Initial descriptions of COVID-19, the pandemic disease caused by severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), characterized it as a primarily respiratory syndrome 1 . However, increasing clinical evidence now implicates multiple organ systems in COVID-19, including the heart, gastrointestinal tract, and kidneys [2] [3] [4] [5] . Notably, multiple independent reports have reported cases of acute COVID-19 associated myopathy [6] [7] [8] , even without prior cardiovascular disease 9 , indicating that SARS-CoV-2 may be directly causing cardiac damage. Meta-analyses identify elevated high-sensitivity troponin-I or natriuretic peptides, biomarkers of cardiac damage, as the strongest predictors of mortality in hospitalized patients, eclipsing both prior congestive obstructive pulmonary disease and cardiovascular disease [8] [9] [10] [11] . Alarmingly, 55% of COVID-19 patients presented abnormal echocardiograms 12 , and a majority of recovered patients continue to suffer from impaired cardiac function, indicating that long-term heart sequelae from COVID-19 may not be limited to severe cases 13 . TMPRSS2, commonly involved in viral cell entry, was not detected in any cell type, cathepsin-L (CTSL) and cathepsin-B (CTSB) were detected in all cells. These observations suggest potential infectivity of CMs by SARS-CoV-2, and predict poor infectivity in ECs and CFs. To validate these predictions, we exposed human iPSC-derived CMs, CFs, ECs, or a mixture of all three to mimic native myocardial composition, to SARS-CoV-2 at a low MOI (MOI = 0.006).
After 48 hours, CFs and ECs showed little to no viral RNA relative to a housekeeping control, whereas CMs expressed >10 4 greater levels of viral RNA than CFs or ECs (Fig. 1A) . There was no significant difference in viral detection between CMs and mixed cultures (Fig. 1A) , and undifferentiated iPSCs were uninfectable (Supp. Fig. S1D ). Differences in viral expression largely correlated with cell-type specific ACE2 expression levels (Supp. Figs. 1A-B) . Plaque assays on the supernatants of virally exposed cells confirmed that CFs, ECs, and iPSCs did not support productive infection, whereas CMs robustly produced replication-competent virions (Supp. Fig. S1E ).
Immunostaining for viral Spike protein and for double-stranded RNA (dsRNA), indicative of replicating virus, further confirmed that only CMs supported viral replication (Fig. 1B, white  arrows) . However, all three cell types exhibited significant cytopathic effects after 48 hours of viral exposure, characterized by fragmented cell bodies, dissociation from neighboring cells (Fig. 1B , red arrows) and significant cell death (Fig. 1C) . Visual cytopathic effects were most prevalent in CFs, while the greatest nuclear loss was observed in ECs, indicating toxicity from viral exposure can occur without detectable viral replication. However, inoculation with heatinactivated SARS-CoV-2 did not cause cell death or dissociation in any of the cell types assayed (Fig. 1C) .
Replication of positive-strand single stranded RNA viruses, including SARS-CoV-2, involves budding of double-membrane vesicles from the endoplasmic reticulum, with viral particle assembly occurring in cisternae of the ER-Golgi intermediate compartment (ERGIC) 32 . In CMs infected with SARS-CoV-2, dsRNA and Spike protein initially (24h post-infection) accumulated near the nucleus in small perinuclear puncta, the typical location of the ERGIC, indicating potential replication centers (Fig. 1D) . At 48h post infection, many cells exhibited dsRNA signal dispersed throughout their cytoplasm, which may correlate with advanced stages of infection. By 72h post infection, SARS-CoV-2 was spread throughout the culture and a large portion of the CMs had died, with the remaining cells displaying disperse viral stain localization, dissociation from neighboring cells, and heavily reduced sarcomeric staining (Fig. 1D) .
Remnants of the ER-Golgi membranes (light blue arrowheads) and large vesicles near the nucleus in infected CMs were readily identified by transmission electron microscopy (TEM). These vesicles, approximately 500-750 nanometers in diameter, contained multiple particles 60-100 nm in diameter that were identified as SARS-CoV-2 virions (Fig. 1E, red arrowheads) . Altogether, these results indicate that SARS-CoV-2 is able to infect, replicate in, and rapidly propagate through CMs. In all experiments, cells were exposed to SARS-CoV-2 virus for 48 hours, unless otherwise specified, at an MOI of 0.006 before lysis or fixation. A. RT-qPCR quantification of viral RNA (Fold change of SARS-CoV-2 N gene, N5, over housekeeping gene transcript, RPP30) in cell cultures exposed to SARS-CoV-2. CF: iPSC-derived cardiac fibroblasts; EC: iPSC-derived endothelial cells; CM: iPSC-derived cardiomyocytes; Mixed: 60:30:10 CM:EC:CF . Error bars: SEM. **: p-val < 0.01, one-way ANOVA with Tukey's multiple comparisons. B. Representative images of immunostaining of cardiac cells exposed to SARS-CoV-2. PECAM-1 (CD31) was used as an endothelial cell marker, and cTnT as a cardiomyocyte specific marker. Cardiac fibroblast (CF) cells expressed GFP constitutively. Viral signal was detected by staining for SARS-CoV-2 Spike protein or viral double stranded RNA (dsRNA), as noted. White boxes represent zoomed in regions in rightmost panels. White arrows denote clusters of dsRNA signal. C. Toxicity of SARS-CoV-2 to cardiac cell types, quantified by nuclear retention. Y-axis depicts the % of nuclei counted (relative to mock). Nuclei were counted automatically at 10x magnification (10 images/condition). We next sought to elucidate the mechanism of SARS-CoV-2 entry into CMs. Pretreatment of cells with an ACE2 blocking antibody or with cathepsin-B and -L inhibitor E64d significantly reduced viral detection in infected CMs ( Fig. 2A, S1F) . Though FURIN is expressed in CMs (Fig. S1A) , FURIN inhibition did not reduce infection (Fig. S1F) . Inhibiting specific cathepsins revealed that CTSL inhibition via Z-Phe-Tyr(tBu)-diazomethylketone (Z-FY-DK) decreased viral detection in infected cells to approximately 10% of vehicle-treated controls, whereas inhibition of CTSB with CA-074 did not attenuate viral levels (Fig. 2B ). In addition, the PIKfyve inhibitor apilimod and autolysosome acidification blocker bafilomycin each successfully reduced viral infection to ~0.1% or 1% of vehicle-treated controls, respectively. In contrast, the TMPRSS2 inhibitors aprotinin and camostat mesilate did not significantly inhibit viral infection. These results suggest that SARS-CoV-2 binds to iPS-CMs via the ACE2 receptor, and utilizes a CTSL (but not CTSB)-dependent endolysosomal route to infection, independent of TMPRSS2/serine protease-mediated activation at the cellular membrane.
We next examined whether priming the innate immune response could reduce SARS-CoV-2 infection of CMs. CMs were treated with IFN-α, IFN-β, IFN-γ, or IFN-λ prior to infection . Only pre-exposure to IFN-β decreased infection, and this effect was reversed by co-treatment with the JAK/Stat inhibitor ruxolitinib (Figs. 2C-D). Our observation that IFN-β but not IFN-α pretreatment reduced infection is in concordance with a previous report of differential antiviral activity of these two type-I interferons in a mouse model of myocarditis 33 , confirming that CMs can mount an antiviral response with appropriate stimulation.
To evaluate the transcriptional impact of SARS-CoV-2 on cardiac cells, we performed RNAsequencing of CFs, ECs, and iPSCs exposed to an MOI of 0.006 and CMs exposed to a range of elevating MOIs (0.001, 0.01, and 0.1). Sequencing recovered SARS-CoV-2 reads in an MOIand cell type-dependent fashion (Fig. 3A) , with SARS-CoV-2 reaching >50% of the recovered reads in CMs at the highest MOI. Principal component analysis (PCA) of the biological conditions revealed clustering based primarily on cell type, with CFs and ECs clustering together and CMs and iPSCs separating into distinct clusters (Fig. 3B) . Loading plots of the principal components supported this interpretation: genes determining the spectrum of variation between CMs and CF/ECs were associated with CMs (MYH7, MYH6, TNNT2) at one pole (Fig.  3C ) and anti-correlated with CF/EC specific genes at the other (FN1, COL1A2, TFPI2, MME) ( Supplementary Fig. S2A) . However, the distance between mock-treated CMs and the furthest infected CMs was greater than the distance between CMs and CFs or ECs, indicating that viral infection altered cellular expression profiles at least as strongly as cell type. Along this axis, the level of transcriptional disruption correlated poorly with MOI across all CM samples, potentially due to natural stochasticity in the kinetics of infection. Regrouping conditions by the level of transcriptional disruption allowed us to more clearly deduce transcriptional trends resulting from viral exposure (Figs. S2A-D). Infected cardiomyocytes downregulated pathways corresponding to cardiac muscle tissue organization and cellular respiration. Meanwhile, pathways related to the innate immune response and apoptosis were upregulated, although the innate response to viral infection decreased, suggesting active repression by SARS-CoV-2 as reported by others 34 ( Fig. 3D) . Anomalous upregulation of pathways associated with olfactory receptors and dysregulation of proteasome catabolism were also observed with increasing transcriptional disruption (Fig. 3D, S2E) .
The differentially regulated genes involved in inflammation and innate immunity reflected the observed preferential infectivity of CMs. Exposed CFs and ECs had a depressed cytokine response compared to CMs at all three MOIs (Supplementary Fig. S2E) , while infected CMs were enriched in genes involved in cytokine production and T-cell activation (OAS2, MX1, IFIT1, IL1B, IL6, TNF) ( Supplementary Fig. S2E) . Interestingly, CMs at each MOI showed clear dysregulation of contractile machinery, proteasomal subunits and ubiquitination (Supplementary Fig. 3A-G) . Genes involved in the Linker of Nucleoskeleton and Cytoskeleton (LINC) complex were disrupted, especially calmin and members of the nesprin family, both critical for anchoring the nucleus to the actin cytoskeleton ( Fig S4) . Furthermore, sarcomeric structural proteins, myosin light chains, and proteasome kinases and chaperones were strongly downregulated, while most myosin heavy chains were significantly upregulated (Fig. 3E, F,  Supplementary Fig. 3A-G) , suggesting a split effect of SARS-CoV-2 infection on the contractile and structural integrity of CMs.
Motivated by the expression changes in structural and contractile genes in our transcriptomic data, we performed high-content imaging of CMs following SARS-CoV-2 infection. We immediately noticed several abnormal structural features in infected CMs that were not seen in mock or heat-inactivated virus-treated CMs. Most notably, we observed widespread myofibrillar disruption throughout the cytoplasm: a unique pattern of specific, periodic cleavage of myofibrils into individual sarcomeric units of identical size without any linear alignment (Figs. 4A-C, Supplementary Figs. 4A-D) . Myofibrillar fragmentation was evident as early as 24 hours after infection, and significantly increased after 48 hours of viral exposure, suggesting that fragmentation increases over the course of infection (Fig. 4B) . This pattern of myofibrillar fragmentation was present in cells independently of actively replicating virus (as per dsRNA staining; Chi-square test for independence p-value = 0.81) (Fig. 4C) .
Co-staining of CMs with the thin filament marker cardiac Troponin T (cTnT), and the Z-disk marker α-actinin 2, revealed that SARS-CoV-2-induced myofibrillar fragments consisted of two bands of cTnT flanking a single α-actinin 2 band (Fig. 4D, Supplementary Figure S5E) . To examine sarcomeric fragmentation in greater detail, we employed TEM imaging of SARS-CoV-2 infected and mock-treated CMs. While intact sarcomeres were clearly identified with a classic dark Z-disk adjacent to a light I-band followed by a dark A-band, single fragmented myofibrils displayed an extended I-band and complete absence of the A-band (Fig. 4E) , indicating a liberation of thick filaments from sarcomere subunits.
Since transcriptomic profiling indicated that viral infection perturbed the proteasome system (Fig. 3F, Supplementary Fig. S3F) , we also examined if proteasome inhibition could recapitulate similar structural abnormalities. Although high doses of the proteasome inhibitor bortezomib induced myofibril fragmentations in CMs, the effect was much less frequent and less severe than the effect of SARS-CoV-2 infection, and was generally accompanied by diffuse cTnT staining (Supplementary Fig. S5G) . Furthermore, the well-known cardiotoxic drug doxorubicin did not induce myofibril fragmentation (data not shown), suggesting that proteasomal inhibition may specifically recapitulate part of the viral effects that lead to myofibrillar fragmentation. Additionally, we also observed a second structural phenotype where infected CMs often lacked nuclear DNA staining. This phenomenon was characterized by withdrawn sarcomeres and abnormally shaped or absent nuclear DNA signal (Fig. 4F) .
Interestingly, this abnormal nuclear structure is consistent with our observed transcriptomic dysregulation of the LINC complex (Supplementary Fig. S4 ).
We next asked whether the SARS-CoV-2 induced phenotypes observed in vitro could predict similar patterns of cardiac cell damage in vivo. We obtained autopsy specimens from three COVID-19 positive patients: one who had been diagnosed with myocarditis (COVID-M1), and two which had no reported cardiac involvement (COVID-A1 and COVID-A2). The myocardium of COVID-M1 revealed the expected immune infiltration of the tissue, in addition to significant disruptions in myocytes lacking DNA staining (Fig. 5A 6B ) both had loss of nuclear DNA comparable to that of the COVID-M1 patient. A slight difference (approximately 10%) in nuclear counts between regions of intact tissue vs disrupted tissue was detected across patients (Supplementary Fig. 6C ).
Hematoxylin & eosin staining did not reveal signs of sarcomeric damage in any samples, so we further probed the autopsy samples by immunostaining for sarcomere proteins. COVID-M1, COVID-A1, and COVID-A2 all revealed regions of severe myofibrillar anomalies. COVID-M1 displayed diffuse staining in myocytes in addition to occasional troponin-positive mononuclear cells in the vasculature (Fig. 5D, Supplementary Fig. 6D) , while COVID-A1 (Fig. 5E ) and COVID-A2 ( Supplementary Fig. 6E ) displayed clear and distinct patterns of disruptions to the sarcomeric banding. We attempted to identify viral presence in the tissues using antibodies against SARS-CoV-2 Spike protein, dsRNA, and two epitopes of the viral nucleocapsid protein, but were unable to detect viral signal in any autopsy patient tissues (Supplementary Fig. 6F) .
Overall, these patterns suggest that myocarditis in the heart leads to significant tissue damage, but is not a requirement for myofibrillar disruption of cardiac tissue. Additionally, these patterns of myocardial damage closely resemble similar cytopathic features observed by in vitro exposure of SARS-CoV-2 to iPSC-CMs.
In this study, we carried out a comprehensive analysis of the cytopathic effects of SARS-CoV-2 on human iPSC-derived cardiac cell types. Cytopathic effects were particularly striking in cardiomyocytes, which manifested a distinctive myofibrillar fragmentation into individual sarcomeres and a loss of nuclear DNA from intact cell bodies. Surprisingly, these cytopathic effects appear to occur independently of the presence of actively replicating SARS-CoV-2 virus, suggesting a larger spectrum of adverse consequences than initially assumed. Guided by these observations, we observed similar sarcomeric structural disruptions and cells lacking nuclear DNA in myocardium specimens from COVID-19 patients. Together, these results provide novel insights in the pathogenesis of SARS-CoV-2 heart damage, indicate new avenues for the development of cardioprotective interventions against COVID-19, and also raise significant concerns about the prevalence and severity of cardiac involvement.
Determining the mechanisms responsible for diminished cardiac function is critically important to develop cardioprotective therapies for COVID-19. We observe that SARS-CoV-2 infection creates extremely precise and ordered disruptions to the myofibrillar structure and dissolution of the cardiac contractile machinery, which would inevitably lead to functional collapse. The striking consistency and periodicity of fragmentation suggests specific protease activity responsible for separation of the sarcomeric thick and thin filaments, as shown by immunofluorescence and TEM. Additionally, our transcriptomic analyses suggest a compensatory overexpression of myosin heavy chain genes in response to targeted degradation (Fig 3D) . We also observed a significant depression of the ubiquitin-proteasome system upon infection, and myofibrillar fragmentation could be partially recapitulated by the addition of the proteasome inhibitor bortezomib (Supplementary Fig. S3; Supplementary Fig.  S5G) . Altogether, we speculate that SARS-CoV-2 infection may trigger cleavage of myofibrils by targeting myosin, which, when unopposed by the proteasome system, may result in the observed myofibrillar fragmentation phenotype.
Interestingly, myosin heavy chain protein family members contain the LKGG↓K sequence, which is known to be one of the sites used by the SARS-CoV-2 papain-like protease (PLpro) to cleave the viral polyprotein 35, 36 . A cleavage of myosin by the viral protease could result in the observed filament-specific cleavage, 37 though this interpretation does not immediately explain why myofibrillar pattern seems to be present in cells exposed to SARS-CoV-2 independently of dsRNA signal. Although further interrogation is needed to determine the precise mechanisms mediating myofibrillar fragmentation, the phenotype still informs expectations for histological specimens and therapeutic discovery.
To date, most myocardial histology from autopsy specimens of COVID-19 patients have revealed only general signs of myopathy, such as edema, occasional mononuclear infiltrate, and mild hypertrophy 18 . Although 30-50% of COVID-19 patients manifest clinical signs of cardiac dysfunction 13, 38 , histological hallmarks of COVID-19 in the heart have remained elusive. Our preliminary examinations of hematoxylin and eosin staining of COVID-19 myocardial samples revealed only minor disruption and generally intact myofibrillar anatomy. However, informed by our in vitro analysis, we identified clear features of similar myocardial damage by immunostaining for sarcomeric proteins. This correlation reveals the potential underlying etiology of SARS-CoV-2's impact on cardiac function and demonstrates that human iPS-derived models of myocardium can predict features of cardiac pathogenesis in COVID-19 patients.
In addition to myofibril disruption, we also identified a lack of nuclear DNA in many CMs after SARS-CoV-2 exposure, which was also observed in COVID-19 patient autopsy specimens.
Physically intact cardiomyocytes lacking nuclear DNA would normally reflect a terminal event such as apoptosis 39 . However, transcriptomic analysis suggests instead that SARS-CoV-2 may be disrupting the nuclear LINC complex, through which the cytoskeleton supports the shape and structure of the nucleus 40, 41 . Understanding the mechanism by which these aberrant nuclear phenotypes arise will be crucial to determine if they are a clinical feature induced by COVID-19 in the heart.
Aside from myocytes in the heart, non-myocyte cardiac stroma may also mediate some of the outcomes observed in COVID-19, such as cardiac hypertrophy and vascular dysfunction 31 . In our studies, CFs and ECs were not expected to be infectable due to their low levels of ACE2 expression. qPCR of exposed CFs and ECs supported these predictions and displayed no replicating virus, but we did observe potent cytopathic effects in both ECs and CFs. Interestingly, heat inactivated virus failed to recapitulate this effect, suggesting that abortive infection may be responsible for cytotoxicity. Future studies identifying specific mechanisms of viral toxicity to the cardiac stroma will be useful to determine how these cells may contribute to SARS-CoV-2-induced cardiac dysfunction.
The ability of iPS-CMs to model and predict cardiopathic consequences of infectious pathogens opens a wide array of potential avenues for discovery and validation of candidate cardioprotective therapies for COVID-19 and other diseases. For example, our finding that SARS-CoV-2 infects CMs via an endolysosomal route may indicate that clinical trials which target TMPRSS2 to prevent COVID-19 42,43 may not afford effective cardioprotection without orthogonal targeting of endosomal proteases 44 . Additionally, although cell-based drug screens exist for many pathogens, including SARS-CoV-2 45 , the unique cytoarchitecture of cardiomyocytes and the specific impacts of SARS-CoV-2 on these cells offer novel screening possibilities. For example, myocyte-specific myofibrillar fragmentation enables cell-based drug screens with an exceptionally sensitive reporter of CM infection. Identification of efficacious cardioprotective therapies may require preventing viral replication and maintaining sarcomeric integrity to achieve optimal therapeutic benefit. Reports of cardiac dysfunction incidence in COVID-19 patients range from 20% to 50%, independent of disease severity 8, 13, 38 . Cardiac damage is strongly associated with disease mortality 3, 8, 18, 20, 46 . In addition, due to the heart's innate lack of regenerative capacity 47,48 , a majority of these patients could suffer long-term cardiac sequelae from COVID-19. Our studies, which are analogous to mild cases of COVID-19 due to the low viral load and short time frame of viral exposure, display signs of striking cytopathic effects similar to those we observe in patient samples without any clinical diagnosis of cardiac involvement. These findings provide clues to the mechanism of cardiac pathology of COVID-19, and anticipate they will help guide the development of efficacious anti-viral and cardioprotective therapies to help manage and prevent heart damage in patients.
Research protocol for evaluation of autopsy specimens was approved by institutional review board at the University of California, San Francisco (UCSF)/Zuckerberg San Francisco General Hospital (ZSFG).
supplemented with 12µM CHIR99021 (day 0), and refreshed with R/B-24 hours later (day 1). From days 2-20, cells were fed every 2 days with cardiac fibroblast basal media (CFBM) (Lonza, CC-3131) supplemented with 75ng/mL bFGF. On day 20, CFs were singularized with Accutase for 10 minutes and replated at 7,000 cells/cm 2 onto tissue culture plastic 10cm dishes in FibroGRO medium (Millipore Sigma, SCMF001). FibroGRO media was changed every two days until the CFs reached approximately 80-90% confluency, at which point they were passaged with Accutase. SHF-CFs were validated to be >80% double-positive for TE-7 and vimentin by flow cytometry.
Endothelial Cell Differentiation WTC11 iPSCs were directed towards an endothelial cell (EC) lineage by the addition of E8 media supplemented with BMP4 (5 ng/ml) and Activin A (25 ng/ml) for 48 hours followed by E7BVi media, consisting of E6 medium supplemented with bFGF (50ng/ml), VEGF-A (50 ng/ml), BMP4 (50 ng/ml) and a TGFβ inhibitor, SB431542, (5 μM) for 72 hours. After 5 days of successive media changes, ECs were split and plated at high density in EGM media (Lonza, CC-3162) on tissue culture flasks coated with fibronectin (1:100, Sigma Aldrich F0895). On day 8, all cells were cryo-preserved and a fraction of ECs were assayed for >95% purity by flow cytometry using antibodies against mature EC markers CD31 and CDH5. ECs were seeded at a density of roughly 60,000 cells/cm 2 , 48 hours prior to infection.
Mixed cultures of iPS-CMs, iPS-ECs, and iPS-CFs were created by combining single cell suspensions of each cell types in a ratio of 60:30:10 CM:EC:CF at a density of 200,000 cells/mL. The mixed suspension was replated onto Matrigel-coated tissue culture plates 48 hours prior to infection at a density of 62,500 cells/cm 2 . 
The WA-1 strain (BEI resources) of SARS-CoV-2 was used for all experiments. All live virus experiments were performed in a Biosafety Level 3 lab. SARS-CoV-2 stocks were passaged in Vero cells (ATCC) and titer was determined via plaque assay on Vero cells as previously described 7 . Briefly, virus was diluted 1:10 2 -1:10 6 and incubated for 1 hour on Vero cells before an overlay of Avicel and complete DMEM (Sigma Aldrich, SLM-241) was added. After incubation at 37°C for 72 hours, the overlay was removed and cells were fixed with 10% formalin, stained with crystal violet, and counted for plaque formation. SARS-CoV-2 infections of iPSc and iPS-derived cardiac cells were done at a multiplicity of infection of 0.006 for 48 hours unless otherwise specified. For heat inactivation, SARS-CoV-2 stocks were incubated at 85°C for 5 min. Plaque assay for supernatant from infected cells was performed as above.
Infected and mock-treated cell cultures in coverslips were washed with Phosphate Buffered Solution (PBS) and fixed in 4% paraformaldehyde (PFA) overnight, followed by blocking and permeabilization with 0.1% Triton-X 100 (T8787, Sigma) and 5% BSA (A4503, Sigma) for one hour at RT. Antibody dilution buffer (Ab buffer) was comprised of PBS supplemented with 0.1% Triton-X 100 and 1% BSA. Samples were incubated with primary antibodies overnight at 4°C (Table 2) , followed by 3 washes with PBS and incubation with fluorescent-conjugated secondary antibodies at 1:250 in Ab buffer for 1 hour at RT (Table 2) . Coverslips were mounted onto SuperFrost Slides (FisherBrand, 12-550-15) with ProLong Antifade mounting solution with DAPI (Invitrogen, P36931. Images were acquired with a Zeiss Axio Observer Z.1 Spinning Disk Confocal (Carl Zeiss) or with an ImageXpress Micro Confocal High-Content Imaging System (Molecular Devices) and processed using ZenBlue and ImageJ.
Paraffin section of healthy and COVID-19 patient hearts were deparaffinized using xylene, rehydrated through a series of decreasing ethanol solutions (100%, 100%, 95%, 80%, 70%) and rinsed in PBS. Hematoxylin and eosin staining was performed according manufacturer instructions and the slides were mounted with Cytoseal 60 (Richard-Allan Scientific) and glass coverslips. For immunofluorescence staining, epitopes were retrieved by immersing slides through 35 min incubation at 95°C in citrate buffer (Vector Laboratories, pH 6) or Tris-EDTA buffer (Cellgro, pH 9). Slides were cooled for 20min at RT and washed with PBS. Samples were permeabilized in 0.2% Triton X-100 (Sigma) in PBS by slide immersion and washed in PBS.
Blocking was performed in 1.5% normal donkey serum (NDS; Jackson ImmunoResearch) and PBS solution for 1h at RT. Primary and secondary antibody cocktails were diluted in blocking solution (Table 2) . PBS washes were performed after primary (overnight, 4 o C) and secondary antibody (1h, RT) incubations. Nuclei were stained with Hoechst and coverslips were mounted on slides using ProLong TM Gold Antifade Mountant. Samples were imaged on the Zeiss Axio Observer Z1. 
Cultured cells were lysed with Qiagen buffer RLT (Qiagen, 79216) supplemented with 1% βmercaptoethanol (Bio-Rad, 1610710) and RNA was isolated using the RNeasy Mini Kit (Qiagen 74104) or Quick-RNA MicroPrep (ThermoFisher, 50444593) and quantified using the NanoDrop 2000c (ThermoFisher). Viral load was measured by detection of the viral Nucleocapsid (N) transcript through one-step quantitative real-time PCR, performed using PrimeTime Gene Expression Master Mix (Integrated DNA Technologies, 1055772) with primers and probes specific to N ('N5') and RPP30 as in internal reference. RT-qPCR reactions were performed on a CFX384 (BioRad) and delta cycle threshold (ΔCt) was determined relative to RPP30 levels. Viral detection levels in pharmacologically treated samples were normalized to DMSO-treated controls. 
To generate libraries for RNA-sequencing, RNA isolate quality was assessed with an Agilent Bioanalyzer 2100 on using the RNA Pico Kit (Agilent, 5067-1513). 10ng of each RNA isolate was then prepared using the Takara SMARTer Stranded Total RNA-Seq Kit v2 -Pico Input