Introduction
Coronaviruses cause respiratory disease in a wide variety of hosts. During the ongoing pandemic of SARS‐CoV‐2 causing coronavirus disease 19 (COVID‐19), clinical signs other than respiratory symptoms have been linked to infection, frequently associated with neurological symptoms such as anosmia and ageusia. These features have been related to an over‐responsiveness of patients' immune system to SARS‐CoV‐2 (Bhaskar
et al,
2020; Han
et al,
2020; Qiu
et al,
2020). Consequently, there is an urgent need to understand the hallmarks of this over‐responsiveness and to find novel therapeutics or repurpose drugs to improve the clinical condition of COVID‐19 patients (Batalha
et al,
2021).
Ivermectin (IVM), a macrocyclic lactone, is a commercially available anti‐parasitic drug which prevents infection by a wide range of endo‐ and ectoparasites (Sajid
et al,
2006; Heidary & Gharebaghi,
2020). IVM is an efficient positive allosteric modulator of the α‐7 nicotinic acetylcholine receptor (nAChR) (Krause
et al,
1998) and of several ligand‐gated ion channels, including the muscle receptor for glutamate (GluCl) in worms (Hibbs & Gouaux,
2011). Furthermore, IVM has been shown to exert an immunomodulatory effect in humans and animals (Sajid
et al,
2006; Heidary & Gharebaghi,
2020) under conditions that are known to involve the α‐7 nAChR (Pavlov & Tracey,
2012), even though its underlying mechanisms are yet to be established (Laing
et al,
2017). A direct or indirect interaction of SARS‐CoV‐2 with nAChR has also been hypothesized, in particular because of sequence homologies between SARS‐CoV‐2 spike proteins and nAChR ligands such as snake venom toxins (Changeux
et al,
2020). IVM has been shown to be active beyond its anti‐parasitic activity in a wide variety of pathologies, including cancer, allergy, and viral infections (Laing
et al,
2017). Recently, IVM has been reported to reduce viral load and improve the clinical status of mice infected by an animal coronavirus, the mouse hepatitis virus (MHV) (Arévalo
et al,
2021).
In vitro inhibition of SARS‐CoV‐2 replication by IVM in Vero/hSLAM cells has also been reported (Caly
et al,
2020), albeit at much higher concentrations (50‐ to 100‐fold) than those clinically attainable in humans (150–400 µg/kg) (Guzzo
et al,
2002; Bray
et al,
2020; Chaccour
et al,
2020).
The aim of this study is to investigate the impact of IVM on the pathogenesis of COVID‐19, in a SARS‐CoV‐2 infection model, the golden Syrian hamster. This species is naturally permissive to this virus and the most reliable and affordable animal model for COVID‐19 (Chan
et al,
2020; Muñoz‐Fontela
et al,
2020). Moreover, it was recently used to demonstrate the importance of lowering the inflammation with intranasal administration of type I IFN to prevent disease progression (Hoagland
et al,
2020). Male and female adult golden Syrian hamsters were intranasally inoculated with 6 × 10
4 PFU of SARS‐CoV‐2 [BetaCoV/France/IDF00372/2020]. This inoculum size was selected as it invariably causes symptomatic infection in golden Syrian hamster, with a high incidence of anosmia and high viral loads in the upper and lower respiratory tracts within 4 days post‐inoculation (dpi) (de Melo
et al,
2021). At the time of infection, animals received a single subcutaneous injection of IVM at the anti‐parasitic dose of 400 µg/kg, commonly used in human clinical setting, and were monitored over 4 days.
Here, we show that the modulation of the host’s inflammatory response using IVM as a repurposed drug strongly diminished the clinical score and severity of the disease (including anosmia) observed in these animals, although it has no impact on viral load. IVM‐treated animals presented a strong modulation in several signaling pathways, including a significant reduction of the type I and III interferon response and of the Il‐6/Il‐10 ratio, along with the presence of M2 macrophages in the lung. These effects were mostly compartmentalized and sex‐dependent, and treated infected females exhibited better clinical outcomes.
Conclusions
Our results demonstrate that IVM improves clinical outcome in SARS‐CoV‐2‐infected animals and is associated with a reduced inflammatory status, but with no impact of SARS‐CoV‐2 loads in the upper and lower respiratory tracts. Thus, in hamsters, as in humans (preprint: Cereda
et al,
2020; Hasanoglu
et al,
2021), symptomatology and therefore the severity of SARS‐CoV‐2 infection is not strictly correlated with viral load. The main effect of IVM in the lungs is on type I and III IFN responses and other related signaling pathways including phospholipases, kinases, and adenylate cyclases, which are important therapeutic targets (Melotti
et al,
2014; Raker
et al,
2016; Hu
et al,
2020; Li
et al,
2020; preprint: Masood
et al,
2020; Isidori
et al,
2021), and this translates clinically into an improved clinical score.
The results presented herein are consistent with a role of type I and III IFN responses in the pathogenesis of SARS‐CoV‐2‐associated lung disease in hamsters. They show that IVM administration limits IFN response and lung inflammation, even though defects in the type I IFN pathways have been associated with severe COVID‐19 (Bastard
et al,
2020; Zhang
et al,
2020a). This result may suggest that while IFN signaling is crucial to control viral replication and prevent severe disease, in infected hamsters, which only develop a moderate disease, IFN signaling may actually increase tissue damage and associated signs such as anosmia. This is consistent with previous reports in animal models where inhibition of IFN pathways was shown to increase viral replication but with lower lung pathology (Boudewijns
et al,
2020; Sun
et al,
2020).
Even if the effects observed may, to some extent, share similarities with the impact of dexamethasone (Horby
et al,
2020) and tocilizumab (anti‐IL‐6) (Rossotti
et al,
2020) on COVID‐19, IVM action is steady and strong in the golden hamsters and is not expected to block the effectors of inflammation but rather to dampen its initiation. Interestingly, the activation of nuclear factor erythroid 2‐related transcription factor (Nrf2) via low‐dose radiotherapy (LDRT) treatment has been proposed to cause a shift from M1 to M2 macrophages and a blockade of NLRP3 inflammasomes (Calabrese
et al,
2021) and to be potentially beneficial on the lungs of COVID‐19 patients. Along the same line, we noticed via our RNA‐seq analyses that IVM treatment increased the gene expression of Nrf2 (or Nfe2l2) in the lungs of female hamsters and slightly in males (Dataset
EV1), which gives additional evidence of the broad and upstream activity of IVM during SARS‐CoV‐2 infection. Further, our data show that these effects are compartmentalized and that the upper and lower respiratory tracts of hamsters respond differently to IVM treatment.
Considerable sex differences are observed in terms of clinical presentation, inflammatory profile, and transcriptomic signatures in the lungs of hamsters, as seen in COVID‐19 human patients, where men tend to develop more severe disease than women (Jin
et al,
2020; Takahashi
et al,
2020), possibly in relation with androgen signaling (vom Steeg & Klein,
2019; Samuel
et al,
2020; Scully
et al,
2020). Interestingly, sex steroids, here female hormones, might also influence both the course of COVID‐19 in hamsters and the effects of IVM, possibly due to the potentiation of cell receptors signaling in females, such as nAChRs (Krause
et al,
1998; Cross
et al,
2017) and GlyRs (Van Den Eynden
et al,
2009; Cerdan
et al,
2020), of which IVM is a positive allosteric modulator.
Moreover, the data presented herein are consistent with those reported in human clinical trials with IVM (Kory
et al,
2021). In humans, IVM is widely used as anti‐helminthic and anti‐scabies at therapeutic doses (150–400 µg/kg) (Guzzo
et al,
2002) that are in the range of those used in our hamster experiments. Further, several clinical trials on COVID‐19 using IVM have been declared, where IVM has been associated with reduction of inflammatory markers and disease severity (preprint: Hill
et al,
2021; Kaur
et al,
2021). Interestingly, in a long‐term care facility where the residents received IVM to control a scabies outbreak, no death or severe COVID‐19 was observed (Bernigaud
et al,
2021). IVM has already been administered to hospitalized COVID‐19 patients, with contrasting outcomes: one study related no efficacy of late IVM administration (8–18 days after symptom onset) in severe COVID‐19 patients treated in combination with other drugs (hydroxychloroquine, azithromycin, tocilizumab, steroids) (Camprubí
et al,
2020), whereas another study reported lower mortality, especially in severe COVID‐19 patients treated with IVM in addition to other treatments (hydroxychloroquine, azithromycin, or both) (Rajter
et al,
2020). Importantly, in a study that administered IVM alone within 72 h of symptom onset, the authors noticed an important diminution of anosmia/hyposmia in the IVM group without differences in PCR positivity between IVM and placebo groups (Chaccour
et al,
2021).
The presently available data support the view that IVM pharmacokinetics is quite stable across species, generally with slow absorption, broad distribution, low metabolism, and slow excretion, even if some conditions, such as route of administration, formulation, and body condition, may modify these features. In humans, the oral route is the only approved, and the administration of a single dose of 30 mg of IVM (corresponding to 347–594 µg/kg) leads to a
Cmax (maximum concentration) of 84.8 ng/ml, a t
max (time to reach the
Cmax) of 4.3 h and a half‐life of 20.1 h (Guzzo
et al,
2002). In animal models, the subcutaneous route is most often used, and comparatively, golden hamsters that received a subcutaneous injection of 400 µg/kg of IVM (as in the present study) showed comparable results with a
Cmax of 80.2 ng/ml and a t
max of ˜4 h (Hanafi
et al,
2011). Further, based on veterinary data (Lifschitz
et al,
2000), simulations using a minimal physiologically based pharmacokinetic (mPBPK) model revealed that the lungs would be exposed to IVM concentrations 2.7× greater than those found in the plasma (Jermain
et al,
2020). Yet, this dose did not suffice to achieve the range of antiviral concentrations reported
in vitro (Caly
et al,
2020).
Consequently, considering the results observed in the golden hamster model, IVM may be considered as a therapeutic agent against COVID‐19, which would not strongly affect SARS‐CoV‐2 replication but limit the pathophysiological consequences of the infection
in vivo, potentially mediated by type I and III IFN responses and several other related signaling pathways, and a favorable M1/M2 myleoid cells ratio in the lungs. A characteristic modulation of the immune response in the lower airways was observed in IVM‐treated hamsters characterized by a transcriptomic profile similar to that observed in humans exhibiting less severe symptoms and a better prognosis (preprint: Masood
et al,
2020; McElvaney
et al,
2020b). Our data are consistent with the hypothesis that this effect is mediated by the cholinergic anti‐inflammatory action of IVM on the vagus nerve reflex (Changeux
et al,
2020; Tizabi
et al,
2020), that should be addressed experimentally. In particular, the precise contribution of the nAChR in IVM action should be elucidated in comparison with that of other possible IVM targets (Zemkova
et al,
2014). Altogether, this study brings the proof of concept that an IVM‐based immunomodulatory therapy improves the clinical condition of SARS‐CoV‐2‐infected hamsters, and in clinical trials, it alleviates symptoms of COVID‐19 in humans and possibly limits post‐COVID‐19 syndrome (also known as long COVID) via an anti‐inflammatory action.
Materials and Methods
Ethics
All animal experiments were performed according to the French legislation and in compliance with the European Communities Council Directives (2010/63/UE, French Law 2013–118, February 6, 2013) and according to the regulations of Pasteur Institute Animal Care Committees. The Animal Experimentation Ethics Committee (CETEA 89) of the Institut Pasteur approved this study (200023; APAFIS#25326‐2020050617114340 v2) before experiments were initiated. Hamsters were housed by groups of 4 animals in isolators and manipulated in class III safety cabinets in the Pasteur Institute animal facilities accredited by the French Ministry of Agriculture for performing experiments on live rodents. All animals were handled in strict accordance with good animal practice.
Production and titration of SARS‐CoV‐2 virus
The isolate BetaCoV/France/IDF00372/2020 (EVAg collection, Ref‐SKU: 014V‐03890) was kindly provided by Sylvie Van der Werf. Viral stocks were produced on Vero‐E6 cells infected at a multiplicity of infection of 1 × 10
−4 plaque‐forming units (PFU). The virus was harvested 3 days post‐infection, clarified, and then aliquoted before storage at −80°C. Viral stocks were titrated on Vero‐E6 cells by classical plaque assays using semisolid overlays (Avicel, RC581‐NFDR080I, DuPont) (Baer & Kehn‐Hall,
2014).
SARS‐CoV‐2 model and ivermectin treatment of hamsters
Male and female Syrian hamsters (Mesocricetus auratus; RjHan:AURA) of 5–6 weeks of age (average weight 60–80 g) were purchased from Janvier Laboratories and handled under specific pathogen‐free conditions. The animals were housed and manipulated in isolators in a Biosafety level‐3 facility, with ad libitum access to water and food. Before manipulation, animals underwent an acclimation period of 1 week.
Animals were anesthetized with an intraperitoneal injection of 200 mg/kg ketamine (Imalgène 1000, Merial) and 10 mg/kg xylazine (Rompun, Bayer) and received one single subcutaneous injection of 200 µl of freshly diluted ivermectin (I8898, Sigma‐Aldrich) at the classical anti‐parasitic dose of 400 µg/kg (Beco
et al,
2001) (or at 100–200 µg/kg for the dose–response experiment). Non‐treated animals received one single subcutaneous injection of 200 µl of physiological solution. 100 µl of physiological solution containing 6 × 10
4 PFU of SARS‐CoV‐2 was then administered intranasally to each animal (50 µl/nostril). Mock‐infected animals received the physiological solution only.
Infected and mock‐infected animals were housed in separate isolators, and all hamsters were followed up daily during 4 days at which the body weight and the clinical score were noted. The clinical score was based on a cumulative 0–4 scale: ruffled fur, slow movements, apathy, and absence of exploration activity.
At day 3 post‐infection (dpi), animals underwent a food finding test to assess olfaction as previously described (Lazarini
et al,
2012; de Melo
et al,
2021). Briefly, 24 h before testing, hamsters were fasted and then individually placed into a fresh cage (37 × 29 × 18 cm) with clean standard bedding for 10 min. Subsequently, hamsters were placed in another similar cage for 2 min when about five pieces of cereals were hidden in 1.5 cm bedding in a corner of the test cage. The tested hamsters were then placed in the opposite corner, and the latency to find the food (defined as the time to locate cereals and start digging) was recorded using a chronometer. The test was carried out during a 15‐min period. As soon as food was uncovered, hamsters were removed from the cage. One minute later, hamsters performed the same test but with visible chocolate cereals, positioned upon the bedding. The tests were realized in isolators in a Biosafety level‐3 facility that were specially equipped for that.
At 4 dpi, animals were euthanized with an excess of anesthetics (ketamine and xylazine) and exsanguination (AVMA,
2020), and samples of nasal turbinates and lungs were collected and immediately frozen at −80°C. Fragments of lungs were also collected and fixed in 10% neutral buffered formalin.
RNA isolation and transcriptional analyses by quantitative PCR from golden hamsters' tissues
Frozen tissues were homogenized with TRIzol (15596026, Invitrogen) in Lysing Matrix D 2‐ml tubes (116913100, MP Biomedicals) using the FastPrep‐24™ system (MP Biomedicals) at the speed of 6.5 m/s during 1 min. Total RNA was extracted using the Direct‐zol RNA MicroPrep Kit (R2062, Zymo Research: nasal turbinates) or MiniPrep Kit (R2052, Zymo Research: lung) and reverse‐transcribed to first‐strand cDNA using the SuperScript™ IV VILO™ Master Mix (11766050, Invitrogen). qPCR was performed in a final volume of 10 μl per reaction in 384‐well PCR plates using a thermocycler (QuantStudio 6 Flex, Applied Biosystems). Briefly, 2.5 μl of cDNA (12.5 ng) was added to 7.5 μl of a master mix containing 5 μl of Power SYBR Green Mix (4367659, Applied Biosystems) and 2.5 μl of nuclease‐free water with nCoV_IP2 primers (nCoV_IP2‐12669Fw: 5′‐ATGAGCTTAGTCCTGTTG‐3′; nCoV_IP2‐12759Rv: 5′‐CTCCCTTTGTTGTGTTGT‐3′) at a final concentration of 1 μM (WHO,
2020). The amplification conditions were as follows: 95°C for 10 min, 45 cycles of 95°C for 15 s and 60°C for 1 min; followed by a melt curve, from 60 to 95°C. Viral load quantification of hamster tissues was assessed by linear regression using a standard curve of eight known quantities of plasmids containing the
RdRp sequence (ranging from 10
7 to 10
0 copies). The threshold of detection was established as 200 viral copies/µg of RNA. The Golden hamster gene targets were selected for quantifying host inflammatory mediator transcripts in the tissues using the
Hprt (hypoxanthine phosphoribosyltransferase), the
γ‐
actin, and/or the actinB genes as reference (Appendix Table
S1). Variations in gene expression were calculated as the
n‐fold change in expression in the tissues from the infected hamsters compared with the tissues of the uninfected ones using the
method (Pfaffl,
2001).
Droplet digital PCR (ddPCR)
Reverse transcription
200 ng of RNA was reverse‐transcribed using iScript Advanced cDNA Synthesis kit for RT–qPCR (1702537, Bio‐Rad) according to the manufacturer's specifications.
Quantitative PCR for γ‐actin and Hprt reference genes
Real‐time PCR was performed in a CFX96 qPCR machine (Bio‐Rad). All samples were measured in duplicate. The 10 μl PCR included 0.8 ng of cDNA, 1× PowerUp PCR master mix (A25742, Applied Biosystems), and 0.5 µM of each primer (Appendix Table
S1). The reactions were incubated in a 96‐well optical plate at 95°C for 2 min, followed by 40 cycles of 95°C for 15 s and 60°C for 1 min.
Droplet digital PCR
ddPCRs were performed on the QX200 Droplet Digital PCR system according to the manufacturer's instructions (Bio‐Rad). Briefly, reaction mixture consisted in 10 μl ddPCR Supermix for probe no dUTP (1863023, Bio‐Rad), 0.25‐1 ng of cDNA, primers and probes for E/IP4 and N/nsp13 duplex reactions used at concentration listed in Appendix Table
S2 in a final volume of 20 μl. PCR amplification was conducted in a iCycler PCR instrument (Bio‐Rad) with the following condition: 95°C for 10 min, 40 cycles of 94°C for 30 s with a ramping of 2°/s, 59°C for 1 min with a ramping of 2°/s, followed by 98°C for 5 min with a ramping of 2°/s and a hold at 4°C. After amplification, the 96‐well plate was loaded onto the QX200 droplet reader (Bio‐Rad) that measures automatically the fluorescence intensity in individual droplets. Generated data were subsequently analyzed with QuantaSoft™ software (Bio‐Rad) based on positive and negative droplet populations. Data are expressed as CPD (copy per droplets) normalized to
γ‐
actin and
Hprt reference gene relative expression.
Viral titration in golden hamsters' lung
Frozen lung fragments were weighted and homogenized with 1 ml of ice‐cold DMEM supplemented with 1% penicillin/streptomycin (15140148, Thermo Fisher) in Lysing Matrix M 2‐ml tubes (116923050‐CF, MP Biomedicals) using the FastPrep‐24™ system (MP Biomedicals) and the following scheme: homogenization at 4.0 m/s during 20 s, incubation at 4°C during 2 min, and new homogenization at 4.0 m/s during 20 s. The tubes were centrifuged at 10,000
g during 1 min at 4°C, and the supernatants were titrated on Vero‐E6 cells by classical plaque assays using semisolid overlays (Avicel, RC581‐NFDR080I, DuPont) (Baer & Kehn‐Hall,
2014).
Transcriptomics analysis in golden hamsters' lung
RNA preparation was used to construct strand‐specific single‐end cDNA libraries according to the manufacturers' instructions (TruSeq Stranded mRNA sample prep kit, Illumina). Illumina NextSeq 500 sequencer was used to sequence libraries. The RNA‐seq analysis was performed with the Sequana framework (Cokelaer
et al,
2017). We used the RNA‐seq pipeline (v0.9.16), which is available online (
https://github.com/sequana/sequana_rnaseq). It is built on top of Snakemake 5.8.1 (Köster & Rahmann,
2012). Reads were trimmed from adapters using Cutadapt 2.10 (Martin,
2011) and then mapped to the golden hamster MesAur1.0 genome assembly from Ensembl using STAR 2.7.3a (Dobin
et al,
2012). FeatureCounts 2.0.0 (Liao
et al,
2014) was used to produce the count matrix, assigning reads to features using annotation MesAur1.0.100 with strand‐specificity information. Quality control statistics were summarized using MultiQC 1.8 (Ewels
et al,
2016). Statistical analysis on the count matrix was performed to identify differentially regulated genes, comparing infected versus non‐infected samples considering all samples and separating by sex. Clustering of transcriptomic profiles was assessed using a principal component analysis (PCA). Differential expression testing was conducted using DESeq2 library 1.24.0 (Love
et al,
2014) scripts based on SARTools 1.7.0 (Varet
et al,
2016) indicating the significance (Benjamini–Hochberg‐adjusted
P‐values, false discovery rate FDR < 0.05) and the effect size (fold change) for each comparison. Finally, enrichment analysis was performed using modules from Sequana, first by converting golden hamster ensembl ids to gene names and then using human annotations for GO terms and KEGG pathways. The GO enrichment module uses PantherDB (Mi
et al,
2019) and QuickGO (Huntley
et al,
2014) services; the KEGG pathways enrichment uses gseapy (
https://github.com/zqfang/GSEApy/), EnrichR (Chen
et al,
2013), KEGG (Kanehisa & Goto,
2000), and BioMart services. All programmatic accesses to the online web services were performed via BioServices (Cokelaer
et al,
2013).
Histopathology
Lung fragments fixed in 10% neutral buffered formalin were embedded in paraffin. Four‐µm‐thick sections were cut and stained with hematoxylin and eosin staining. The slides were then scanned using Axioscan Z1 Zeiss slide scanner, using the Zen 2 blue edition software.
Immunofluorescence
Lung fragments fixed in 10% neutral buffered formalin were washed in PBS and then embedded in O.C.T compound (4583, Tissue‐Tek), frozen on dry ice, and cryostat‐sectioned into 20‐µm‐thick sections. Sections were rinsed in PBS, and epitope retrieval was performed by incubating sections for 20min in citrate buffer pH 6.0 (C‐9999, Sigma‐Aldrich) at 96°C for 20 min. Sections were then blocked in PBS supplemented with 10% goat serum, 4% fetal calf serum, and 0.4% Triton X‐100 for 2 h at room temperature, followed by overnight incubation at 4°C with primary antibodies: rat anti‐Ly6G (1/100, 551459, BD‐Biosciences), chicken anti‐Iba1 (1/500, 234006, Synaptic Systems), rabbit anti‐Arg1 (1/250, PA5‐29645, Invitrogen), and rabbit anti‐SARS‐CoV nucleoprotein (1/500, provided by Dr Nicolas Escriou, Institut Pasteur, Paris). After rinsing, slides were incubated with the appropriate secondary antibodies (1/500: goat anti‐rat Alexa Fluor 546, A11081, Invitrogen; goat anti‐rabbit Alexa Fluor 488, A11034, Invitrogen; goat anti‐chicken Alexa Fluor 647, A32933, Invitrogen) for 2 h at room temperature. All sections were then counterstained with Hoechst (H3570, Invitrogen), rinsed thoroughly in PBS, and mounted in Fluoromount‐G (15586276, Invitrogen) before observation with a Zeiss LM 710 inverted confocal microscope through a Plan Apochromat 20x/0.8 Ph2 M27 lens. Cell quantification was performed in an automated manner using ImageJ. Single‐channel images were extracted, thresholded, and converted to binary images. Cells were then counted using the Particles Analyzer ImageJ plug‐in.
Statistics
Statistical analysis was performed using Prism software (GraphPad, version 9.0.0, San Diego, USA), with P < 0.05 considered significant. Quantitative data were compared across groups using log‐rank test or two‐tailed Mann–Whitney test. Randomization and blinding were not possible due to pre‐defined housing conditions (separated isolators between infected and non‐infected animals). Ex vivo analysis was blinded (coded samples). All animals were included, and data were provided from 2 replications, except food finding in males, that were replicated 3 times.
Author contributions
JPC and HB conceived the experimental hypothesis. GDM, FLaz, FLar, and HB designed the experiments. GDM, FLaz, FLar, LF, LK, SL, AM, and DH performed the experiments. GDM, FLaz, FLar, LF, EK, SL, AM, TC, PP, ML, and P‐ML analyzed the data. GDM, J‐PC, and HB wrote the manuscript, and all authors edited it.
Acknowledgements
The SARS‐CoV‐2 strain was supplied by the National Reference Centre for Respiratory Viruses hosted by Institut Pasteur (Paris, France) and headed by Dr. Sylvie van der Werf. The human sample from which strain 2019‐nCoV/IDF0372/2020 was isolated has been provided by Dr. X. Lescure and Pr. Y. Yazdanpanah from the Bichat Hospital (Paris, France). This work was supported by Institut Pasteur TASK FORCE SARS COV2 (NicoSARS, NeuroCovid, and Cov‐DROP projects) and received help from the European Union's Horizon 2020 Framework Programme for Research and Innovation under Specific Grant Agreement No. 945539 (Human Brain Project SGA3). We thank Elodie Turc and Laure Lemée, Biomics Platform, C2RT, Institut Pasteur, Paris, France, supported by France Génomique (ANR‐10‐INBS‐09‐09), IBISA and the Illumina COVID‐19 Projects' offer. We would like to thank Marion Berard, Laetitia Breton, Rachid Chennouf, Hamidou Diakhate, and Eddie Maranghi for their help in implementing experiments in the Institut Pasteur animal facilities and Nicolas Escriou Innovation laboratory: Vaccines, Institut Pasteur, Paris, for providing the anti‐SARS‐CoV‐2 nucleoprotein antibody. We thank Magali Tichit and Johan Bedel for the help with histopathology. We thank Gerard Orth, Arnaud Tarantola, and Andrew Holtz for critical reading of the manuscript. The synopsis illustration was created with BioRender.com.