Single influenza A viruses induce nanoscale cellular reprogramming at the virus-cell interface

Broich, Lukas; Wullenkord, Hannah; Osman, Maria Kaukab; Fu, Yang; Müsken, Mathias; Reuther, Peter; Brönstrup, Mark; Sieben, Christian

doi:10.1038/s41467-025-58935-8

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Published: 25 April 2025

Single influenza A viruses induce nanoscale cellular reprogramming at the virus-cell interface

Nature Communications volume 16, Article number: 3846 (2025) Cite this article

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Abstract

During infection, individual virions trigger specific cellular signaling at the virus-cell interface, a nanoscale region of the plasma membrane in direct contact with the virus. However, virus-induced receptor recruitment and cellular activation are transient processes that occur within minutes at the nanoscale. Hence, the temporal and spatial kinetics of such early events often remain poorly understood due to technical limitations. To address this challenge, we develop a protocol to covalently immobilize labelled influenza A viruses on glass surfaces before exposing them to live epithelial cells. Our method extends the observation time for virus-plasma membrane association while minimizing viral modifications, facilitating live imaging of virus-cell interactions. Using single-molecule super-resolution microscopy, we investigate virus-receptor interaction showing that viral receptors exhibit reduced mobility at the virus-binding site, which leads to a specific local receptor accumulation and turnover. We further follow the dynamics of clathrin-mediated endocytosis at the single-virus level and demonstrate the recruitment of adaptor protein 2 (AP-2), previously thought to be uninvolved in influenza A virus infection. Finally, we examine the nanoscale organization of the actin cytoskeleton at the virus-binding site, showing a local and dynamic response of the cellular actin cortex to the infecting virus.

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Introduction

Influenza A viruses (IAVs) are a major human health threat that cause annual epidemics and sporadic pandemics¹. IAVs are enveloped particles, and their membrane harbors the two major viral envelope proteins, hemagglutinin (HA) and neuraminidase (NA). To initiate viral replication and ensure the production of progeny virions, IAVs need to infect susceptible host cells. The infection process begins with the binding of the viral HA to sialylated glycan receptors on the host cell plasma membrane². While this initial interaction is critical for IAV host tropism¹, it is of inherently low affinity³. We could recently show that IAVs overcome this limitation by forming multivalent contacts through the binding of receptor nanoclusters within the plasma membrane⁴. However, subsequent steps of the infection process remained elusive at the single-virus level.

Following receptor binding, IAVs enter cells by clathrin-mediated endocytosis (CME) or macropinocytosis^5,6. For CME-based cell entry, IAVs were shown to use pre-existing clathrin-coated pits or induce the de novo formation of new endocytosis sites, suggesting a specific signaling function into the cell⁷. Since sialylated receptors do not possess any signaling capacity, glycoproteins were hypothesized early on, and more recently a number of candidates have been proposed to be involved in IAV cell entry^8,9,10,11,12. Among them are tyrosine kinases such as the epidermal growth factor receptor (EGFR) or c-Met¹¹. siRNA-mediated knock-down of EGFR in susceptible cell lines could reduce the infection rate of human H1N1 IAV strain A/Puerto Rico/8/34 (PR8) by about 60%¹¹. Co-localization on the plasma membrane could also be shown, suggesting a direct interaction between IAV and EGFR^4,11. We have recently performed single-virus tracking experiments to better understand how IAVs engage with functional EGFR nanodomains. Our results suggested that IAVs explore the plasma membrane in a receptor-dependent way, thereby eventually reaching plasma membrane nanodomains that harbor both sialylated receptors and EGFRs, which then become activated, leading to downstream signal transduction and induction of CME⁴. However, the spatial and temporal visualization of this process at the single-virus level remained difficult due to the transient nature of the cell entry process and the small size of the IAV-cell interface.

In this work, we present a method to stabilize the interaction between viruses and cells, allowing for a comprehensive examination of the dynamics at the virus-cell interface, including receptor recruitment and the subsequent induction of clathrin-mediated endocytosis (CME). We developed a protocol to modify microscopy glass substrates for the immobilization of unmodified IAVs, thereby generating a stable viral interface that can then be brought in contact with live cells for microscopic investigation. With our method, we were able to prolong the plasma membrane contact between virus and cells to investigate the interaction between IAVs and individual EGFR molecules, using advanced single-molecule localization microscopy (SMLM) and single-particle tracking. By determining the mobility of individual receptors, we could identify a significant decrease of receptor diffusion in proximity to IAV, indicating a direct interaction between receptor and virus. In subsequent experiments, we show the induction of CME as well as the dynamics and nanoscale response of the actin cytoskeleton at the virus binding site. Taken together, our approach enables a nanoscale view of IAV-induced cellular reprogramming at the virus-cell interface.

Results

Unmodified IAVs can be covalently linked to glass substrates before live cell attachment

To generate a stable virus-cell interface enabling high-resolution visualization of virus-cell interaction, we chemically modified glass coverslips for the covalent immobilization of native viruses and the attachment of live cells. The coverslips were coated with a silane-PEG₅₀₀₀-NHS linker that could further react with primary amines present on the viral envelope proteins. To passivate the regions between the viruses and reduce the amount of NHS-linkers, silane-PEG₅₀₀₀-NHS was mixed with silane-PEG₂₀₀ at a ratio of 1:1 (Fig. 1a). We immobilized fluorescently-labeled IAV PR8 particles and quantified the number of attached particles per 500 µm², which corresponds to the projected 2D surface area of an attached A549 human lung epithelial cell. In parallel, we performed immunostaining and transmission electron microscopy of our fluorescently labeled IAVs to evaluate the quality of our virus preparation. The majority of DID-labeled particles stained positive for H1 and also contained viral nucleoprotein (NP), confirming the presence of intact and H1 decorated virions (Supplementary Fig. 1A, B). In addition, transmission electron microscopy of our virus preparation revealed that it consists almost exclusively of particles resembling pleomorphic IAVs in size and shape (Supplementary Fig. 1C). The number of immobilized virus particles and their integrated intensity were then determined by confocal microscopy. Representative images and the quantification of viruses detected per area are shown in Fig. 1b. In comparison to an unreactive PEG-passivated slide, we found a 2.5-fold higher number of viruses on the NHS-modified slide, with very few particles being attached to silane-PEG₂₀₀ coated slides (Fig. 1b, violin plot). We then investigated the intensity distribution of the immobilized IAV particles, which showed a single peak that overlaps with the intensity distribution of 100 nm fluorescent beads, immobilized using the same approach and recorded in a parallel experiment. Representative images and the intensity quantification are shown in Fig. 1c. This indicates that virus particles did not aggregate during the labeling and immobilization procedure despite the pleomorphism of IAVs^13,14. Together, this one-step silanization protocol provides a homogenously distributed viral attachment pattern on the microscopy glass slides.

The next step was to enable the attachment of live mammalian cells on top of the immobilized IAVs. To aid live cell attachment, we conjugated a small cyclic pentapeptide with an arginine-glycine-aspartate motif (Cyclo(-RGDfK), further referred to as cRGD), a subunit of the extracellular integrin-binding matrix protein fibronectin¹⁵, to the silane-PEG₅₀₀₀-NHS linker. We then analyzed the time course of cell adhesion using fluorescently labeled A549 cells. Quantification of the cellular area over time showed that the cRGD coating indeed enhanced the speed of cell attachment with a saturation point of the growth curve already after 1 h. Cell adhesion was slower and less pronounced on untreated and silane-PEG-treated glass slides, as exemplified by the 30% smaller cellular area (Fig. 1d). To adhere both, IAV and live A549 cells to the slide simultaneously for subsequent microscopic analyses, we mixed the labeled viruses with 0.5 mM cRGD solution and immobilized both at the same time onto the slides. We then cultured A549 on the immobilized viruses and used confocal microscopy to investigate the 3D organization of the sample. Following cell fixation and staining with Alexa488-conjugated phalloidin to visualize the cellular periphery, we observed that immobilized viruses were indeed located under the cells on the same slide (Fig. 1e). We would like to add that we cannot exclude the possibility that the viruses also bind to a certain extent to cRGD in this configuration. However, IAV particles remain immobile during the experiment, are not washed away during the slide preparation, and don’t enter the cells, thereby indicating covalent attachment through NHS.

In summary, our glass modification protocol allows the immobilization of native unmodified virus particles and is compatible with the attachment of live mammalian cells. Our platform can now be used to investigate the formation and signaling function of the virus-cell interface using fluorescence microscopy.

Immobilized IAVs recruit EGFR in a sialic acid-dependent way

To successfully infect a cell, viruses need to interact with plasma membrane receptors. While it is well described that IAVs bind host cells via sialic acid receptors, the nature of other cellular proteins involved in virus entry remains less well understood. Previously, EGFR was suggested as a potential candidate glycoprotein involved in infection of IAV PR8¹¹. To investigate the role of EGFR in the infection of host cells by IAV, we immobilized fluorescently labeled IAV PR8 particles on glass surfaces and cultured A549 cells expressing endogenous levels of EGFR-GFP on top of these viruses. We then used live-cell total internal reflection (TIRF) microscopy to investigate the distribution of EGFR-GFP in proximity to immobilized viruses. We found an accumulation of the EGFR-GFP signal colocalizing with fluorescently labeled IAV particles (example in Fig. 2a). To test whether this accumulation was dependent on HA-sialic acid interactions, the cells were treated with 250 mU/ml sialidase for 1 h prior to imaging. The accumulation of EGFR-GFP was not visible anymore, indicating a sialic-acid-dependent IAV-EGFR interaction (Supplementary Fig. 2).

Virus-receptor interaction occurs within the virus-cell interface, a nanoscale patch of the plasma membrane. To gain deeper insights into the dynamic interaction between IAVs and individual EGFR molecules, we used photoactivated localization microscopy (PALM), a super-resolution microscopy method that relies on the sparse photoswitching and subsequent localization of individual photoactivatable proteins^16,17. For this purpose, A549 cells expressing EGFR tagged with mEos3.2 (FPbase ID: VUXRFA), a PALM-compatible fluorophore¹⁸, were cultured on top of immobilized IAV particles. Individual live cells were imaged for 300–600 s, and EGFR-mEos3.2 localizations determined using established single-molecule localization routines (see methods). Importantly, since we used live cells, EGFR molecules could be followed (tracked) over several frames by single-particle tracking PALM (sptPALM¹⁹), which allowed the localization of individual receptor molecules and the reconstruction of their movements with nanometer precision. On average, about 10,000 receptor trajectories were reconstructed per cell. For each EGFR trajectory, the mean squared displacement (MSD), as well as the initial diffusion coefficient (D_1-3) was calculated (see methods).

For cells grown on linker-modified slides without any immobilized viruses, we determined the median initial diffusion coefficient (D_1-3) for EGFR to be 0.051 ± 0.013 µm²/s. Notably, similar values of approximately 0.05 µm²/s or 0.048 ± 0.065 µm²/s have been reported for EGFR in A431 cells, a comparable human epithelial cell line^8,9. This indicates that the modified glass surface did not have a negative impact on the diffusion properties of EGFR.

To then investigate the effect of virus binding on the EGFR dynamics, we repeated the experiment with cells cultured on top of immobilized IAV particles. Median diffusivities were compared either at the whole cell (WC) level or for receptors found in 800 × 800 nm squares around immobilized viruses (Fig. 2b, red square). We determined the initial diffusion coefficient (D_1-3) of all EGF receptors to be 0.040 ± 0.009 µm²/s (Fig. 2c, d). This is within one standard deviation from the measurements without the virus (Fig. 2d and Supplementary Fig. 3). We thus conclude that the mere presence of the virus does not appear to alter the overall receptor diffusivity at the whole-cell level. MSD vs. lag time plots allow the discriminate different particle movement patterns by analyzing the shape of the resulting curve²⁰. Comparing MSD vs. lag time plots of receptors in viral proximity with receptors in other membrane patches (Fig. 2b, red/blue squares) showed a distinct diffusive behavior for both groups (Fig. 2c, right panel) indicating slower EGFR diffusion in viral proximity (+/− virus mobile). Indeed, when we specifically looked at receptors within membrane patches located around the viruses, we observed a significant reduction in diffusivity to 0.024 ± 0.012 µm²/s (Fig. 2d, on the virus, p = 0.0040). To again test the effect of sialic acid on the reduction of IAV-mediated EGFR slowdown, we treated the cells with sialidase before the measurement. Sialic acid digestion reversed this effect, resulting in a diffusion coefficient of 0.054 ± 0.015 µm²/s for receptors in viral proximity (Fig. 2d, on virus + sia). We further compared the fraction of slowed receptors between the different samples, considering receptors with D_1-3 < 0.01 µm²/s as slowed down^10,11. A significantly higher fraction of slowed receptors was observed in the proximity of the viruses (Fig. 2d, right panel, p = 0.0398). Complementing these results, a nanoscale cluster analysis revealed significantly larger EGFR clusters colocalizing with immobilized viruses (Supplementary Figs. 4 and 5).

Finally, we wanted to investigate the dynamic structural organization of EGFR within the virus-cell interface. Interestingly, while we observed EGFR clusters as reported previously (Supplementary Fig. 5)⁴, they appeared only transiently over time, indicating an exchange of receptors from the plasma membrane reservoir (Fig. 2e, Supplementary Fig. 6 and Supplementary Movies 1 and 2). Due to the shortness of sptPALM trajectories, EGFR movement in and out of the virus-cell interface could only rarely be observed. We thus used EGFR-GFP-expressing A549 cells and indeed observed a long-term exchange (minutes) of EGFR molecules within the virus-cell interface (Supplementary Fig. 7).

In summary, these results suggest that immobilized IAVs form a virus-cell interface capable of clustering EGFR, and this interaction depends on sialic acid. The virus-cell interface is not a static platform. Instead, receptors are slowed down, allowing a characteristic enrichment (Supplementary Figs. 5 and 8) and dynamic exchange of receptors from the larger plasma membrane reservoir.

Immobilized IAVs recruit proteins involved in clathrin-mediated endocytosis

After we could observe and quantify the interaction between IAV and host cellular EGFR, we wanted to test if the interaction induced downstream cellular signaling. The subsequent major event of IAV infection would be the induction of endocytosis, a crucial step leading to the internalization of viral particles into the host cell cytoplasm. For non-immobilized IAVs, CME has been shown to be the main endocytosis pathway. Depending on the IAV strain, the majority of viruses (~ 60%) is internalized by CME, while the remaining fraction uses clathrin-independent pathways^7,21. Further research suggested that CME-based IAV entry is dependent on the adapter protein epsin-1²².

To further assess the functionality of our assay, we cultured A549 cells expressing epsin1-GFP on top of immobilized IAV particles (Fig. 3a). The cells were imaged using TIRF microscopy under live cell conditions. We found that epsin-1-GFP displayed discrete point-like structures and that 38 ± 8% (n = 49) of immobilized IAVs colocalized with epsin1, similar to the previously published value (~ 50%) for apical IAV infection²².

**Fig. 3: Immobilized IAVs recruit proteins involved in clathrin-mediated endocytosis.**

EGFR was shown to associate with different classes of clathrin-coated pits (CCPs), which are discriminated by the presence of adapter protein 2 (AP2), where AP2-negative CCPs would rely on other endocytic adapter proteins such as eps15 and epsin-1²³. AP2 is a major constituent and pioneering factor of CCPs at the plasma membrane. To test if immobilized IAVs also recruit AP2 to the plasma membrane, we used cells stably expressing endogenous levels of AP2-RFP together with clathrin light chain A (ClC) fused to GFP. These cells were then cultured on top of immobilized IAVs and observed by live-cell TIRF imaging. We found that 63.33 ± 12.02% (n = 45) of immobilized viruses colocalized with both AP2 and ClC (Fig. 3b), confirming that IAVs indeed recruit AP2 to the plasma membrane. Interestingly, when we monitored the signal intensity over a period of 10 minutes, we detected fluctuations in the fluorescence signals for both ClC-GFP and AP2-RFP, indicating a recurring interaction between the virus and the cellular CME machinery. We identified signal peaks, quantified the average time between two intensity peaks, and found that ClC-GFP peaks had a mean interval of 62.10 ± 27.93 s, while AP2-RFP peaks could be observed every 73.94 ± 32.95 s (Fig. 3d, n = 28). Similar recruitment dynamics have been shown before for reoviruses immobilized by click-chemistry²⁴, which are in a similar size range as IAV particles.

From these experiments, we conclude that immobilized IAVs are in a continuous interaction with their host cells, with repeated formation of clathrin-coated pits at the viral binding sites. We document recruitment of epsin-1 as shown before²² but surprisingly also AP2, which was previously suggested to be dispensable for IAV infection²⁵.

Immobilized IAVs lead to local redistribution of the cellular actin cytoskeleton

The dense cortical F-actin network beneath the plasma membrane plays a central role in many cellular processes, including CME. During CME, polymerization of the globular G-actin into the filamentous F-actin was suggested to support membrane deformation and scission²⁶. While IAV cell entry was shown to depend on the actin cytoskeleton only in polarized cells²⁷, a recent phosphoproteomic study revealed a fast cellular re-programming of actin regulatory proteins upon IAV-cell attachment²⁸. We thus hypothesized that IAV-cell binding induces a local F-actin reorganization.

To investigate the F-actin organization precisely at the virus-binding site, we stained the cellular F-actin using Alexa647-conjugated phalloidin in A549 cells cultured on top of immobilized IAV particles (Fig. 4a). We imaged the cells in TIRF to visualize F-actin structures close to the plasma membrane and found that about 15% of IAVs show a distinct accumulation of F-actin (Fig. 4a and Supplementary Fig. 9). To further investigate the nanoscale organization of the F-actin accumulation in proximity of cell-bound IAVs, we performed stochastic optical reconstruction microscopy (STORM). STORM is a super-resolution microscopy method that relies on the chemically-induced photoswitching of organic dye molecules, which allows their localization with nanometer precision²⁹. STORM reconstructions revealed the complex and diverse nanoscale organization of the F-actin network within the TIRF field of adherent A549 cells (Fig. 4b). While we could detect many larger filamentous structures with diameters around 89 ± 21 nm (FWHM of 20 filaments), immobilized IAVs seem to overlap with distinct structures that we then classify based on their appearing structural organization into three groups: 1) local enrichment (i.e., blob), 2) absence of signal (i.e., hole) or 3) ring- or arc-like structures (Fig. 4b, white zoom boxes). With respect to previous reports on the nanoscale organization of F-actin during CME^30,31, we hypothesized that the observed structures (1-3) are intermediates of the same process. We thus performed live-cell imaging of F-actin at the virus-binding site. To this end, we cultured A549 cells expressing LifeAct-GFP on top of immobilized IAV particles (Fig. 4c and Supplementary Movie 3). The cells were imaged using TIRF microscopy under live-cell conditions. We observed a similar diversity of structures as observed after phalloidin staining, indicating that F-actin was efficiency labeled in both cases³². When following the LifeAct-GFP signal in the proximity of individual viruses (Supplementary Video 3), we could indeed observe fluctuations with transitions between different structures that resemble the patterns (1-3) observed by STORM. Snapshots of the LifeAct-GFP fluctuations around one virus particle (Fig. 4c, insert) are shown in Fig. 4d. Line plot measurements (Fig. 4c, insert, white line) disclosed the different F-actin patterns transitioning from a hole to a ring and into a blob that overlaps with the viral signal before turning back into a ring-like organization. Since our live-cell imaging showed a very dynamic F-actin re-organization, local enrichment at the virus binding site was short-lived and only visible in 10–20% of the frames. This observation is thus consistent with our observation in fixed cells (15% overlap, Fig. 4a) considering a larger number of viruses and assuming similar actin turnover.

**Fig. 4: Local re-organization of the actin cytoskeleton at the virus-cell interface.**

From these experiments, we conclude that immobilized IAVs induce local F-actin re-organization that can be imaged in live cells or investigated by STORM. Our findings highlight the dynamics of the local F-actin cytoskeleton in response to an attached virus and open many possibilities towards exploring the functional implications of the observed structures.

Discussion

Virus-cell entry is a dynamic process that involves the recruitment of cellular glycoproteins and their activation, which is followed by specific downstream signaling events that eventually lead to virus endocytosis. Despite the relevance of this process for virus infection, its transient nature and the small scale of the virus-cell interface have made it difficult to study at high spatial and temporal resolution. Other strategies to immobilize virus particles have been developed before but require genetic or chemical modification of viral particles^24,33. These assays are time- and resource-intensive and may also be impractical for certain viruses as the modification of the envelope proteins might affect their ability to successfully infect a cell. We have developed a universal method to immobilize native virus particles on glass slides compatible with the subsequent cultivation of live mammalian host cells. Our method allows to investigate the early interaction between an infecting virus and its host cell at the single-virus level. Importantly, the technique has the capability to measure the interaction between a host cell and multiple viruses at the same time and is compatible with a variety of different microscopy methods, including single-molecule super-resolution microscopy. We have demonstrated the versatility of our approach by studying IAV-receptor interaction, CME induction, and local F-actin reorganization, key steps involved in IAV endocytosis.

As a model system, we chose the human IAV strain A/Puerto Rico/8/34 (PR8), which was shown before to activate and use EGFR for cell entry¹¹. We could previously show that IAVs use plasma membrane nanodomains for multivalent cell attachment and subsequent activation of EGFR⁴. Here we applied sptPALM of EGFR-mEos3.2 to investigate the dynamics of IAV-EGFR interaction in live cells. We found that IAV binding slows down EGFR within the virus-cell interface and that this reduced mobility is sialic acid-dependent. We then reasoned that an active virus-cell interface should be able to recruit downstream proteins, such as those involved in CME. We could indeed observe recruitment of epsin-1 to the virus binding site as previously shown for IAV strain X-31²². Since IAV PR8 and X-31 both activate EGFR³⁴, which itself was shown to use multiple CME pathways, we further tested the recruitment of AP2 and clathrin to the virus-binding site. Although AP2 was previously suggested not to be involved in IAV-cell entry²⁵, we found recurring recruitment of both proteins with frequencies similar to those observed before for reoviruses²⁴. From these experiments, we conclude that immobilized viruses induce CME and that this process involves both epsin-1 and AP2-dependent pathways. Our experiments demonstrate that AP2 is recruited to the virus-binding site; however, the significance of AP2 in IAV entry and its interaction with other adapter proteins requires further investigation. While we show a specific sialic acid-dependent slowdown of receptor proteins, we cannot exclude curvature-dependent CME induction independent of receptor activation, as previously shown for reoviruses and synthetic surfaces^24,35. Because most IAVs use multiple entry receptors³⁶, our assay could be applied for more detailed studies to elucidate the contribution of specific receptor proteins in IAV-cell entry. In a parallel study, we have used the inverted attachment approach to investigate the interaction between H18 IAVs and MHCII receptors. We found a more pronounced receptor slowdown as compared to the PR8-EGFR case, which was accompanied by a strong receptor clustering³⁷. This indicates nanoscale clustering as a universal mechanism of IAV-mediated receptor interaction. It is important to note that a prerequisite for sptPALM is a sparse photo-activation density, ensuring that only one photoactivated protein emits light within a diffraction-limited spot (~ 300 nm in diameter). The virus-cell interface is such a diffraction-limited spot and hence the nature of the measurement is not compatible with the direct visualization of multiple proteins at once as they come together to form a cluster. To effectively visualize the dynamic processes at the virus-binding site, we thus created time-binned movies from our sptPALM data. This approach combines multiple localizations over an extended period, enabling us to visualize the buildup of receptor nanoclusters (Supplementary Fig. 6). Lastly, we use our assay to probe the local response of the actin cytoskeleton to virus-cell binding. We observed different structural arrangements (ring, hole, blobs) that are consistent with similar structures observed in live-cell imaging. The small fraction of viruses (~ 15%) showing a distinct colocalization with Lifeact-GFP could be a result of the transient nature of the observed F-actin fluctuations. Also, while CME in yeast is strictly actin-dependent³⁸, not all CME sites in mammalian cells seem to require actin accumulation³⁹, here presumably to counteract membrane tension^27,40. We thus conclude that the observed structures are intermediates of the same process. Since viruses cannot be internalized, the actin fluctuations are recurring as we also show for the recruitment of CME-associated proteins.

In summary, super-resolution and advanced quantitative microscopy has grown into an important tool in studying IAV cell biology^{4,37,41,42,43,44,45}. We believe our method will enhance research on the processes that govern early viral infections, including receptor specificity and cellular tropism in live cells. Our results shed light on the nanoscale organization and dynamics of the virus-cell interface. Here, alternative super-resolution microscopy methods such as structured illumination microscopy (SIM) or stimulated emission depletion microscopy (STED) could be used in the future to directly visualize nanocluster dynamics⁴⁶. Moreover, because the method can easily be applied to other cargo particles, such as synthetic amine-functionalized nanoparticles, virus surrogates, or bacteria, it may find utility for multiple areas of infection research at the single-cell level.

Methods

Cell lines and culture

A549 wild-type cells (A549 WT, ATCC: CCL-185) and A549 cells endogenously expressing GFP-tagged EGFR (Sigma-Aldrich, CLL1141) were cultured and maintained in DMEM (Gibco, 10566016) supplemented with 10% FCS (Capricorn, FBS-LE-12A). A549 WT cells were modified by retroviral transduction to generate a stable cell line overexpressing EGFR-mEos3.2. To keep the selection pressure on these cells, 1.25 µg/ml puromycin (Gibco, A1113803) was added to the culture medium. MDA-MB-231 cells (provided by David Drubin, UC Berkeley) endogenously expressing ClC-GFP and AP2-RFP were cultivated in DMEM/F-12 (Gibco, 10565018) medium supplemented with 10% FCS. All cells were maintained in a humidified incubator at 37 °C and 5% CO₂ and passaged every 3 to 4 days. For passaging or seeding cells for an experiment, cells were detached using 0.5% Trypsin/EDTA (Gibco, 25300054), counted, and resuspended in fresh growth medium at the desired concentration. Prior to all measurements, cells were starved in FCS and phenol red-free medium for 1 h (Gibco, 31053028). For sialidase treatment, cells were starved in the same medium supplemented with 250 mU/ml sialidase (Clostridium perfringens, Roche, 11585886001). Plasmids and cell lines are available from the corresponding authors on request.

Influenza virus production

The viruses used for these experiments were produced in specific pathogen-free eggs from ValoBiomedia. Eggs were incubated for 10 days at 37 °C and 50–70% humidity and rotated regularly. Influenza A/Puerto Rico/8/34; H1N1 (kindly provided by Klaus Schughart, HZI) was diluted 1:1000 in sterile PBS. Egg surfaces were disinfected (100 mg/ml Povidone-iodine) and punctuated. 200 µl virus suspension was injected into the eggs, and the injection site was sealed with glue. Subsequently, eggs were incubated for 48 h at 37 °C and 50–70% humidity, without rotation. After the incubation period, eggs were stored at 4 °C overnight. Eggs were opened, and allantoic fluid was aspirated and collected. The virus content of each sample was individually assessed in a hemagglutination assay (Chicken RBC, Fiebig Animal Blood Products). Positive samples were pooled together, centrifuged at 2000g at 4 °C for 30 minutes, aliquoted, and frozen at − 70 °C.

Lentivirus production and transduction

To generate an EGFR-mEos3.2 overexpressing cell line, A549 WT cells were genetically modified by retroviral transduction. Retroviruses were produced in HEK-293T cells (ATCC: CRL-3216) cultured in a 6-well plate. When cells were at ~ 70% confluency, they were co-transfected with a lentiviral packaging plasmid (psPAX2, Addgene #12260), a lentiviral envelope plasmid (pMD2.G, Addgene #12259) and a lentiviral transfer plasmid, using Lipofectamine 2000. The transfer plasmid encodes EGFR-mEos3.2 as well as a puromycin resistance (custom construct by Genscript, pLVX-M-puro backbone, Addgene #125839). Two days post-transfection, the cell culture supernatant was harvested and cleared of cellular debris by centrifuging for 5 min at 1300 × g. The supernatant was transferred to A549 cells growing in a 6-well plate in a culture medium at ~ 30% confluency. To facilitate viral infection, the medium was supplemented with 20 µg/ml polybrene (Sigma-Aldrich, TR-1003-G). Two days post-transduction, the infection medium was removed, and cells were cultured in a growth medium supplemented with 1.25 µg/ml puromycin for selection.

Glass slide preparation

To clean and activate the glass coverslips (round 25 mm, No. 1.5, Epredia and 24 mm × 60 mm, No. 1.5, Marienfeld Superior), they were submerged in a freshly prepared piranha solution (5:1 H₂SO₄/H₂O₂) for 1 h. Subsequently, the slides were washed twice with deionized water, dried using a nitrogen stream, and stored for up to 1 month at room temperature in a desiccator under a nitrogen atmosphere. To modify the activated surface, slides were treated with different ratios of silane-PEG₂₀₀₀/silane-PEG₅₀₀₀-NHS (tebu-bio, Offenbach, Germany) dissolved in dry DMSO for 45 min at RT. As a negative control, an unreactive surface consisting of only silane-PEG₂₀₀₀ was used. Slides were washed three times in dry DMSO and twice in HEPES (0.1 M, pH 8) immediately before the subsequent reaction to avoid hydrolysis of the unstable NHS ester. As an alternative source, we also performed experiments on commercially available reactive glass slides (3D epoxy glass slides, PolyAn, Berlin).

Virus labeling and sample chamber preparation

To fluorescently label IAVs, lipid-dye conjugates (DiO, DiI and DiD, Invitrogen, V22889) were incorporated into the viral membrane. IAVs were diluted to a protein concentration of 1 mg/ml in a final volume of 50 µl PBS. 1 µl of 10 mM lipid-dye solution was added, and the suspension was shortly vortexed, spun down, and incubated in the dark at RT for at least 1 hour. To separate unbound dye from labeled viruses, the suspension was fractionated by gel filtration (NAP-5, Cytiva 17-0853-01). The absorbance of the collected fractions was measured at 280 nm together with the respective dye absorbance wavelength (488 nm, 561 nm, or 647 nm) to determine protein and dye concentrations (Nanodrop One, Thermofisher). Fractions that showed a peak at both wavelengths were collected and pooled together. To remove bigger aggregates of viruses, the viruses were mixed with 400 ml PBS and sterile filtered over a 0.22 µm filter right before immobilization. Sterile-filtered viruses were diluted 1:150 in HEPES (0.1 M, pH 8) buffer, homogenized, mixed with 0.5 mM cRGD solution, and added to the slide. Nanobeads (TetraSpeck, Invitrogen, T7279), were diluted 1:100 in HEPES before immobilization. Slides were kept at 4 °C for 16 h or for 1 h at RT. For the use of commercial reactive 3D epoxy glass slides (PolyAn, Berlin), viruses were diluted in NaHPO₄ buffer (150 mM NaHPO₄, 50 mM NaCl, pH 8.5), filtered, and added to the slide. The slides were then incubated at 4 °C for 16 h, protected from light. Here, no cRGD was required for efficient cell binding. Following sample immobilization, the remaining reactive groups were quenched using a quenching buffer (50 mM ethanolamine, 100 mM TRIS, pH 9.0, sterile filtered using 0.22 µm syringe filter). To measure multiple conditions on one slide, sticky-slide 8-well high chambers (Ibidi, 80828) were glued onto the slides. After incubation, the virus suspension was removed, and the slides were washed three times with PBS, prior to seeding the cells. Per chamber, 25,000 cells were seeded for all experiments.

Immunolabelling of immobilized viruses

DID-labeled viruses were immobilized as described above. The slides were incubated in a blocking buffer containing 0.2% BSA for 1 h at RT. For staining of NP, the buffer further contained 0.2 % Triton X-100. The slides were washed and incubated with primary antibodies in a blocking buffer for at least 1 h at RT. Anti-NP (Merck, MAB8257) and anti-H1 (Thermo Fischer, PA534929) antibodies were used at 1:500 dilution. After washing, secondary antibodies (Thermo Fischer) were diluted 1:500 in blocking buffer and added to the slides for 1 h at RT.

Cellular adherence measurement

Slides were prepared as described above. 0.05 mM cRGD (Cyclo(-RGDfK, MedChemExpress, HY-P0023) in HEPES (0.1 M, pH 8) was added and incubated for 1 h to covalently couple cRGD to the NHS moiety on the linker. Slides were washed 3x in deionized water and dried in a nitrogen stream. Hoechst/DiO stained A549 cells (density: 1.25 × 10⁵ cells/ml) were seeded on top of different slides (untreated, 20 mg/ml silane-PEG₂₀₀₀, 20 mg/ml silane-PEG₅₀₀₀-NHS with cRGD), and cell adherence (area/time) was observed over the course of 3 h. The attachment was monitored using a Keyence BZ-X800 fluorescence microscope with a stage-top incubation chamber (Tokai Hit STR), and images were taken every 10 min. The adherence of the cells was analyzed using CellProfiler⁴⁷.

SMLM and image processing

SMLM was performed on a Nikon Ti Eclipse N-STORM microscope equipped with four laser lines at 405 nm (Coherent), 488 nm (Sapphire, Coherent), 561 nm (Sapphire, Coherent) and 646 (MPB Communications). The sample was observed through a Nikon Apo TIRF 100x Oil DIC N2 NA 1.49 objective, and emitted light was detected on an Andor iXon3 DU-897 EMCCD camera (Oxford Instruments). Emitted light was directed onto the camera using a dichroic mirror and filtered through a Bright Line HC 609/64 (PALM) emission filter. We typically took between 10k (PALM) and 40k (STORM) frames at 30 ms integration time using the NIS elements software (AR 5.21.03 64-bit). Single emitter positions were localized using the software DECODE⁴⁸. Separate models were trained for STORM and sptPALM acquisitions. To obtain the microscope-specific parameters needed to train a DECODE model, z-stacks of 100 nm fluorescent beads (TetraSpeck, Thermo Fisher, T7279) were acquired with a step size of 10 nm. Z-stacks were analyzed using SMAP²¹ to determine the necessary fitting parameters. These values were used as initial parameters to train the DECODE models. Each model was trained until it reached the maximum number of epochs or the loss function did not reasonably improve anymore. Localizations were further processed using custom MatLab (MathWorks) or Python routines. Lateral sample drift was corrected via redundant cross-correlation⁴⁹ using Thunderstorm v1.3¹⁷. For clustering, we used the build-in DBSCAN function of MatLab R2020b (Mathworks). Images were rendered and cropped for visualization using Thunderstorm and Fiji⁵⁰. Plotting and statistical analysis were performed using Prism 9 (GraphPad). Image resolution was determined using Fourier Ring Correlation (FRC) and image decorrelation analysis^51,52.

sptPALM

A549 cells expressing EGFR-mEos3.2 were seeded on top of the immobilized viruses and incubated overnight until sufficiently attached to the glass surface. Prior to imaging, the growth medium was removed, and cells were starved for 1 h in the infection medium (DMEM, 0.2% BSA). For sialidase treatment, 250 mU/ml sialidase were added to the infection medium. Images were acquired in HEPES buffered DMEM (Gibco, 21063029) using a Nikon Ti Eclipse N-STORM microscope. For each measurement, 10,000 frames were acquired. A standardized procedure was use to filter the resulting single emitter positions: The initial 10% of frames (1000 frames) were discarded for all measurements as well as the 5% of localizations with highest uncertainties in x and y direction. In addition, the 2.5% dimmest and brightest localizations were discarded. From the remaining localizations, trajectories were reconstructed with the use of the TrackPy¹⁸ python library. Localizations were linked using a KDTree algorithm with a maximum search range of 0.8 pixels and a zero-frame-gap memory. The adaptive stop for solving oversized subnets was set to 0.1 with a step size multiplicator of 0.9. For further analysis, only tracks with a minimum length of 10 frames were considered. Ensemble drift xy(t) was calculated and subtracted from the tracks. Mean squared displacement (MSD, Eq. 1) was determined and plotted individually for all molecules. For all tracks, the initial diffusion coefficients (D_1-3) were calculated from the initial slope of the first three data pairs of individual MSD plots, by applying a linear fit in log space (Eq. 2)^19,20. MSD and diffusion coefficients for virus spots were calculated within an 800 × 800 nm square around an immobilized virus. Receptors with D_1-3 < 0.01 µm²/s were considered immobile^10,11.

$${MSD}\,\left(T\right)=\frac{1}{T}{\sum }_{t=1}^{T}{(x\left(t\right)-{x}_{0})}^{2}$$

(1)

$${MSD}\left(T\right)=D * 2n * T$$

(2)

Live cell CME visualization

To visualize adapter protein recruitment in live cells, A549 WT cells were transfected with Epsin1-GFP (addgene, #22228) using Lipofectamine 2000 (LifeTechnologies). One day later, the cells were detached using trypsin and subsequently cultivated on top of immobilized viruses. Live-cell imaging of AP2-RFP and ClC-GFP was performed using gene-edited MDA-MB-231 cells⁵³ (kindly provided by David Drubin). The cells were imaged using a Nikon Ti2 microscope equipped with a NikonApo TIRF 100x Oil DIC N2 NA 1.49 objective. The samples were excited using a 488 nm laser (Coherent) and emitted light was detected on a pco.edge 4.2 sCMOS camera (Excellitas). The cells were observed using TIRF illumination.

F-actin labeling and imaging

For STORM imaging of F-actin, A549 WT cells were cultivated on top of immobilized viruses. At ~ 80% confluency, the cells were fixed in 4% PFA for 20 min at RT. Samples were washed three times with PBS and blocked/permeabilized for 1 h in a blocking buffer (0.2% Triton, 0.2 % BSA in PBS). Afterward, the cells were incubated in 0.56 µM phalloidin-A647 (Thermo Fisher, A22287) solution in PBS for 2 h at RT in the dark. The samples were washed three times with PBS. STORM imaging was performed using an established photo-switching buffer made of 50 mM Tris with 10 mM NaCl, supplemented with 10% glucose, 50 mM 2-mercaptoethylamine, 0.5 mg ml⁻¹ glucose oxidase, and 40 µg ml⁻¹ catalase. All chemicals were purchased from Sigma. For live-cell imaging of F-actin structures, A549 cells were transfected with LifeAct-GFP one day before detachment using Lipofectamine 2000 (Thermo Fisher) and subsequently cultivated on top of immobilized viruses. The samples were imaged as described above for live-cell CME visualization.

Transmission electron microscopy

To visualize virus particles via the negative stain method, a thin carbon support film was floated on a droplet of virus solution to allow adsorption of particles for approx. 45 s. Afterward, the carbon was bound to a copper grid (mesh 300), washed twice on water droplets, and finally put on a drop of 4% (w/v) aqueous uranyl acetate, pH 5.0, for 45 s. The exogenous liquid was taken away with filter paper, and the grid was heat-dried on a light bulb. Samples were examined in a Zeiss Libra120 Plus transmission electron microscope (Carl Zeiss, Oberkochen, Germany) at an acceleration voltage of 120 kV at calibrated magnifications using ITEM Software (Olympus Soft Imaging Solutions, Münster).

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Example datasets are available via https://zenodo.org/records/14222289. All datasets generated or analyzed during the current study are available in the paper or are appended as supplementary data. Source data are provided in this paper.

Code availability

Newly generated, custom-written MatLab and Python scripts are available at https://github.com/christian-7/NIBI/tree/main/code_Broich_et_al or via https://doi.org/10.5281/zenodo.15005579.

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Acknowledgements

C.S. would like to thank Jennifer Fricke for excellent technical assistance. This work was supported by the Helmholtz Association (VH-NG-1526 to C.S.) and a PhD fellowship of the China Scholarship Council (CSC to Y.F.). C.S. received funding from the European Union under the Horizon Europe grant agreement No. 101191739 – Project COMBINE. Views and opinions expressed are, however, those of the author(s) only and do not necessarily reflect those of the European Union or the European Research Executive Agency (REA). Neither the European Union nor REA can be held responsible for them. C.S. and M.B. acknowledge support through the cooperativity and creativity project call (CCC, project 8) of HZI. M.K.O. is a member of the Spemann Graduate School of Biology and Medicine (SGBM). P.R. is supported by the Hans A. Krebs Medical Scientist Program of the Medical Faculty of the University of Freiburg. P.R. and M.K.O. were supported by grants from the European Research Council (ERC, NUMBER 882631—Bat Flu).

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Authors and Affiliations

Nanoscale Infection Biology Group, Helmholtz Centre for Infection Research, Braunschweig, Germany
Lukas Broich, Yang Fu & Christian Sieben
Department of Chemical Biology, Helmholtz Centre for Infection Research, Braunschweig, Germany
Hannah Wullenkord & Mark Brönstrup
Institute of Virology, Medical Center - University of Freiburg, Freiburg, Germany
Maria Kaukab Osman & Peter Reuther
Faculty of Medicine, University of Freiburg, Freiburg, Germany
Maria Kaukab Osman & Peter Reuther
Spemann Graduate School of Biology and Medicine, University of Freiburg, Freiburg, Germany
Maria Kaukab Osman
Central Facility for Microscopy, Helmholtz Centre for Infection Research, Braunschweig, Germany
Mathias Müsken
German Center for Infection Research (DZIF), Site Hannover-Braunschweig, Braunschweig, Germany
Mark Brönstrup
Institute of Organic Chemistry, Leibniz University Hannover, Hannover, Germany
Mark Brönstrup
Institute of Genetics, Technische Universität Braunschweig, Braunschweig, Germany
Christian Sieben

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Contributions

L.B., M.B., and C.S. conceived and designed the study. L.B., H.W., M.K.O., Y.F., P.R., and C.S. performed experiments and analyzed the data. M.M. performed the TEM analysis. L.B. and C.S. wrote the manuscript with input from all authors.

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Broich, L., Wullenkord, H., Osman, M.K. et al. Single influenza A viruses induce nanoscale cellular reprogramming at the virus-cell interface. Nat Commun 16, 3846 (2025). https://doi.org/10.1038/s41467-025-58935-8

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Received: 27 July 2024
Accepted: 03 April 2025
Published: 25 April 2025
DOI: https://doi.org/10.1038/s41467-025-58935-8

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