slices were obtained from mouse lungs using a slightly modified published protocol (Pieretti et al., 2014). Briefly, C57BL/6 mice were euthanized with overdose of Avertin (1 ml of 2.5% Avertin). The peritoneum was opened, and the descending aorta cut allowing blood to pool in the abdomen. The trachea was cannulated, and the lungs inflated with 37°C 1.5% low-melting-point agarose (Cat# 50111, Lonza) prepared with RPMI 1640 media. Subsequently, the lungs were excised and rinsed with RPMI 1640 media. Isolated left lungs were sectioned into six to eight 1-mm-thick transverse slices using a #10 scalpel blade. Lung slices were placed in a pre-warmed cover glass chamber slide (Nalgene, Nunc) under a metal flat washer (M8-5/16th inches diameter). The chamber slide was then placed into the temperature control chamber on the microscope. The temperature of air was monitored and maintained at 37.0 ± 0.5°C for 5% CO₂. Mounted lung sections on the chamber slide were microscopically analyzed using a Leica SP8 confocal microscope equipped with an HC PL APO CS2 40× (NA, 1.30) oil objective (Leica Microsystem, Inc) and images were processed with Leica LAS AF software (Leica Microsystem, Inc). Lung slices were imaged a range of depths (10–50 μm). Time-lapse images were processed with Imaris software v.9.9.1 64× (Oxford Instruments plc).

Intravital imaging

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The microanatomy of liver, spleen, heart, Peyer’s patch, and inguinal LN were delineated by tail vein injection of labeled antibodies before imaging. Antibodies used included CD31, blood vessels; F4/80, Kupffer cells and macrophages; CD169, subcapsular macrophages; and Ly6G; neutrophils. The antibody mixtures were injected 10 min before starting animal surgery. Fluorescently labeled Spike proteins were injected intravenously or at the mouse tail base as indicated. To image the liver or spleen a slightly modified published protocol was used (Matsumoto et al., 2018). Briefly, after initial anesthesia (Avertin 300 mg/kg, i.p.) the skin and peritoneum were cut to expose the left lobe of the liver, or the left flank was cut below the costal margin to expose the spleen. The visible organs were glued with n-butyl cyanoacrylate to a custom-made metal holder. After attachment, the mouse was placed over a pre-warmed cover glass (Brain Research Laboratories) window on universal mounting frame AK-Set (PECON). The exposed organs were kept moist with saline wetted gauze. The mounting frame was placed into the temperature control chamber on the microscope and maintained at 37.0 ± 0.5°C. Once stabilized onto imaging stage/insert the mice received isoflurane (Baxter; 2% for induction of anesthesia, and 1–1.5% for maintenance, vaporized in an 80:20 mixture of oxygen and air). For liver and spleen four-dimensional analysis of cell behavior, stacks of various numbers of section (z-step = 3, 5) were acquired every 5–30 s to provide an imaging volume of 20–50 μm in depth. Intact organ imaging of the heart, inguinal lymph node, or Peyer’s patches was performed after mouse sacrifice using same imaging procedure as used for liver/spleen imaging.

To image neutrophils in the cremaster muscle (Yan et al., 2021), mice received an intrascrotal injection of PBS, BSA, IL-1β (50 ng in 300 µl saline, R&D Systems), Spike proteins, or IL-1b and Spike proteins. 90 min prior to imaging, the mice received injections of Avertin (300 mg/kg, i.p.) and fluorescently labeled antibodies intravenously. Antibodies used directed against Gr-1, neutrophils; GPIbβ, platelets; and CD31, blood vessel endothelium. The isolated cremaster tissue was exteriorized and stabilized onto the imaging stage/insert with the tissue directly contacting the cover glass. The exposed tissue was kept moist with pre-warmed saline (37°C). Once stabilized onto an imaging stage/insert the mouse received isoflurane (Baxter; 2% for induction of anesthesia, and 1–1.5% for maintenance, vaporized in an 80:20 mixture of oxygen and air), and placed into a temperature-controlled chamber. Image stacks of optical sections (z-step = 1) were routinely acquired at 20–30 s intervals to provide an imaging volume of 15–25 µm. All imaging was performed with a Leica SP8-inverted five-channel confocal microscope (Leica Microsystems) equipped with HC PL APO CS2 40× (NA, 1.30) oil objective. Sequences of image stacks were transformed into volume-rendered four-dimensional videos using Imaris software v.9.9.1 64× (Oxford Instruments plc). Video editing was performed using Adobe Premiere Pro 2022 (Adobe Systems Incorporated).

Quantifications and statistical analyses

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All experiments were performed at least three times. Representative images were placed in figures. Primary image data which was analyzed and calculated by Leica LAS AF software (Leica Microsystem, Inc) or Imaris software v.9.9.1 64× (Oxford Instruments plc). was acquired and processed with Microsoft Excel software. Error bars with ± SEM, and p values were calculated with unpaired t-test or two-way ANOVA multiple comparisons in GraphPad Prism 9.3.1 (GraphPad software). p<0.05 was considered significantly different.

Figure 1—figure supplement 2—source data 1, Figure 2—source data 1,2, Figure 2—figure supplement 1—source data 1, Figure 3—source data 1, Figure 3—figure supplement 1—source data 1, Figure 4—figure supplement 1—source data 1, Figure 5—source data 1,2, Figure 7—source data 1–3, and Figure 8—source data 1 contain the numerical data used to generate the figures. Primary imaging files are available as Figure 1—source data 1–6, Figure 1—figure supplement 2—source data 2,3, Figure 3—source data 2, Figure 4—figure supplement 3—source data 1, and Figure 7—source data 4.

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Author details

1. Chung Park

   B-Cell Molecular Immunology Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, United States

   Contribution

   Conceptualization, Resources, Data curation, Formal analysis, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-7819-5333

2. Il-Young Hwang

   B-Cell Molecular Immunology Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-4498-4382

3. Serena Li-Sue Yan

   B-Cell Molecular Immunology Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, United States

   Contribution

   Data curation, Formal analysis, Methodology

   Competing interests

   No competing interests declared

4. Sinmanus Vimonpatranon

   1. Immunopathogenesis Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, Bethesda, United States
   2. Department of Retrovirology, Armed Forces Research Institute of Medical Sciences – United States Component, Bangkok, Thailand

   Contribution

   Resources, Methodology

   Competing interests

   No competing interests declared

5. Danlan Wei

   Immunopathogenesis Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, Bethesda, United States

   Contribution

   Resources

   Competing interests

   No competing interests declared

6. Don Van Ryk

   Immunopathogenesis Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, Bethesda, United States

   Contribution

   Resources, Methodology

   Competing interests

   No competing interests declared

7. Alexandre Girard

   Immunopathogenesis Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, Bethesda, United States

   Contribution

   Resources, Methodology

   Competing interests

   No competing interests declared

8. Claudia Cicala

   Immunopathogenesis Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, Bethesda, United States

   Contribution

   Conceptualization, Resources, Methodology

   Competing interests

   No competing interests declared

9. James Arthos

   Immunopathogenesis Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, Bethesda, United States

   Contribution

   Conceptualization, Resources, Methodology

   Competing interests

   No competing interests declared

10. John H Kehrl

    B-Cell Molecular Immunology Section, Laboratory of Immunoregulation, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, United States

    Contribution

    Conceptualization, Supervision, Funding acquisition, Investigation, Writing - original draft, Project administration, Writing - review and editing

    For correspondence

    jkehrl@niaid.nih.gov

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-6526-159X

Funding

No external funding was received for this work.

Acknowledgements

The authors thank Dr. Anthony Fauci for long-standing encouragement. This work was supported by Intramural Research Program of National Institute of Allergy and Infectious Diseases.

Ethics

This study strictly adhered to the recommendations in the Guide for the Care and Use of Laboratory Animals by the National Institutes of Health. All animals were handled according to approved institutional animal care and use committee (IACUC) protocols (LIR-15) of the National Institutes of Allergy and Infectious Disease. The protocol received approval from the Committee on the Ethics of Animal Experiments of the National Institutes of Allergy and Infectious Disease. All surgeries were performed under anesthesia, with every effort made to minimize suffering.

Version history

1. Sent for peer review: March 13, 2023
2. Preprint posted: March 14, 2023 (view preprint)
3. Preprint posted: June 6, 2023 (view preprint)
4. Preprint posted: February 15, 2024 (view preprint)
5. Version of Record published: March 20, 2024 (version 1)
6. Version of Record updated: April 2, 2024 (version 2)

Cite all versions

You can cite all versions using the DOI https://doi.org/10.7554/eLife.86764. This DOI represents all versions, and will always resolve to the latest one.

Copyright

This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.

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