Regular Article
Investigating virus–host cell interactions: Comparative binding forces between hepatitis C virus-like particles and host cell receptors in 2D and 3D cell culture models

https://doi.org/10.1016/j.jcis.2021.02.067Get rights and content

Abstract

Cell cultures have been successfully used to study hepatitis C virus (HCV) for many years. However, most work has been done using traditional, 2-dimensional (2D) cell cultures (cells grown as a monolayer in growth flasks or dishes). Studies have shown that when cells are grown suspended in an extra-cellular-matrix-like material, they develop into spherical, ‘organoid’ arrangements of cells (3D growth) that display distinct differences in morphological and functional characteristics compared to 2D cell cultures. In liver organoids, one key difference is the development of clearly differentiated apical and basolateral surfaces separated and maintained by cellular tight junctions. This phenomenon, termed polarity, is vital to normal barrier function of hepatocytes in vivo. It has also been shown that viruses, and virus-like particles, interact very differently with cells derived from 2D as compared to 3D cell cultures, bringing into question the usefulness of 2D cell cultures to study virus–host cell interactions. Here, we investigate differences in cellular architecture as a function of cell culture system, using confocal scanning laser microscopy, and determine differences in binding interactions between HCV virus-like particles (VLPs) and their cognate receptors in the different cell culture systems using atomic force microscopy (AFM). We generated organoid cultures that were polarized, as determined by localization of key apical and basolateral markers. We found that, while uptake of HCV VLPs by both 2D and 3D Huh7 cells was observed by flow cytometry, binding interactions between HCV VLPs and cells were measurable by AFM only on polarized cells. The work presented here adds to the growing body of research suggesting that polarized cell systems are more suitable for the study of HCV infection and dynamics than non-polarized systems.

Introduction

Since infectious hepatitis C virus (HCV) was produced in cell culture for the first time in 2005 [1], [2], [3], traditional monolayer (2-dimensional) cell cultures (i.e. growth on the surface of a cell culture flask or dish) have been used extensively to study HCV, and to investigate virus–host cell interactions [4]. However, the extracellular environment to which cells in culture are exposed has been shown to have a profound impact on cell biology, influencing features such as differentiation, development of signaling pathways, and development of multicellular structures [5]. When grown under certain conditions, cells in culture have been shown to form multicellular spheroid structures – organoids – that more closely resemble the in vivo cellular architecture and function of the organs from which they were derived, and hence differ greatly from monolayer cultures. For example, a key difference between monolayer (2D) and organoid (3D) hepatocyte cell cultures is that, in 3D cultures, cells become polarized, with distinct apical and basolateral surfaces separated and maintained by tight junctions between adjacent cells, as seen in hepatocytes in vivo [6]. This polarization is important for the correct functioning of hepatocytes. Indeed, the loss of polarization in hepatocytes in vivo has been linked to oncogenesis [7]. Further, polarization in cell culture systems leads to significantly different virus–cell interactions as compared to those in non-polarized cells [6], [8], [9]. Organoid cultures are proving to be an invaluable tool in the study of pathogenesis. For example, primary human liver organoids have been utilized as a model system to study hepatitis B virus (HBV) infection and resultant tumorigenesis, as well as for virus production and the study of HBV transcription [10], [11], while organoid cultures of biopsy-derived liver cancer tissue may allow for the development of personalized therapies [12]. Primary human liver organoids have further been investigated as a means to generate healthy mature hepatocytes for engraftment [13], [14]. Similarly, primary intestinal endothelial organoids have been used to investigate cystic fibrosis [15] and colorectal cancer, working towards personalized medicine [16], and to model rotavirus [17], Salmonella [18] and Helicobacter [19] infection.

The human hepatocarcinoma cell line, Huh7, is commonly used to study HCV pathogenesis. Monolayer cultures of Huh7 cells are known to be non-polarized [20], and are therefore potentially a less-than-ideal system to investigate HCV. However, under certain growth conditions, Huh7 cells have been shown to form organoids. These organoid cultures more closely mimic conditions in vivo than do monolayer cell cultures [21]. In recent work, organoid cultures of Huh7 cells grown in suspension in extracellular matrix (ECM) were seen to form structures resembling bile canaliculi (BC), and to display polarization with clearly defined apical and basolateral surfaces. Adjacent cells were connected through tight cell–cell interactions (tight junctions), defining and maintaining polarity of cells. Further, key markers (Na+K+-ATPase, Zonula Occuldins-1 (ZO-1) and Radixin) were seen to display clearly polarized localization. Additionally, these organoids were seen to perform in vivo-like functions, such as the export of a labeled bile analogue to the canalicular space; behavior absent in monolayer cell cultures of Huh7 cells [21]. The organoid cultures were also shown to support HCV infection at levels equivalent to that of 2D Huh7 cultures.

Hepatitis C virus has been shown to display extremely restricted tropism, almost exclusively infecting hepatocytes. This is partly due to the complex attachment and entry pathway of HCV, and the absolute requirement for specific cell receptors. HCV entry into host cells requires the tetraspanin molecule CD81 and the scavenger receptor class B type 1 (SR-B1), which bind the HCV envelope glycoprotein E2 [22], [23], [24]. Endothelial growth factor receptor (EGFR) has also been shown to be vital in the assembly of the HCV receptor complex [25], [26]. In 2D Huh7.5 cells, Coller et al. [27] demonstrated that SR-B1, CD81, and the tight junction proteins Claudin-1 (CLDN1) and Occludin (OCLN) are essential for HCV entry to hepatocytes. How the virus is able to access tight junction proteins in vivo was, however, for many years a matter of debate. The requirement for CLDN1 and OCLN for HCV entry has been well documented in various siRNA and knockout experiments [28], [29], [30], [31], [32], [33]. However, despite earlier experiments indicating that cytoskeletal re-arrangements could traffic HCV–CD81–SR-B1 complexes to tight junctions to allow for interaction with tight-junction-associated proteins CLDN1 and OCLN [34], Coller et al. [27] did not see evidence of HCV entry occurring at tight junctions in monolayer Huh7.5 cell cultures. The exact role of these tight-junction-associated proteins was not elucidated until very recently, when in 2018 Glen Randall’s group (Baktash et al. [6]) imaged HCV entry into Huh7.5 organoid cell cultures. They demonstrated that host cell entry by HCV virions proceeds in a complex and ordered fashion, involving initial association, at the basolateral membrane, with SR-B1, CD81 and EGFR. The complex is then trafficked to the tight junctions in an actin-dependent manner and, upon interaction with CLDN1 and OCLN within the tight junctions, is internalized via clatherin-dependent endocytosis. CD81 was shown to be essential for the trafficking of HCV to the tight junctions to allow subsequent internalization. In 3D cells, EGFR was found to be essential for triggering internalization of CLDN1–HCV receptor complexes (once migration to the tight junctions had occurred), but not for initial interaction between CLDN1 and CD81 [6], as was earlier determined in monolayer cell cultures [35], highlighting the importance of organoid cultures in the study of HCV pathogenesis.

The binding of a ligand to its cognate receptor can be measured as adhesive forces, i.e. the amount of ‘pulling’ force required to dissociate the bond is indicative of the binding affinity. Determination of biological adhesion forces by Atomic Force Microscopy (AFM) is an established technique; it has been more than two decades since Lee et al. [36] first used the technique to determine the strength of the biotin–streptavidin bond, with a rupture force of 1 nN required to dissociate the bond – placing the bond strength between that of van der Waals forces and an ionic bond – and Hinterdorfer et al. [37] reported that a force of 240 nN was required to dissociate a single antibody-antigen complex. Functionalized AFM experiments involve attaching specific molecules to AFM probes to determine interaction forces between the probe molecule (ligand) and its receptor(s). The technique has been used to investigate adhesion molecules involved in cell–cell adhesion in mammalian cells [38], [39], and various characteristics of bacterial cells [40]. Recently, the development of functionalized AFM protocols allowing single-cell force spectroscopy (CSFS) and single-molecule force spectroscopy (SMFS) measurements to be taken have led to observations such as the rearrangement of cellular receptors in response to both chemical and mechanical signals [41]. More recently, protocols have been developed for the functionalization of AFM tips with viruses, allowing virus binding to cognate receptors on living cells to be investigated [42], [43]. Newton et al. have described refined protocols utilizing the technique, combined with confocal microscopy, to determine binding forces between enveloped viruses and live cells expressing the virus’s cognate receptor [44]. However, these experiments have, to date, been carried out on model systems, utilizing cells engineered to express certain receptors, in combination with model viruses engineered for ectopic expression of exogenous viral proteins.

iIn this work, we present the first investigation of binding of HCV VLPs to a hepatocyte cell line, utilizing organoid cell cultures which closely mimic the native target cells of HCV. Confocal scanning laser microscopy (CSLM) was used to visualize differences between organoid and monolayer culture cells, with localization of key cell surface proteins indicating polarization of organoid cells, while no polarization was observed for monolayer culture cells. HCV VLP functionalized AFM cantilever tips were used to differentiate the spatial arrangement of binding of the VLPs to the hepatic cells in situ. Interestingly, distinct binding interactions were only observed for the polarized cells, whereas traditional, 2D cultured cells displayed negligible VLP interactions. These binding interactions, as measured by atomic force spectroscopy, were completely inhibited by the introduction of anti-HCV VLP IgG antibodies. We therefore investigated blocking of binding and uptake of HCV VLPs by HCV VLP antiserum and polyclonal IgG purified from serum from pigs inoculated with an HCV VLP-based vaccine candidate [45] in polarized and non-polarized cells using flow cytometry. We found both serum and IgG blocked binding and uptake of HCV VLPs in both organoid and monolayer cultured Huh7 cells. Therefore, differences in HCV VLP–host cell interactions as measured by atomic force spectroscopy are likely indicative of differences in the formation of the initial cell-VLP attachment complex between 2D and 3D cell cultures.

Section snippets

VLP production

HCV genotype 1a VLPs were produced by expression of HCV core, E1 and E2 proteins in Huh7 cells transfected with a recombinant adenovirus vector and purified by ultracentrifugation as previously described [46], [47]. For VLP labeling with fluorescein isothiocyanate (FITC; Sigma USA. F7250, ≥90%), 20 µg of VLP was incubated overnight at 4 °C in 200 µL 2 mg/mL FITC in 100% dimethyl sulfoxide (DMSO, Sigma, 99%), then dialyzed overnight at 4 °C, in 100 kDa cutoff dialysis tubing, against 1 L of

Generation of polarized Huh7 organoids

Huh7 cells were grown as organoids and investigated using immunofluorescence techniques (Fig. 1, Fig. 2). In vivo, hepatocytes are highly polarized cells forming continuous networks of tube-like structures with external, or basal, cell surfaces in contact with the hepatic blood supply and the apical cell surfaces giving structure to the internal lumen into which bile is secreted – the bile canaliculi. These basal and apical surfaces are defined and maintained by tight junctions between adjacent

Discussion

We present here the first report of AFM measurements of binding interactions between HCV VLPs and the hepatocyte cell line, Huh7. We found that binding interactions were only discernable between HCV VLPs and cells that had been grown suspended in extracellular matrix, i.e. polarized organoid cultures. We also found that when adhesive forces could be measured, they were localized in distinct microdomains on cells. In work conducted on various coronaviruses (CoV), Hantak et al. [54] have recently

Conclusion

Here, we report the generation of organoid cultures that were polarized, as determined by localization of key cellular markers, and found that binding interactions between HCV VLPs and cells, as measured by atomic force spectroscopy, were measurable only on polarized cells. Adhesive forces in the range of 400–800 pN were recorded, with adhesion events seen to be clustered into microdomains. These binding forces were 1–2 orders of magnitude higher than previously reported values [57], [58], most

CRediT authorship contribution statement

Simon Collett: Data curation, Writing - original draft, Software, Visualization. Joseph Torresi: Funding acquisition, Project administration, Writing - original draft, Software, Visualization. Linda Earnest-Silveira: Data curation, Writing - original draft, Software, Visualization. Vi Khanh Truong: Data curation, Visualization. Dale Christiansen: Data curation, Writing - original draft, Software, Visualization. Bang M. Tran: Data curation, Visualization. Elizabeth Vincan: Data curation,

Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgements

This research was performed in part at the RMIT Micro Nano Research Facility (MNRF) in the Victorian Node of the Australian National Fabrication Facility (ANFF). We acknowledge the facilities, and the scientific and technical assistance of the RMIT Microscopy & Microanalysis Facility (RMMF), a linked laboratory of Microscopy Australia. S.C. is supported by a research training program stipend scholarship from the Australian Government, Department of Education and Training. A.E. acknowledges

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