Endothelial thrombomodulin downregulation caused by hypoxia contributes to severe infiltration and coagulopathy in COVID-19 patient lungs

Published:January 12, 2022DOI:https://doi.org/10.1016/j.ebiom.2022.103812

Background

Thromboembolism is a life-threatening manifestation of coronavirus disease 2019 (COVID-19). We investigated a dysfunctional phenotype of vascular endothelial cells in the lungs during COVID-19.

Methods

We obtained the lung specimens from the patients who died of COVID-19. The phenotype of endothelial cells and immune cells was examined by flow cytometry and immunohistochemistry (IHC) analysis. We tested the presence of severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) in the endothelium using IHC and electron microscopy.

Findings

The autopsy lungs of COVID-19 patients exhibited severe coagulation abnormalities, immune cell infiltration, and platelet activation. Pulmonary endothelial cells of COVID-19 patients showed increased expression of procoagulant von Willebrand factor (VWF) and decreased expression of anticoagulants thrombomodulin and endothelial protein C receptor (EPCR). In the autopsy lungs of COVID-19 patients, the number of macrophages, monocytes, and T cells was increased, showing an activated phenotype. Despite increased immune cells, adhesion molecules such as ICAM-1, VCAM-1, E-selectin, and P-selectin were downregulated in pulmonary endothelial cells of COVID-19 patients. Notably, decreased thrombomodulin expression in endothelial cells was associated with increased immune cell infiltration in the COVID-19 patient lungs. There were no SARS-CoV-2 particles detected in the lung endothelium of COVID-19 patients despite their dysfunctional phenotype. Meanwhile, the autopsy lungs of COVID-19 patients showed SARS-CoV-2 virions in damaged alveolar epithelium and evidence of hypoxic injury.

Interpretation

Pulmonary endothelial cells become dysfunctional during COVID-19, showing a loss of thrombomodulin expression related to severe thrombosis and infiltration, and endothelial cell dysfunction might be caused by a pathologic condition in COVID-19 patient lungs rather than a direct infection with SARS-CoV-2.

Funding

This work was supported by the Johns Hopkins University, the American Heart Association, and the National Institutes of Health.

Keywords

 Evidence before this study

Normally functioning endothelial cells protect the blood vessel by preventing unwanted coagulation. In COVID-19 patients, plasma levels of soluble markers associated with endothelial cell activation and dysfunction are elevated. To understand the phenotypic changes of vascular endothelial cells during COVID-19, we searched the PubMed database with the terms “COVID-19 endothelial cells” in combination with “phenotype”, “flow cytometry”, or “immunohistochemistry”. However, there was no article showing a comprehensive profile of endothelial cell phenotype in COVID-19.

 Added value of this study

In this study, we report a phenotypic profile of endothelial cells in the COVID-19 patient lungs by using flow cytometry and immunohistochemistry (IHC) analysis. Pulmonary endothelial cells of COVID-19 patients exhibited a dysfunctional phenotype related to thrombosis. Especially, we found a loss of thrombomodulin expression, a potent anticoagulant and anti-inflammatory molecule, in endothelial cells of COVID-19 patients. As a mechanism of endothelial dysfunction in COVID-19, we tested if SARS-CoV-2 can directly infect endothelial cells. We found no clear evidence of SARS-CoV-2 infection in endothelial cells using IHC stain, electron microscopy, and in vitro viral infection.

 Implications of all the available evidence

These findings provide evidence that endothelial cells may be a critical player to contribute to severe thrombosis in the lungs of COVID-19 patients and that endothelial thrombomodulin expression would be a potential therapeutic target for COVID-19-related immunothrombosis.

Introduction

The global pandemic of coronavirus disease 2019 (COVID-19) is caused by a novel severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) infection that primarily affects the respiratory tract.1 These respiratory effects can range from mild upper respiratory disease to severe pneumonia and acute respiratory distress syndrome (ARDS).1 However, patients hospitalized with COVID-19 also experienced an increased incidence of venous and arterial thromboembolic complications including acute pulmonary embolism (PE), deep vein thrombosis (DVT), ischemic stroke, or myocardial infarction.234 The incidence of thrombotic complications in intensive care unit patients has been reported between 16 and 49 percent.234 In a meta-analysis of 277 autopsy cases, small vessel thrombi were found in 10·8 percent of cases, and 19·1 percent had macrovascular thrombi such as DVT, PE, and mural thrombi.5 Patients hospitalized with COVID-19 have elevated levels of D-dimer, a thrombosis biomarker, and fibrin degradation products.6 Given the prominence of microthrombi in COVID-19, it was proposed to rename COVID-19 as MicroCLOTS (microvascular COVID-19 lung vessels obstructive thromboinflammatory syndrome).7Normally functioning vascular endothelium protects blood vessels by regulating vascular tone and preventing unwanted coagulation, playing an essential role in preventing ARDS.8,9 Endothelial cells express key molecules that regulate the balance between hemostasis and thrombosis such as thrombomodulin and von Willebrand factor (VWF).9101112 In COVID-19 patients, elevated plasma levels of soluble VWF, thrombomodulin, intercellular adhesion molecule 1 (ICAM-1), vascular adhesion molecule 1 (VCAM-1), and P-selectin have been shown, suggesting endothelial cell activation and dysfunction.13141516 Several researchers predicted that vascular endothelial cells in COVID-19 increased adhesion molecule expression for platelet and leukocyte migration and pro-inflammatory cytokine and chemokine production.8,17,18 Moreover, endothelial cell activation and damage were proposed to play a central role in the pathogenesis of COVID-19 associated with ARDS, vascular leakage, pulmonary edema, and a procoagulant state.8,18,19 Endothelial cell dysfunction was also expected to cause children’s multisystem inflammatory syndrome, which develops in a small percentage of children infected with SARS-CoV-2.20However, the phenotypic and transcriptional changes of endothelial cells in COVID-19 have not been fully described. Many single-cell RNA sequencing (scRNA-seq) studies on COVID-19 were performed with non-tissue samples such as peripheral blood mononuclear cells, bronchoalveolar lavage fluid, nasopharyngeal swab, and sputum, barely containing endothelial cells.2122232425 Several studies using the lung tissues from human or non-human primates have been conducted, but they intended to investigate a host entry for SARS-CoV-2 – angiotensin-converting enzyme 2 (ACE2) and transmembrane serine protease 2 (TMPRSS2) – not showing a comprehensive profile of endothelial cell phenotype and transcriptome.2627282930 Furthermore, endothelial transmembrane molecules such as thrombomodulin and ICAM-1 are generally cleaved by enzymes and released into the blood in diseased conditions, suggesting that a transcriptional profile of endothelial cells would not reflect their dysfunctional phenotype during COVID-19.11,12,31 Also, it remains unclear whether endothelial cell dysfunction and damage in COVID-19 are induced by direct SARS-CoV-2 infection or indirectly through pathologic conditions such as hypoxia and pro-inflammatory cytokine milieu. Some studies have shown ACE2 expression on endothelial cells with limited evidence for the direct infection by SARS-CoV-2.32,33 However, others have disputed ACE2 expression on endothelial cells and further infection by SARS-CoV-2.3435363738394041Here, we provide a unique perspective on the phenotypic changes of endothelial cells in the autopsy lungs, hearts, and kidneys of COVID-19 patients using flow cytometry and immunohistochemistry (IHC) analyses. Pulmonary endothelial cells showed a prothrombotic and dysfunctional phenotype during COVID-19, especially a loss of thrombomodulin expression, an anticoagulant and anti-inflammatory molecule. In addition, macrophages, monocytes, and T cells were increased and activated in the lungs of COVID-19 patients. However, we found no evidence of SARS-CoV-2 infection in the pulmonary endothelium, while the alveolar epithelium exhibited severe SARS-CoV-2 infection and damage, causing hypoxia during COVD-19. We confirmed hypoxic stress on endothelial cells and immune cells in the COVID-19 patient lungs.

Methods

 Human samples

We obtained seven lung samples, seven heart samples, and eight kidney samples from nine autopsies of patients who died of COVID-19 at the Johns Hopkins University (Table 1 and Supplementary Table S1). We were unable to collect all three tissues from all patients for technical reasons. As a non-COVID-19 control, eight lung specimens, eleven heart specimens, and ten kidney specimens were obtained from autopsies of patients who had no respiratory, cardiovascular, and renal diseases. Samples were fixed for histology study or cryopreserved for flow cytometry and RT-PCR analysis. For cryopreservation, samples were stabilized in HypoThermosol FRS preservation media (Stemcell) and then frozen in CryoStor CS10 cryopreservation media (Stemcell) at -80°C.Table 1Clinical and demographic characteristics of patients.

COVID-19 patients
Number of patients9
% of males55·6% (5/9)
Age (years)66·1 ± 14·2
Ventilation (days)12·0 ± 12·3
ICU admission (days)9·9 ± 12·2
Hospitalization (days)10·6 ± 11·9
SaO2 (%)76·8 ± 17·5
CRP (mg/ml)55·1 ± 46·7
D-dimer (µg/ml)11·2 ± 12·0
Date of tissue collection4/7/2020 – 5/2/2020
Controls
Number of patients12
% of males41·7% (5/12)
Age (years)56·4 ± 18·9

 Ethics

Written informed consent was obtained from subjects, and the study protocol was approved by the Committee for the Protection of Human Subjects (IRB NA_00036610). This study was conducted in accordance with the guidelines in the Declaration of Helsinki.

 SARS-CoV-2 in vitro infection of primary human endothelial cells

Human coronary artery endothelial cells (HCAECs) were purchased from Cell Applications (300-05a). Cells were cultured in Human Meso Endo Growth Medium (Cell Applications). All infections were performed in a biosafety level (BSL) 3 environment. HCAECs were seeded onto 6-well plates at a density of 200,000 cells/well. At the time of infection, the growth medium was replaced with media containing SARS-CoV-2 (SARS-CoV-2/USA/DC-HP00007/2020, GISAID EPI_ISL_434688) at a multiplicity of infection (MOI) of 5 infectious units per cell.42 Cells were then incubated for 1 hour at 37°C. Media containing the virus was aspirated and replenished with fresh media. Samples were then incubated for 24 hours. Cell images were taken using an EVOS XL Core Imaging System (Invitrogen).

 Histology study

Lung, heart, and kidney tissues were paraffin-embedded, cut as 4-μm-thick sections, and stained with H&E. Coagulation abnormalities were graded by microscopic assessment of the severity of thrombus formation (graded from 0 to 5) and hemorrhage (from 0 to 5), and the sum of two item scores was shown. Infiltration was separately scored from 0 to 5. Grading was performed by two independent blinded investigators. Images were acquired on a BX43 microscope (Olympus) with a DS-Fi3 camera (Nikon) using NIS-Elements D Software (Nikon).

 Immunohistochemistry staining

The tissue slides were deparaffinized, rehydrated, and blocked with Duel Endogenous Enzyme Block (Dako). Tissue sections were probed with the primary antibody against fibrin (59D8; Millipore), CD42b (SP219; Abcam; RRID: AB_2814749), VWF (F8/86; Invitrogen; RRID: AB_11001165), PECAM-1 (JC/70A; Abcam; RRID: AB_307284), or SARS-CoV-2 spike S1 protein (007; Sino Biological; AB_2827979) and then treated with HRP-conjugated secondary goat anti-rabbit or anti-mouse IgG antibody (Leica). The slides were counterstained with 50% hematoxylin in water.

 Cell isolation from autopsy tissue samples

All procedures involving SARS-CoV-2-infected tissue samples were performed in a BSL3 environment until the virus is inactivated. Tissues were minced using a single-edge scalpel and then incubated with 5 mL of tissue digestion enzyme solution for 30 minutes at 37°C with intermittent vortexing. The digestion enzyme solution was prepared with 200 U/ml Collagenase (Worthington) for lungs or 400 U/ml for hearts and kidneys and supplemented with 50 U/ml DNase I (Worthington) and 50 U/ml Hyaluronidase (Sigma) for all organs. Cells were washed and filtered through 40 μm cell strainers (Falcon).

 Flow cytometry

Flow cytometry samples from autopsy tissues were prepared in a BSL3 condition. Single cells were first stained with Live/Dead Fixable Aqua (Invitrogen) and 7-AAD (BioLegend). After blocking with TruStain FcX (BioLegend; RRID: AB_2818986) and normal mouse serum (Invitrogen), cells were surface-stained with fluorochrome-conjugated antibodies (Supplementary Table S2). Cells were then fixed, permeabilized with Cyto-Fast Fix/Perm Buffer (BioLegend), and stained with antibodies against intracellular antigens (Supplementary Table S2). Sample acquisition was performed on a BD FACSymphony flow cytometer (BD) running FACSDiva (BD). Results were analysed using FlowJo software (BD).

 Real-time RT-PCR

Total RNA from tissues or cultured cells was extracted using RNeasy Plus Mini Kit (Qiagen) or TRIzol (Invitrogen) in a BSL3 environment. Single‐stranded cDNA was synthesized with iScript Reverse Transcription Supermix (Bio-Rad). Target genes were amplified using Power SYBR Green PCR Master Mix (Applied Biosystems), and real-time cycle thresholds were detected via MyiQ2 thermal cycler (Bio-Rad). Data were analysed by the 2−ΔΔCt method and were normalized to GAPDH or HPRT expression and then to biological controls.

 SARS-CoV-2 RNA detection

Total RNA extracted from autopsy tissues or cultured cells were reverse transcribed and then amplified with primers specific for SARS-CoV-2 nucleocapsid (N) genes. Primers for SARS-CoV-2 N1, SARS-CoV-2 N2, and internal control RNase P were obtained from the Integrated DNA Technologies. According to the Centers for Disease Control and Prevention (CDC) guideline, a test is considered positive for SARS-CoV-2 if both SARS-CoV-2 N1 and N2 markers show Ct value <40.

 Electron microscopy

The lung tissue was excised from patient autopsies and cut into 2–3 mm3 pieces, and immediately fixed in 2·5% glutaraldehyde (EM grade; Electron Microscopy Sciences) dissolved in 0·1 M Na cacodylate, pH 7·4, for 2 hours at room temperature. Samples were processed as previously described by the Electron Microscopy Laboratory in the Department of Pathology at the Johns Hopkins University School of Medicine before examination with an electron microscope CM120 (Philips) under 80 kV.43 Images were acquired with Image Capture Engine V602 (Advanced Microscopy Techniques).

 Mice

Wild-type male C57BL/6 mice (JAX 000664) were purchased from the Jackson Laboratory. Mice were housed in specific pathogen-free animal facilities at the Johns Hopkins University School of Medicine. Experiments were conducted with 6- to 10-week-old age-matched mice. For hypoxia exposure, mice were housed in Plexiglas hypoxia chambers with 10·0% O2 that had continuous airflow.44 The O2 concentration was monitored and controlled with a ProOx model 350 unit (BioSpherix) by infusion of N2. CO2 and ammonia were removed throughout the experiment. Control mice were exposed to normal room air (20·8% O2). On day 8 of hypoxia exposure, lungs were perfused with PBS and then harvested. All methods and protocols were approved by the Animal Care and Use Committee of the Johns Hopkins University.

 Statistics

GraphPad Prism 8 software was used for statistical analysis. Data shown are mean values of two or three independent experiments. Data were analysed using non-parametric Mann-Whitney test or Kruskal-Wallis test followed by post-hoc Dunn’s multiple comparison test. Statistically significant comparisons were represented by asterisks in figures: *P < 0·05; **P < 0·005; ***P < 0·0005. Actual P-values were reported in figure legends. Linear regression using Spearman correlation was tested for correlation analysis. Actual r- and P-values were reported in figures.

 Role of the funding source

The funding sources had no involvement in study design, data collection, data analyses, interpretation, or writing of this report.

Results

 Coagulation abnormality and infiltration are presented in the lungs of COVID-19 patients

Thromboembolism is the most common pathologic finding in the postmortem lungs of patients who died of COVID-19.45464748 Thrombus formation is also observed in the autopsy hearts and kidneys from COVID-19 patients.46,48,49 To test whether thrombosis is present in the lungs, hearts, and kidneys from COVID-19 patient autopsies collected at the Johns Hopkins University, we analysed histologic sections of those tissue specimens. Clinical characteristics of our COVID-19 patient cohort are summarized in Table 1 and Supplementary Table S1. We tested seven lung samples, seven heart samples, and eight kidney samples collected from the autopsy of COVID-19 patients. Many postmortem lungs from COVID-19 patients showed severe coagulation abnormalities such as vascular congestion, thrombus formation, and hemorrhage, which were not observed in non-COVID-19 autopsy controls (Fig. 1a and Supplementary Fig. S1a). Fibrin deposition normally occurs at the site of the damaged blood vessel during coagulation, however, excessive formation or impaired clearance of a fibrin clot contributes to thrombosis development in pathologic conditions and diseases.50 IHC studies confirmed the development of fibrin- and platelet-rich thrombi in the lungs of COVID-19 patients (Fig. 1b and c and Supplementary Fig. S1b and c). The autopsy hearts and kidneys of COVID-19 patients exhibited congested blood vessels and fibrin-rich thrombi (Fig. 1a and b and Supplementary Fig. S1a and b). However, the severity of coagulation abnormalities in the hearts and kidneys of COVID-19 patients was milder compared to the lungs (Fig. 1d). No platelet-rich thrombi were found in the hearts and kidneys of COVID-19 patient autopsies (data not shown). Elevated plasma D-dimer levels confirmed thrombosis development in our COVID-19 patient cohort tested upon their hospitalization (Fig. 1e). In addition, we found that the number of platelets in the COVID-19 patient lungs was increased compared with controls by using flow cytometry (Fig. 1f and Supplementary Fig. S2). The number of platelets in the COVID-19 patient lungs was correlated with the severity of hypercoagulability (Fig. 1g). In the hearts and kidneys, the platelet counts were comparable between COVID-19 and control groups (Fig. 1f).

Fig 1
Figure 1Hypercoagulable state and infiltration in lungs during COVID-19. (a) Representative images of H&E staining on postmortem lungs, hearts, and kidneys from COVID-19 patients and non-COVID-19 controls. Scale bars: 25 µm. (b) Representative images of immunohistochemistry (IHC) for fibrin (brown) on autopsy lungs, hearts, and kidneys of COVID-19 patients. Scale bars: 25 µm. (c) Representative image of IHC for CD42b (brown) on autopsy lungs of COVID-19 patients. Scale bar: 25 µm. (d) Coagulation abnormality score of COVID-19 patient lungs, hearts, and kidneys examined by H&E stain. Kruskal-Wallis test with Dunn’s multiple comparisons test was used for statistical analysis. P-values: 0·0260 (Lung vs Heart); 0·0201 (Lung vs Kidney); 0·9999 (Heart vs Kidney). (e) Plasma D-dimer level in COVID-19 patients upon their hospitalization. (f) Number of platelets in postmortem lungs, hearts, and kidneys from COVID-19 patients and controls determined by flow cytometry. Platelets were gated on EpCAMCD45PECAM-1dimCD41+ (Supplementary Fig. S2). Mann-Whitney test was used for statistical analysis. P-values: 0·0205 (Lung); 0·3845 (Heart); 0·9999 (Kidney). (g) Correlation between platelet number and coagulation abnormality score in autopsy lungs of COVID-19 patients. Spearman correlation was used for statistical analysis. (h) Number of CD45+ cells in COVID-19 patient lungs, hearts, and kidneys. CD45+ immune cells were gated on EpCAMCD45+ (Supplementary Fig. S2). Mann-Whitney test was used for statistical analysis. P-values: 0·0037 (Lung); 0·0122 (Heart); 0·8286 (Kidney). (i) Correlation between CD45+ infiltrating cell number examined by flow cytometry and infiltration score histologically graded in postmortem lungs of COVID-19 patients. Spearman correlation was used for statistical analysis. In COVID-19 patient group, seven lung samples, seven heart samples, and eight kidney samples were tested. In controls, eight lung samples, eleven heart samples, and ten kidney samples were used. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005.View Large ImageFigure ViewerDownload Hi-res imageDownload (PPT)

Severe infiltration was another histological finding in the lungs of COVID-19 patients (Fig. 1a). The number of CD45+ infiltrating cells was significantly increased in the COVID-19 patient lungs compared with controls (Fig. 1h and Supplementary Fig. S2). The number of CD45+ cells in the COVID-19 patient lungs analysed by flow cytometry was correlated with the infiltration score histologically graded (Fig. 1i). In contrast, CD45+ cell number was reduced in the hearts of COVID-19 patients compared to controls (Fig. 1h). The kidneys of COVID-19 patients presented a comparable number of CD45+ infiltrating cells to the controls (Fig. 1h). These results indicate that severe hypercoagulability and infiltration develop in the lungs of COVID-19 patients.

 Platelets and endothelial cells show a prothrombotic phenotype in COVID-19 patient lungs

To investigate if platelets are activated in the lungs of COVID-19 patients, causing a severe hypercoagulable state, we characterized the phenotype of platelets in the autopsy lungs, hearts, and kidneys. In COVID-19 patients, blood platelets showed upregulated P-selectin and other receptors contributing to aggregation and activation of platelets.51,52 Our flow cytometry analysis revealed that platelet P-selectin level was increased in the hearts but not in the lungs and kidneys from COVID-19 patient autopsies compared to controls (Fig. 2a and b). We also tested the platelet expression of PF4 (CXCL4), which promotes blood coagulation and is a target of autoantibodies causing heparin-induced thrombocytopenia.53 Platelet PF4 expression showed a significant increase in the lungs of COVID-19 patients compared to controls (Fig. 2c and d). PF4 level in the heart platelets was also increased in the COVID-19 group, although it was not significant (Fig. 2c and d). In the kidneys, no different PF4 level was found between groups (Fig. 2c and d). Platelet expression of other receptors such as CD40L, CD42b (glycoprotein Ibα), and CXCR4 in the COVID-19 autopsy tissues was comparable to controls (data not shown). In addition, we found a massive production of VWF, a procoagulant, in most COVID-19 patient autopsy tissues – lungs, hearts, and kidneys – using IHC staining (Fig. 2e and Supplementary Fig. S3). VWF was detected in the outer layer of the thrombus and CD31-expressing vascular endothelium in the COVID-19 patient tissues, suggesting VWF expression by both activated platelets and endothelial cells. In contrast, control tissues showed a low level of VWF production only in the vascular endothelial lining (Fig. 2e and Supplementary Fig. S3).

Fig 2
Figure 2Prothrombotic phenotype of platelets and endothelial cells in COVID-19 patient lungs. (a and b) Flow cytometry analysis (a) and frequencies (b) of P-selectin expression on platelets in lungs, hearts, and kidneys of COVID-19 patients and controls. Platelets were gated on EpCAMCD45PECAM-1dimCD41+ (Supplementary Fig. S2). Mann-Whitney test was used for statistical analysis. P-values: 0·2319 (Lung); 0·0003 (Heart); 0·0676 (Kidney). (c and d) Histograms (c) and mean fluorescent intensity (MFI) (d) of PF4 expression in platelets of lungs, hearts, and kidneys from COVID-19 patients and controls. Mann-Whitney test was used for statistical analysis. P-values: 0·0289 (Lung); 0·1422 (Heart); 0·7577 (Kidney). (e) Representative images of immunohistochemistry staining for VWF (red) and CD31 (brown) on lung, heart, and kidney sections from COVID-19 and control autopsies. Scale bars: 12·5 µm. (f) Histograms for thrombomodulin (TM) and endothelial protein C receptor (EPCR) expression on endothelial cells in postmortem lungs, hearts, and kidneys. Endothelial cells were gated on EpCAMCD45PECAM-1hiCD34+ (Supplementary Fig. S2). (g and h) MFI of endothelial TM (g) and EPCR (h) expression in lungs, hearts, and kidneys from COVID-19 and control autopsies. Mann-Whitney test was used for statistical analysis. P-values: 0·0022 (Lung); 0·2109 (Heart); 0·1728 (Kidney) (g). P-values: 0·0003 (Lung); 0·0556 (Heart); 0·5884 (Kidney) (h). In COVID-19 patient group, seven lung samples, seven heart samples, and eight kidney samples were tested. In controls, eight lung samples, eleven heart samples, and ten kidney samples were used. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005, ***P < 0·0005.View Large ImageFigure ViewerDownload Hi-res imageDownload (PPT)

Platelets exhibited an activated phenotype in the lungs, hearts, and kidneys of COVID-19 patients (Fig. 2a-e). However, severe hypercoagulability mainly developed in the lungs of our samples and barely in the hearts and kidneys (Fig. 1a-g). To test whether endothelial cells provide a prothrombotic environment specifically in the lungs during COVID-19, we examined a phenotype of endothelial cells in the lungs, hearts, and kidneys of COVID-19 patients using flow cytometry analysis. We found that endothelial cell expression of anticoagulants thrombomodulin and endothelial cell protein C receptor (EPCR) were significantly downregulated in the lung of COVID-19 patients compared with controls (Fig. 2f-h). Endothelial cells in the hearts of COVID-19 patients showed a slight decrease in thrombomodulin and EPCR expression, but it was not significant (Fig. 2f-h). The kidney endothelial cells expressed thrombomodulin and EPCR with no differences between groups (Fig. 2f-h). In hemostatic conditions, endothelial cells produce nitric oxide and prostaglandin I2 by utilizing the enzymes endothelial nitric oxide synthase (eNOS) and cyclooxygenase 2 (COX-2), respectively, to inhibit platelet aggregation and activation9. In the COVID-19 patient organs that we examined, endothelial cells expressed a comparable level of those enzymes to controls (data not shown). Collectively, endothelial cells represent a prothrombotic phenotype in COVID-19 patient lungs, leading to platelet recruitment and coagulation abnormalities.

 Macrophages, monocytes, and T cells are increased and activated in COVID-19 patient lungs

We found an increased CD45+ cell infiltration in the lungs of COVID-19 patients (Fig. 1a and h). To further investigate the immune profile of COVID-19 patient lungs, we analysed the number and phenotype of different immune cells in the autopsy lungs of COVID-19 patients using flow cytometry. Macrophages were the largest immune cell population in both COVID-19 and control lungs and significantly increased in the COVID-19 patients compared with controls (Fig. 3a and Supplementary Fig. S4a). Similarly, the number of monocytes, CD4+ T cells, and CD8+ T cells was increased in the lungs of COVID-19 patients, while the neutrophil count was comparable between COVID-19 and controls (Fig. 3a and Supplementary Fig. S4a). In the phenotype analysis, macrophages in the COVID-19 patient lungs exhibited downregulated HLA-DR expression compared to controls (Fig. 3b and c). CD40 expression on macrophages was increased in the COVID-19 patients, although it was not significant (Fig. 3b and d). Monocytes in the COVID-19 patient lungs revealed upregulated CD16 expression compared with controls but not significantly (Fig. 3e and f). In CD4+ T cells of the COVID-19 patient lungs, CCR7+CD45RA central memory T cells (TCM) were proportionally increased compared to controls, while CCR7CD45RA+ effector memory T cells re-expressing CD45RA (TEMRA) were decreased (Fig. 3g and h). However, the absolute number of most CD4+ T cell subsets including CCR7+CD45+ naïve T cells (TN), TCM cells, and CCR7CD45RA effector memory T cells (TEM) was significantly increased in the COVID-19 patient lungs compared to controls (Fig. 3i). Similarly, the number of CD4+ TEMRA cells was inclined in the COVID-19 patient, although there was no significance (Fig. 3i). The cell frequencies and numbers of CD8+ T cell subsets in the COVID-19 patient lungs exhibited similar results to CD4+ T cells (Supplementary Fig. S4b and c). These data demonstrate that immune cells such as macrophages, monocytes, and T cells are increased and activated in the lungs during COVID-19.

Fig 3
Figure 3Increased macrophages, monocytes, and T cells with activated phenotype in lungs of COVID-19 patients. (a) Numbers of macrophages (CD68+CD206+), monocytes (CD68+CD206CD169), CD4+ T cells (CD3+CD4+), CD8+ T cells (CD3+CD8+), and neutrophils (CD15+CD66b+) in autopsy lungs of COVID-19 patients and controls. The gating strategy for flow cytometry analysis was shown in Supplementary Fig. S4a. Mann-Whitney test was used for statistical analysis. P-values: 0·0041 (Macrophages); 0·0140 (Monocytes); 0·0205 (CD4+ T cells); 0·0093 (CD8+ T cells); 0·4634 (Neutrophils). (b) Histograms of macrophage HLA-DR and CD40 expression in COVID-19 patient lungs. (c and d) Mean fluorescent intensity (MFI) of HLA-DR expression (c) and frequency of CD40 expression (d) in lung macrophages from COVID-19 and control autopsies. Mann-Whitney test was used for statistical analysis. P-values: 0·0175 (c); 0·0541 (d). (e and f) Flow cytometry analysis (e) and frequency (f) of CD16 expression in monocytes of autopsy lungs from COVID-19 patients. Mann-Whitney test was used for statistical analysis. P-value: 0·1807. (g) CCR7 and CD45RA expression profiles of CD4+ T cells from COVID-19 and control lungs. (h and i) Frequencies of T cell subsets among CD4+ T cells (h) and absolute numbers (i) in postmortem lungs of COVID-19 patients and controls. Mann-Whitney test was used for statistical analysis. P-values: 0·6943 (CCR7+CD45RA+); 0·0289 (CCR7+CD45RA); 0·6126 (CCR7CD45RA); 0·0037 (CCR7CD45RA+) (h). P-values: 0·0037 (CCR7+CD45RA+); 0·0022 (CCR7+CD45RA); 0·0205 (CCR7CD45RA); 0·0721 (CCR7CD45RA+) (i). Seven COVID-19 lung samples and eight control lung samples were tested. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005.View Large ImageFigure ViewerDownload Hi-res imageDownload (PPT)

 Adhesion molecule expression is reduced in endothelial cells of COVID-19 patient lungs

We and others found increased immune cell infiltration in the autopsy lungs of COVID-19 patients (Fig. 3).45,48 Several researchers predicted that endothelial cell expression of adhesion molecules such as ICAM-1, VCAM-1, E-selectin, and P-selectin would be upregulated to recruit immune cells in COVID-19.8,17,18 We tested whether adhesion molecule expression on endothelial cells is altered in the postmortem lungs, hearts, and kidneys from COVID-19 patients. Unexpectedly, endothelial adhesion molecule expression was reduced in the COVID-19 patient lungs compared to controls (Fig. 4a-f). In the lungs, endothelial cells in both COVID-19 and control groups were positive for ICAM-1 (Fig. 4a and b). However, we found a significant downregulation of ICAM-1 in the COVID-19 lungs. Endothelial cells in the control lungs expressed a low level of VCAM-1, E-selectin, and P-selectin, while the COVID-19 lungs showed even lower expression of those molecules (Fig. 4a, c, d, e, and f). In the COVID-19 autopsy hearts, ICAM-1 and E-selectin were significantly downregulated compared to controls, while VCAM-1 and P-selectin expression levels showed no differences (Fig. 4a-f). Endothelial cells in the kidneys from COVID-19 patients expressed a comparable level of adhesion molecules to controls except VCAM-1 (Fig. 4a-f). VCAM-1 was significantly decreased in the COVID-19 kidneys (Fig. 4a and c).

Fig 4
Figure 4Downregulated adhesion molecules in lung endothelial cells during COVID-19. (a) Flow cytometry analysis of ICAM-1 and VCAM-1 expression on endothelial cells of postmortem lungs, hearts, and kidneys from COVID-19 patients and controls. (b) Mean fluorescent intensity (MFI) of ICAM-1 expression on endothelial cells in lungs, hearts, and kidneys from COVID-19 and control autopsies. Mann-Whitney test was used for statistical analysis. P-values: 0·0037 (Lung); 0·0154 (Heart); 0·2031 (Kidney). (c) Frequencies of VCAM-1+ cells among endothelial cells in postmortem lungs, hearts, and kidney specimens. Mann-Whitney test was used for statistical analysis. P-values: 0·0401 (Lung); 0·2463 (Heart); 0·0263 (Kidney). (d) Flow cytometry plots of E-selectin and P-selectin expression on endothelial cells of autopsy lungs, hearts, and kidneys from COVID-19 patients and controls. (e and f) MFI of E-selectin (e) and P-selectin (f) expression in endothelial cells from postmortem lungs, hearts, and kidneys. Mann-Whitney test was used for statistical analysis. P-values: 0·0059 (Lung); 0·0268 (Heart); 0·5726 (Kidney) (e). P-values: 0·0939 (Lung); 0·0853 (Heart); 0·8968 (Kidney) (f). (g and h) Correlation between endothelial ICAM-1 (g) or thrombomodulin (TM) (h) expression and infiltration score histologically evaluated. Spearman correlation was used for statistical analysis. In COVID-19 patient group, seven lung samples, seven heart samples, and eight kidney samples were tested. In controls, eight lung samples, eleven heart samples, and ten kidney samples were used. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005.View Large ImageFigure ViewerDownload Hi-res imageDownload (PPT)

In the COVID-19 patient lungs, a loss of ICAM-1 expression in endothelial cells was not associated with the severity of immune cell infiltration, suggesting that other mechanisms would be involved in the pulmonary infiltration (Fig. 4g). Decreased expression of thrombomodulin in endothelial cells was significantly correlated with an increased infiltration in the lungs of COVID-19 patients (Fig. 4h). Thrombomodulin plays an anti-inflammatory role as well as an anticoagulant function.11,12,54,55 These data suggest that endothelial cell dysfunction with downregulated adhesion molecules occurs in the lungs of COVID-19 patients and that downregulated thrombomodulin expression in endothelial cells is related to severe infiltration during COVID-19.

 SARS-CoV-2 does not directly infect endothelial cells or induce their dysfunction

To understand if SARS-CoV-2 directly infects endothelial cells leading to a prothrombotic and dysfunctional state, we first tested the presence of the SARS-CoV-2 genome in the lungs, hearts, and kidneys from COVID-19 patients. All COVID-19 lung specimens were positive for viral RNA in the RT-PCR test (Fig. 5a and Supplementary Fig. S5a). However, two of seven heart samples and six of eight kidney samples tested positive for SARS-CoV-2 RNA (Fig. 5a and Supplementary Fig. S5b and c). Second, we searched SARS-CoV-2 spike protein in the COVID-19 autopsy tissues using IHC stain to detect virus-infected cells. The spike protein was present in all SARS-CoV-2 RNA-positive tissue specimens and was located outside the vascular endothelial layer in the lungs, hearts, and kidneys of COVID-19 patients (Fig. 5b and Supplementary Fig. S5d). Significantly less viral antigen was detected in the heart and kidney tissues compared to the lungs. We could not detect SARS-CoV-2 virions in the vascular endothelium of COVID-19 patient lungs using electron microscopy but were able to identify virus-like particles in the epithelium (Fig. 5c). Those particles represented a SARS-CoV-2-specific ultrastructure described by Miller and Goldsmith.56 In addition, alveolar epithelial cells of the COVID-19 patients showed an abnormal ultrastructure, which has been observed in ciliary dyskinesia induced by respiratory syncytial virus (RSV) infection (Fig. 5c).57

Fig 5
Figure 5No endothelial cell infection or dysfunction by SARS-CoV-2. (a) Pie charts showing the presence of SARS-CoV-2 RNA tested by RT-PCR in postmortem lungs, hearts, and kidneys from COVID-19 patients. Each pie slice represents one patient. (b) Representative images of immunohistochemistry for SARS-CoV-2 spike protein (brown) on lungs, hearts, and kidneys from COVID-19 autopsies. Scale bars: 25 µm (top); 6·25 µm (bottom). Arrows indicate the blood vessel. (c) Representative images of SARS-CoV-2 in the pulmonary epithelium of COVID-19 patient. Scale bars: 2 µm (left); 200 nm (right). (d) Microscopic images of primary human coronary artery endothelial cells (HCAECs) with in vitro SARS-CoV-2 or mock infection. Images were acquired 16 hours after virus inoculation. Scale bars: 125 µm. (e) Frequencies of thrombomodulin (TM)+, ICAM-1+, and VCAM-1+ cells among HCAECs with in vitro SARS-CoV-2 or mock infection assessed by flow cytometry. Mann-Whitney test was used for statistical analysis. P-values: 0·7000 (TM); 0·9999 (ICAM-1); 0·9999 (VCAM-1). Seven lung samples, seven heart samples, and eight kidney samples were tested. Data shown are mean values of two or three independent experiments.View Large ImageFigure ViewerDownload Hi-res imageDownload (PPT)

Since endothelial cells of the COVID-19 patient lungs exhibited a prothrombotic and dysfunctional phenotype despite no signs of a direct SARS-CoV-2 infection, we hypothesized that endothelial cell dysfunction is not caused directly by the viral infection. To test this hypothesis, we infected HCAECs with SARS-CoV-2 in vitro. HCAECs showed no cytopathic changes at 16 hours post-infection (Fig. 5d). In the RT-PCR test, we found no detectable level of virus RNA in both SARS-CoV-2 and mock infection groups (Supplementary Fig. S5e). Importantly, no downregulation of thrombomodulin, ICAM-1, and VCAM-1 was shown in the SARS-CoV-2-infected HCAECs compared with mock infection control (Fig. 5e). Thus, we did not find evidence of direct endothelial cell infection by SARS-CoV-2 despite a dysfunctional phenotype of endothelial cells in COVID-19 patient lungs.

 Epithelial cells are involved in endothelial dysfunction, coagulation, and infiltration in COVID-19 patient lungs

Nasal epithelial cells and type II alveolar epithelial cells in the respiratory system are a primary target of SARS-CoV-2 entry, and the resultant lung injury can cause hypoxia in COVID-19.585960 Since we found the SARS-CoV-2 infection and ultrastructural damage in alveolar epithelial cells of COVID-19 patients (Fig. 5c), we tested whether a hypoxic injury occurred in the lungs. In our COVID-19 patient cohort, the average oxygen saturation before death was 76·8% and ranged between 49% and 100% (Fig. 6a, Table 1 and Supplementary Table S1). The mRNA level of HIF1A, a hypoxia-induced gene, was upregulated in the COVID-19 patient lungs compared to controls, while the hearts and kidneys showed no differences (Fig. 6b). In the COVID-19 patients, lung epithelial cells showed increased GLUT1 expression, another hypoxia-induced molecule, compared to controls, although epithelial HIF1α expression was comparable between COVID-19 and control groups (Fig. 6c). Endothelial cells and immune cells in the lungs of COVID-19 patients exhibited increased HIF1α and GLUT1 expression, confirming hypoxic stress in the lungs (Fig. 6d and Supplementary Fig. S6). To investigate if hypoxia can directly cause endothelial cell dysfunction in the lungs, we housed naïve mice in the hypoxia chamber with 10% O2 for eight days (Fig. 6e). In the hypoxia-exposed mice, the lung endothelial cells showed decreased thrombomodulin and ICAM-1 expression compared to the normal oxygen control, mimicking the dysfunctional phenotype of lung endothelial cells in the COVID-19 patients (Fig. 6f and Supplementary Fig. S7). Other adhesion molecules including VCAM-1, E-selectin, and P-selectin were expressed at comparable levels between the hypoxia and control groups (Fig. 6f and data not shown).

Fig 6
Figure 6Hypoxic stress induced by activated epithelial cells in lungs of COVID-19 patients. (a) Arterial oxygen saturation (SaO2) of COVID-19 patients. (b) HIF1A gene expression in autopsy lungs, hearts, and kidneys of COVID-19 patients and controls determined by RT-PCR. Mann-Whitney test was used for statistical analysis. P-values: 0·0089 (Lung); 0·5358 (Heart); 0·0553 (Kidney). (c) Mean fluorescent intensity (MFI) of HIF1α and GLUT1 expression on lung epithelial cells in COVID-19 autopsies and controls. Epithelial cells were gated on EpCAM+CD45 (Supplementary Fig. S2). Mann-Whitney test was used for statistical analysis. P-values: 0·3969 (left); 0·0003 (right). (d) MFI of HIF1α expression in endothelial cells and CD45+ immune cells in postmortem lungs of COVID-19 patients and controls. Mann-Whitney test was used for statistical analysis. P-values: 0·0541 (left); 0·0012 (right). (e) Scheme of experimental timeline for 10% O2 hypoxia exposure to C57BL/6 mice. Mice were sacrificed on day 8 of hypoxia. (f) MFI of thrombomodulin (TM), ICAM-1, and VCAM-1 expression on lung endothelial cells collected from mice exposed to hypoxia or normal level of oxygen. Mouse lung endothelial cells were gated on EpCAMCD45PECAM-1+CD34+ (Supplementary Fig. S7). P-values: 0·0079 (TM); 0·0079 (ICAM-1); 0·3095 (VCAM-1). (g) Flow cytometry analysis of tissue factor (TF) and ICAM-1 expression in pulmonary epithelial cells from COVID-19 patients and controls. (h) MFI of TF and ICAM-1 expression in epithelial cells of COVID-19 patient lungs. Mann-Whitney test was used for statistical analysis. P-value: 0·0059 (left); 0·2810 (right). (i) Correlation between epithelial TF expression and platelet number in lungs of COVID-19 patients. Spearman correlation was used for statistical analysis. In COVID-19 patient group, seven lung samples, seven heart samples, and eight kidney samples were tested. In controls, eight lung samples, eleven heart samples, and ten kidney samples were used. In mouse experiment, five mice were used for each group. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005; ***P < 0·0005.View Large ImageFigure ViewerDownload Hi-res imageDownload (PPT)

Next, we tested if damaged epithelial cells are directly involved in a hypercoagulable state and immune cell infiltration during COVID-19. We found that pulmonary epithelial cells in the COVID-19 patients expressed a higher level of tissue factor, a procoagulant, compared with controls (Fig. 6g and h). Elevated tissue factor expression level in epithelial cells was significantly associated with increased platelet number in the COVID-19 patient lungs (Fig. 6i). In addition, unlike endothelial cells, pulmonary epithelial cells exhibited an increase of ICAM-1 expression in the COVID-19 patients compared to controls, although it was not significant (Fig. 6g and h). These data suggest that pulmonary epithelial cells might participate in endothelial cell dysfunction, platelet activation and aggregation, and immune cell infiltration directly or indirectly via hypoxia during COVID-19.

Discussion

Many researchers have speculated about a phenotype of activated endothelial cells in COVID-19.8,17,18 However, direct evidence was missing. Most human studies on SARS-CoV-2 infection have used the blood or pharyngeal swab specimens that can be conveniently acquired but barely contain endothelial cells.61 In this study, we elucidated the dysfunctional phenotype of endothelial cells associated with hypercoagulability and severe infiltration in the lungs during fatal COVID-19. As previously shown by other researchers, we found severe coagulation abnormalities and infiltration in the autopsy lungs from COVID-19 patients.45464748 Pulmonary endothelial cells exhibited prothrombotic and dysfunctional phenotype with upregulated procoagulant and downregulated anticoagulants and adhesion molecules in COVID-19. However, we were unable to find clear evidence of direct endothelial cell infection by SARS-CoV-2 in the postmortem COVID-19 specimens and the primary cell in vitro culture. This suggests that endothelial cell dysfunction might not be directly caused by SARS-CoV-2 infection but rather induced by the pathophysiologic conditions during COVID-19. Unlike the lungs, the hearts and kidneys in COVID-19 showed a mild level of hypercoagulable state, infiltration, and endothelial cell dysfunction.Several researchers predicted activated and dysfunctional phenotype of endothelial cells showing strong expression of procoagulant VWF in COVID-19.17,18 Indeed, we found that VWF expression was strikingly increased in endothelial cells of the lungs, hearts, and kidneys from COVID-19 patients. The VWF is a multimeric glycoprotein secreted exclusively by endothelial cells and platelets into the circulation and involved in thrombus formation by tethering platelets to endothelial cells.9,10 In addition, we found the downregulation of anticoagulants thrombomodulin and EPCR in the lung endothelial cells of COVID-19 autopsies. Thrombomodulin is a transmembrane glycoprotein expressed mainly on endothelial cells but also found in immune cells, vascular smooth muscle cells, keratinocytes, and lung alveolar epithelial cells.9,11,12 Thrombomodulin is a potent anticoagulant that binds to thrombin and then deactivates it to initiate an anticoagulation cascade. Thrombin-thrombomodulin complex activates protein C, which is a crucial mediator to inactivate coagulant Factors Va and VIIIa and to further reduce thrombin generation in the anticoagulation processes.11,12,62 EPCR, another coagulant, presents the protein C to the thrombin-thrombomodulin complex to accelerate protein C activation.11,62 This suggests that endothelial cell dysfunction with increased procoagulant production and decreased anticoagulant expression is associated with severe thrombus formation and hypercoagulability in the lung of COVID-19 patients.In the hearts and kidneys of COVID-19 patients, platelets showed an activated phenotype with increased P-selectin, PF4, and VWF expression at a similar level as the lungs. Furthermore, endothelial cells largely produced VWF to a similar degree in the lungs, hearts, and kidneys in the COVID-19 patients. However, the hearts and kidneys from COVID-19 patients exhibited only mild coagulation abnormalities, while the lungs developed fatal thrombosis. This suggests that platelet activation and partial dysfunction of endothelial cells are insufficient for developing a severe multi-organ hypercoagulable state during COVID-19. A loss of thrombomodulin and EPCR expression on endothelial cells seems critical for the progression to fatal thrombotic disorders. Previous studies showed that endothelial expression of those anticoagulant molecules is downregulated in thrombosis associated with meningococcal sepsis, purpura fulminans, and cerebral malaria.636465 In mice, endothelial cell-specific thrombomodulin deletion resulted in lethal thrombosis development.66 We found a significant loss of thrombomodulin and EPCR expression in the lungs of COVID-19 patients. In contrast, those anticoagulants in the hearts and kidneys were relatively preserved. Recently, decreased endothelial thrombomodulin and ECPR expression in COVID-19 patient lungs was reported by using IHC stain.67In COVID-19 patients, it is reported that the plasma level of soluble thrombomodulin was elevated.13,15,16 Since pathophysiological conditions such as infection, sepsis, inflammation, and ischemic disease can cause thrombomodulin release from the endothelial cell membrane into the blood, soluble thrombomodulin is considered as a useful biomarker to diagnose endothelial damage and dysfunction.11,12 Meanwhile, those disease conditions can downregulate endothelial thrombomodulin mRNA transcription through hypoxic stress and pro-inflammatory cytokine milieu.11 We found hypoxic stress in the endothelial cells of COVID-19 patient lungs. Thus, our finding of decreased thrombomodulin expression on endothelial cells in COVID-19 might be mediated by two regulation processes – release in a soluble form and downregulation of gene transcription.We elucidated an increase of macrophages, monocytes, CD4+ T cells, and CD8+ T cells in the postmortem lungs of COVID-19 patients using flow cytometry analysis, while other researchers have mainly utilized IHC or scRNA-seq analysis.686970 In our analysis, lung macrophages of COVID-19 patients expressed a lower level of HLA-DR and a higher level of CD40 than those of controls. Severe COVID-19 patients showed an increased number of HLA-DRlo monocytes in the blood, a dysregulated or immunosuppressive subset of monocytes.25,717273 This suggests that abnormally or alternatively activated macrophages emerged in the lungs during COVID-19. The role of CD40 expressed on macrophages in COVID-19 remains unclear. However, SARS-CoV-2 infection increased CD40 expression and decreased HLA-DR expression in M2 macrophages in vitro, supporting our findings on the lung macrophages of COVID-19 patients.74 Our data revealed an increase of CD16+ inflammatory monocytes in the autopsy lungs of COVID-19 patients. Some studies showed an elevated number of CD14CD16+ non-classical monocytes in COVID-19 patient blood, while others disagreed.25,71,75,76 Additionally, we showed that the number of CD4+ TEM and CD8+ TEM cells was significantly increased in the COVID-19 patient lungs. In agreement with this, other studies showed an elevated frequency or number of activated T cells in the blood during COVID-19, although they have used different markers to define T cell subsets than ours.77,78In SARS-CoV-2 infection, it is speculated that vascular endothelial cells exhibit increased expression of adhesion molecules.8,17,18 ICAM-1 upregulation is generally mediated by various biological stimuli such as inflammation, bacteria or virus infection, and oxidative stress.79 Endothelial cell expression of other adhesion molecules including VCAM-1, E-selectin, and P-selectin would be increased similarly to ICAM-1 upregulation. In pathologic conditions, adhesion molecules anchored in endothelial cell transmembrane are cleaved by matrix metalloproteinases (MMPs) and released into the blood.31 Several studies showed that soluble forms of ICAM-1, VCAM-1, E-selectin, and P-selectin are highly elevated in severely ill COVID-19 patients.13,14 In our flow cytometry analysis, the lung endothelial cells in COVID-19 exhibited reduced expression of surface ICAM-1, VCAM-1, E-selectin, and P-selectin despite increased pulmonary infiltration. It is unclear why adhesion molecule expression was decreased in activated endothelial cells in COVID-19. Since CD45+ cell number was increased in the lungs of COVID-19 patients, it could be hypothesized that endothelial cells upregulate adhesion molecules upon SARS-CoV-2 infection to recruit immune cells from the blood, and then the adhesion molecule expression is downregulated after the infiltration. As a possible cause of downregulation, activated endothelial cells release endothelial microparticles (EMPs), which induce ICAM-1 downregulation on adjacent endothelial cells.80 In addition, COVID-19 patients showed excessively increased plasma MMP level and enzymatic activity, suggesting the massive cleavage of membrane-bound adhesion molecules on endothelial cells.818283 More studies will be necessary to determine how ICAM-1 and other adhesion molecules are downregulated on endothelial cells after the immune cell infiltration in COVID-19. Meanwhile, we found that decreased endothelial thrombomodulin expression was significantly associated with increased infiltration in the lungs of COVID-19 patients, while ICAM-1 expression showed no correlation. Thrombomodulin plays an anti-inflammatory role besides its anticoagulant function by inhibiting immune cell adhesion and suppressing complement system activation.11,12,54,55 Thus, a loss of thrombomodulin expression on endothelial cells could trigger or accelerate a pathogenic infiltration of immune cells into the lungs during COVID-19.Since we observed prothrombotic endothelial cells with downregulated adhesion molecule expression in the COVID-19 patient lungs, we wanted to examine the cause of endothelial cell dysfunction. We investigated the hypothesis that SARS-CoV-2 directly infects endothelial cells leading to dysfunction. Although we detected SARS-CoV-2 RNA by RT-PCR test in all autopsy lung specimens with COVID-19, the spike protein was not seen in endothelial cells by IHC stain. We found the SARS-CoV-2 particles in the pulmonary epithelium but not in the endothelium using electron microscopy. Interestingly, the SARS-CoV-2 virions were mostly located around the ciliated epithelium, where highly expresses ACE2 and TMPRSS2 allowing a strong SARS-CoV-2 infection.22 In addition, we were unable to infect endothelial cells in vitro with SARS-CoV-2 or induce endothelial cell dysfunction by the virus infection. Thus, endothelial cells are unlikely to be infected directly by SARS-CoV-2. Several recent studies showed the low susceptibility of endothelial cells to SARS-CoV-2 due to low ACE2 expression.35363738394041 This is supported by the fact that recombinant ACE2 transduction is required for sufficient SARS-CoV-2 infection of endothelial cells.40 Some researchers published electron microscope studies showing SARS-CoV-2 particles present in endothelial cells.33,47 However, others clarified that those shown particles were indeed cellular ribosomal complexes or vesicles.34,84We sought a pathophysiological condition in SARS-CoV-2 infection that can mediate endothelial cell dysfunction. We found a low oxygen saturation in the COVID-19 patients of our cohort and an increased HIF1A gene expression in their lungs, supporting hypoxic stress during COVID-19. The protein levels of HIF1α or GLUT1 were upregulated in epithelial cells, endothelial cells, and immune cells of COVID-19 patient lungs, confirming a hypoxic condition in SARS-CoV-2-infected lungs. Hypoxia can cause endothelial cell dysfunction and further thrombosis.85,86 In our mouse experiment, hypoxia induced endothelial cell dysfunction exhibiting thrombomodulin and ICAM-1 downregulation, which we observed in the COVID-19 patient lungs. Thus, pulmonary epithelial cells infected and damaged by SARS-CoV-2 could cause hypoxia leading to endothelial dysfunction in COVID-19 patients. Wang et al. reported that epithelial cells were more susceptible to SARS-CoV-2 infection compared with endothelial cells in their in vitro co-culture system.38 Co-cultured endothelial cells in the system revealed altered proteomic profile despite no direct SARS-CoV-2 infection, suggesting crosstalk between SARS-CoV-2-infected epithelial cells and endothelial cells. In addition, we found an increased expression of tissue factor and ICAM-1 in pulmonary epithelial cells of COVID-19 patients. Thus, injured epithelial cells may be directly involved in thrombosis and infiltration in the lungs of COVID-19 patients. Alternatively, microparticles derived from platelets, monocytes, and endothelial cells can act as a procoagulant contributing to thrombosis development.87 Elevated circulating microparticles have been reported in COVID-19 patients, suggesting a pathogenic role of microparticles in COVID-19-related immunothrombosis.88 Future studies should explore whether microparticles are involved in thrombosis development and further endothelial cell dysfunction in the lungs of COVID-19 patients.Microbial infection can be a common cause of inflammation-involved thrombosis.89 Notably, one-half of COVID-19 non-survivors revealed a secondary infection diagnosed by clinical symptoms, signs, or culture tests in Wuhan.90 In our COVID-19 patient cohort, only two revealed a suspected secondary infection tested by sputum culture, suggesting thrombosis mainly caused by primary SARS-CoV-2 infection or following pathologic conditions.In this study, we demonstrated the downregulation of endothelial thrombomodulin in the COVID-19 autopsies and suggested a prominent role of thrombomodulin in preventing severe thrombosis and infiltration. In murine models, overexpression of human thrombomodulin or the engineered preservation of thrombomodulin expression protected the lungs from thrombotic disorders and inflammation.54,91 Moreover, recombinant thrombomodulin or thrombomodulin analog Solulin treatment ameliorated the infarct volume in ischemic stroke.92,93 Recombinant soluble thrombomodulin administration was also beneficial in patients with disseminated intravascular coagulation and different inflammatory diseases.12,55,94 Based on our finding of endothelial thrombomodulin downregulation in COVID-19, recombinant thrombomodulin treatment could be considered as a potential therapy.In conclusion, we found a procoagulant phenotype of endothelial cells with upregulated VWF and downregulated thrombomodulin and EPCR in the lungs and, to a lesser extent, hearts and kidneys during COVID-19. The number of macrophages, monocytes, and T cells was increased in the lungs of COVID-19 patients, and all of them exhibited activated phenotype. Despite increased infiltration, endothelial cell expression of ICAM-1, VCAM-1, E-selectin, and P-selectin was downregulated in the lungs of COVID-19 patients. However, decreased thrombomodulin expression in endothelial cells was related to increased infiltration in the COVID-19 patient lungs. We showed that endothelial cell dysfunction was not caused directly by SARS-CoV-2 infection. Interestingly, we found pulmonary epithelial cell infection and damage by SARS-CoV-2 in the COVID-19 patients. Hypoxia caused by infected epithelial cells would lead to endothelial cell dysfunction in COVID-19. The main limitation of our study is the small sample size. In addition, since we did not have access to biopsy samples of COVID-19 patients, our study was conducted using autopsy specimens. Further studies confirming our findings in fresh biopsy samples from COVID-19 patients are needed.

Contributors

T.W. and D.C. conceptualized study. J.S., C.R.P., A.Z.R., and J.E.H. collected autopsy samples. T.W., M.K.W., D.M.H., M.V.T., Z.M., J.T.S., B.A., I.C., and A.P. performed experiments and analysed data. I.C., A.P., and D.C. verified the underlying data. A.P. supervised BSL3 experiments. T.W. and D.C. wrote the manuscript. E.G., M.K.H., A.G.H., D.-H.K., I.C., R.A.J., N.A.G., J.E.H., A.P., and D.C. reviewed the manuscript and provided intellectual input. All authors reviewed and approved the final version of the manuscript. All authors had full access to all the data in this study and accepted the responsibility to submit for publication.

Declaration of interests

None exist.

Acknowledgments

This work was supported by the Johns Hopkins COVID-19 Research Response Program to D.C. and C.R.P., the COVID-19 Rapid Response Grant 814664 from the American Heart Association (AHA) to D.C., the National Institutes of Health (NIH) Johns Hopkins Center of Excellence in Influenza Research and Surveillance HHSN272201400007C to A.P., the NIH/National Heart, Lung, and Blood Institute (NHLBI) R01HL122606 and NIH/National Institute of General Medicine Sciences (NIGMS) R01GM110674 to R.A.J., and the NIH/National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK) Supplemental Award R01DK093770-09S1 for COVID-19 Research to C.R.P. D.C. is supported by the NIH/NHLBI R01HL118183 and R01HL136586, and AHA 20TPA35490421 and 19TPA34910007. D.C. and N.A.G. are supported by the Matthew Vernon Poyner Memorial Foundation. J.E.H. received support from NIH/National Cancer Institute (NCI) P30CA006973, Cancer Clinical Care support grant and P50CA06924, supplement to a GI SPORE. N.A.G. is supported by the AHA 20SFRN35380046 and NIH/NHLBI R01HL118183. D.-H.K is supported by the NIH/National Institute of Allergy and Infectious Diseases (NIAID) R01AI143773. A.G.H. is supported by NIH/NHLBI R01HL147660. T.W. is supported by the 2018 Rhett Lundy Memorial Research Fellowship from the Myocarditis Foundation. M.K.W. is funded by the NIH/National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS) F31AR077406. D.M.H. is funded by the NIH/NHLBI F31HL149328. The funding sources had no involvement in study design, data collection, analyses, interpretation, or writing of this report.

Data sharing statement

All data reported in this article and supplementary materials are available for sharing by requesting to the corresponding author.

Appendix. Supplementary materials

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Published: January 12, 2022Accepted: December 30, 2021Received in revised form: November 28, 2021Received: August 18, 2021

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DOI: https://doi.org/10.1016/j.ebiom.2022.103812

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Figures

  • Fig 1Figure 1Hypercoagulable state and infiltration in lungs during COVID-19. (a) Representative images of H&E staining on postmortem lungs, hearts, and kidneys from COVID-19 patients and non-COVID-19 controls. Scale bars: 25 µm. (b) Representative images of immunohistochemistry (IHC) for fibrin (brown) on autopsy lungs, hearts, and kidneys of COVID-19 patients. Scale bars: 25 µm. (c) Representative image of IHC for CD42b (brown) on autopsy lungs of COVID-19 patients. Scale bar: 25 µm. (d) Coagulation abnormality score of COVID-19 patient lungs, hearts, and kidneys examined by H&E stain. Kruskal-Wallis test with Dunn’s multiple comparisons test was used for statistical analysis. P-values: 0·0260 (Lung vs Heart); 0·0201 (Lung vs Kidney); 0·9999 (Heart vs Kidney). (e) Plasma D-dimer level in COVID-19 patients upon their hospitalization. (f) Number of platelets in postmortem lungs, hearts, and kidneys from COVID-19 patients and controls determined by flow cytometry. Platelets were gated on EpCAMCD45PECAM-1dimCD41+ (Supplementary Fig. S2). Mann-Whitney test was used for statistical analysis. P-values: 0·0205 (Lung); 0·3845 (Heart); 0·9999 (Kidney). (g) Correlation between platelet number and coagulation abnormality score in autopsy lungs of COVID-19 patients. Spearman correlation was used for statistical analysis. (h) Number of CD45+ cells in COVID-19 patient lungs, hearts, and kidneys. CD45+ immune cells were gated on EpCAMCD45+ (Supplementary Fig. S2). Mann-Whitney test was used for statistical analysis. P-values: 0·0037 (Lung); 0·0122 (Heart); 0·8286 (Kidney). (i) Correlation between CD45+ infiltrating cell number examined by flow cytometry and infiltration score histologically graded in postmortem lungs of COVID-19 patients. Spearman correlation was used for statistical analysis. In COVID-19 patient group, seven lung samples, seven heart samples, and eight kidney samples were tested. In controls, eight lung samples, eleven heart samples, and ten kidney samples were used. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005.
  • Fig 2Figure 2Prothrombotic phenotype of platelets and endothelial cells in COVID-19 patient lungs. (a and b) Flow cytometry analysis (a) and frequencies (b) of P-selectin expression on platelets in lungs, hearts, and kidneys of COVID-19 patients and controls. Platelets were gated on EpCAMCD45PECAM-1dimCD41+ (Supplementary Fig. S2). Mann-Whitney test was used for statistical analysis. P-values: 0·2319 (Lung); 0·0003 (Heart); 0·0676 (Kidney). (c and d) Histograms (c) and mean fluorescent intensity (MFI) (d) of PF4 expression in platelets of lungs, hearts, and kidneys from COVID-19 patients and controls. Mann-Whitney test was used for statistical analysis. P-values: 0·0289 (Lung); 0·1422 (Heart); 0·7577 (Kidney). (e) Representative images of immunohistochemistry staining for VWF (red) and CD31 (brown) on lung, heart, and kidney sections from COVID-19 and control autopsies. Scale bars: 12·5 µm. (f) Histograms for thrombomodulin (TM) and endothelial protein C receptor (EPCR) expression on endothelial cells in postmortem lungs, hearts, and kidneys. Endothelial cells were gated on EpCAMCD45PECAM-1hiCD34+ (Supplementary Fig. S2). (g and h) MFI of endothelial TM (g) and EPCR (h) expression in lungs, hearts, and kidneys from COVID-19 and control autopsies. Mann-Whitney test was used for statistical analysis. P-values: 0·0022 (Lung); 0·2109 (Heart); 0·1728 (Kidney) (g). P-values: 0·0003 (Lung); 0·0556 (Heart); 0·5884 (Kidney) (h). In COVID-19 patient group, seven lung samples, seven heart samples, and eight kidney samples were tested. In controls, eight lung samples, eleven heart samples, and ten kidney samples were used. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005, ***P < 0·0005.
  • Fig 3Figure 3Increased macrophages, monocytes, and T cells with activated phenotype in lungs of COVID-19 patients. (a) Numbers of macrophages (CD68+CD206+), monocytes (CD68+CD206CD169), CD4+ T cells (CD3+CD4+), CD8+ T cells (CD3+CD8+), and neutrophils (CD15+CD66b+) in autopsy lungs of COVID-19 patients and controls. The gating strategy for flow cytometry analysis was shown in Supplementary Fig. S4a. Mann-Whitney test was used for statistical analysis. P-values: 0·0041 (Macrophages); 0·0140 (Monocytes); 0·0205 (CD4+ T cells); 0·0093 (CD8+ T cells); 0·4634 (Neutrophils). (b) Histograms of macrophage HLA-DR and CD40 expression in COVID-19 patient lungs. (c and d) Mean fluorescent intensity (MFI) of HLA-DR expression (c) and frequency of CD40 expression (d) in lung macrophages from COVID-19 and control autopsies. Mann-Whitney test was used for statistical analysis. P-values: 0·0175 (c); 0·0541 (d). (e and f) Flow cytometry analysis (e) and frequency (f) of CD16 expression in monocytes of autopsy lungs from COVID-19 patients. Mann-Whitney test was used for statistical analysis. P-value: 0·1807. (g) CCR7 and CD45RA expression profiles of CD4+ T cells from COVID-19 and control lungs. (h and i) Frequencies of T cell subsets among CD4+ T cells (h) and absolute numbers (i) in postmortem lungs of COVID-19 patients and controls. Mann-Whitney test was used for statistical analysis. P-values: 0·6943 (CCR7+CD45RA+); 0·0289 (CCR7+CD45RA); 0·6126 (CCR7CD45RA); 0·0037 (CCR7CD45RA+) (h). P-values: 0·0037 (CCR7+CD45RA+); 0·0022 (CCR7+CD45RA); 0·0205 (CCR7CD45RA); 0·0721 (CCR7CD45RA+) (i). Seven COVID-19 lung samples and eight control lung samples were tested. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005.
  • Fig 4Figure 4Downregulated adhesion molecules in lung endothelial cells during COVID-19. (a) Flow cytometry analysis of ICAM-1 and VCAM-1 expression on endothelial cells of postmortem lungs, hearts, and kidneys from COVID-19 patients and controls. (b) Mean fluorescent intensity (MFI) of ICAM-1 expression on endothelial cells in lungs, hearts, and kidneys from COVID-19 and control autopsies. Mann-Whitney test was used for statistical analysis. P-values: 0·0037 (Lung); 0·0154 (Heart); 0·2031 (Kidney). (c) Frequencies of VCAM-1+ cells among endothelial cells in postmortem lungs, hearts, and kidney specimens. Mann-Whitney test was used for statistical analysis. P-values: 0·0401 (Lung); 0·2463 (Heart); 0·0263 (Kidney). (d) Flow cytometry plots of E-selectin and P-selectin expression on endothelial cells of autopsy lungs, hearts, and kidneys from COVID-19 patients and controls. (e and f) MFI of E-selectin (e) and P-selectin (f) expression in endothelial cells from postmortem lungs, hearts, and kidneys. Mann-Whitney test was used for statistical analysis. P-values: 0·0059 (Lung); 0·0268 (Heart); 0·5726 (Kidney) (e). P-values: 0·0939 (Lung); 0·0853 (Heart); 0·8968 (Kidney) (f). (g and h) Correlation between endothelial ICAM-1 (g) or thrombomodulin (TM) (h) expression and infiltration score histologically evaluated. Spearman correlation was used for statistical analysis. In COVID-19 patient group, seven lung samples, seven heart samples, and eight kidney samples were tested. In controls, eight lung samples, eleven heart samples, and ten kidney samples were used. Data shown are mean values of two or three independent experiments. *P < 0·05; **P < 0·005.
  • Fig 5Figure 5No endothelial cell infection or dysfunction by SARS-CoV-2. (a) Pie charts showing the presence of SARS-CoV-2 RNA tested by RT-PCR in postmortem lungs, hearts, and kidneys from COVID-19 patients. Each pie slice represents one patient. (b) Representative images of immunohistochemistry for SARS-CoV-2 spike protein (brown) on lungs, hearts, and kidneys from COVID-19 autopsies. Scale bars: 25 µm (top); 6·25 µm (bottom). Arrows indicate the blood vessel. (c) Representative images of SARS-CoV-2 in the pulmonary epithelium of COVID-19 patient. Scale bars: 2 µm (left); 200 nm (right). (d) Microscopic images of primary human coronary artery endothelial cells (HCAECs) with in vitro SARS-CoV-2 or mock infection. Images were acquired 16 hours after virus inoculation. Scale bars: 125 µm. (e) Frequencies of thrombomodulin (TM)+, ICAM-1+, and VCAM-1+ cells among HCAECs with in vitro SARS-CoV-2 or mock infection assessed by flow cytometry. Mann-Whitney test was used for statistical analysis. P-values: 0·7000 (TM); 0·9999 (ICAM-1); 0·9999 (VCAM-1). Seven lung samples, seven heart samples, and eight kidney samples were tested. Data shown are mean values of two or three independent experiments.
  • Fig 6Figure 6Hypoxic stress ind

How does coronavirus kill? Clinicians trace a ferocious rampage through the body, from brain to toes

Authors: By Meredith WadmanJennifer Couzin-FrankelJocelyn KaiserCatherine MatacicApr. 17, 2020 , 6:45 PM

On rounds in a 20-bed intensive care unit one recent day, physician Joshua Denson assessed two patients with seizures, many with respiratory failure and others whose kidneys were on a dangerous downhill slide. Days earlier, his rounds had been interrupted as his team tried, and failed, to resuscitate a young woman whose heart had stopped. All shared one thing, says Denson, a pulmonary and critical care physician at the Tulane University School of Medicine. “They are all COVID positive.”

As the number of confirmed cases of COVID-19 surges past 2.2 million globally and deaths surpass 150,000, clinicians and pathologists are struggling to understand the damage wrought by the coronavirus as it tears through the body. They are realizing that although the lungs are ground zero, its reach can extend to many organs including the heart and blood vessels, kidneys, gut, and brain.

“[The disease] can attack almost anything in the body with devastating consequences,” says cardiologist Harlan Krumholz of Yale University and Yale-New Haven Hospital, who is leading multiple efforts to gather clinical data on COVID-19. “Its ferocity is breathtaking and humbling.”

Understanding the rampage could help the doctors on the front lines treat the fraction of infected people who become desperately and sometimes mysteriously ill. Does a dangerous, newly observed tendency to blood clotting transform some mild cases into life-threatening emergencies? Is an overzealous immune response behind the worst cases, suggesting treatment with immune-suppressing drugs could help? What explains the startlingly low blood oxygen that some physicians are reporting in patients who nonetheless are not gasping for breath? “Taking a systems approach may be beneficial as we start thinking about therapies,” says Nilam Mangalmurti, a pulmonary intensivist at the Hospital of the University of Pennsylvania (HUP).

What follows is a snapshot of the fast-evolving understanding of how the virus attacks cells around the body, especially in the roughly 5% of patients who become critically ill. Despite the more than 1000 papers now spilling into journals and onto preprint servers every week, a clear picture is elusive, as the virus acts like no pathogen humanity has ever seen. Without larger, prospective controlled studies that are only now being launched, scientists must pull information from small studies and case reports, often published at warp speed and not yet peer reviewed. “We need to keep a very open mind as this phenomenon goes forward,” says Nancy Reau, a liver transplant physician who has been treating COVID-19 patients at Rush University Medical Center. “We are still learning.”

The infection begins

When an infected person expels virus-laden droplets and someone else inhales them, the novel coronavirus, called SARS-CoV-2, enters the nose and throat. It finds a welcome home in the lining of the nose, according to a preprint from scientists at the Wellcome Sanger Institute and elsewhere. They found that cells there are rich in a cell-surface receptor called angiotensin-converting enzyme 2 (ACE2). Throughout the body, the presence of ACE2, which normally helps regulate blood pressure, marks tissues vulnerable to infection, because the virus requires that receptor to enter a cell. Once inside, the virus hijacks the cell’s machinery, making myriad copies of itself and invading new cells.

As the virus multiplies, an infected person may shed copious amounts of it, especially during the first week or so. Symptoms may be absent at this point. Or the virus’ new victim may develop a fever, dry cough, sore throat, loss of smell and taste, or head and body aches.

If the immune system doesn’t beat back SARS-CoV-2 during this initial phase, the virus then marches down the windpipe to attack the lungs, where it can turn deadly. The thinner, distant branches of the lung’s respiratory tree end in tiny air sacs called alveoli, each lined by a single layer of cells that are also rich in ACE2 receptors.

Normally, oxygen crosses the alveoli into the capillaries, tiny blood vessels that lie beside the air sacs; the oxygen is then carried to the rest of the body. But as the immune system wars with the invader, the battle itself disrupts this healthy oxygen transfer. Front-line white blood cells release inflammatory molecules called chemokines, which in turn summon more immune cells that target and kill virus-infected cells, leaving a stew of fluid and dead cells—pus—behind. This is the underlying pathology of pneumonia, with its corresponding symptoms: coughing; fever; and rapid, shallow respiration (see graphic). Some COVID-19 patients recover, sometimes with no more support than oxygen breathed in through nasal prongs.

But others deteriorate, often quite suddenly, developing a condition called acute respiratory distress syndrome (ARDS). Oxygen levels in their blood plummet and they struggle ever harder to breathe. On x-rays and computed tomography scans, their lungs are riddled with white opacities where black space—air—should be. Commonly, these patients end up on ventilators. Many die. Autopsies show their alveoli became stuffed with fluid, white blood cells, mucus, and the detritus of destroyed lung cells.

For More Information: https://www.sciencemag.org/news/2020/04/how-does-coronavirus-kill-clinicians-trace-ferocious-rampage-through-body-brain-toes

Skin rash should be considered as a fourth key sign of COVID-19

May 22, 2021

Data from the COVID Symptom Study shows that characteristic skin rashes and ‘COVID fingers and toes’ should be considered as key diagnostic signs of the disease, and can occur in the absence of any other symptoms. 

The COVID Symptom Study, led by researchers from King’s College London and health science company ZOE, asks participants to log their health and any new potential symptoms of COVID-19 on a daily basis. After noticing that a number of participants were reporting unusual skin rashes, the researchers focused on data from around 336,000 regular UK app users. 

Researchers discovered that 8.8% of people reporting a positive coronavirus swab test had experienced a skin rash as part of their symptoms, compared with 5.4% of people with a negative test result. Similar results were seen in a further 8.2% of users with a rash who did not have a coronavirus test, but still reported classic COVID-19 symptoms, such as cough, fever or anosmia (loss of smell).

To investigate further, the team set up a separate online survey, gathering images and information from nearly 12,000 people with skin rashes and suspected or confirmed COVID-19. The team particularly sought images from people of colour, who are currently under-represented in dermatology resources. Thank you to all who submitted photographs of their rashes.

17% of respondents testing positive for coronavirus reported a rash as the first symptom of the disease. And for one in five people (21%) who reported a rash and were confirmed as being infected with coronavirus, the rash was their only symptom.

The rashes associated with COVID-19 fall into three categories: 

  • Hive-type rash (urticaria): Sudden appearance of raised bumps on the skin which come and go quite quickly over hours and are usually very itchy. It can involve any part of the body, and often starts with intense itching of the palms or soles, and can cause swelling of the lips and eyelids. These rashes can present quite early on in the infection, but can also last a long time afterwards.
  • ‘Prickly heat’ or chickenpox-type rash (erythemato-papular or erythemato-vesicular rash): Areas of small, itchy red bumps that can occur anywhere on the body, but particularly the elbows and knees as well as the back of the hands and feet. The rash can persist for days or weeks.
  • COVID fingers and toes (chilblains): Reddish and purplish bumps on the fingers or toes, which may be sore but not usually itchy. This type of rash is most specific to COVID-19, is more common in younger people with the disease, and tends to present later on.

For More Information: https://covid.joinzoe.com/us-post/skin-rash-covid

Covid-19 Vaccine Analysis: The most common adverse events reported so far

Authors: DATED: AUGUST 6, 2021 BY SHARYL ATTKISSON 

As of July 19, 2021 there were 419,513 adverse event reports associated with Covid-19 vaccination in the U.S., with a total of 1,814,326 symptoms reported. That’s according to the federal Vaccine Adverse Event Reporting System (VAERS) database.

Report an adverse event after vaccination online here.

Each symptom reported does not necessarily equal one patient. Adverse event reports often include multiple symptoms for a single patient.

Reporting of illnesses and symptoms that occur after Covid-19 vaccination does not necessarily mean they were caused by the vaccine. The system is designed to collect adverse events that occur after vaccination to uncover any patterns of illnesses that were not captured during vaccine studies.

Read CDC info on Covid-19 vaccine here.

Scientists have estimated that adverse events occur at a rate many fold higher than what is reported in VAERS, since it is assumed that most adverse events are not reported through the tracking system. Reports can be made by doctors, patients or family members and/or acquaintances, or vaccine industry representatives. 

Read: Exclusive summary: Covid-19 vaccine concerns.

Some observers claim Covid-19 vaccine adverse events are not as likely to be underreported as those associated with other medicine, due to close monitoring and widespread publicity surrounding Covid-19 vaccination.

Approximately 340 million doses of Covid-19 vaccine have been given in the U.S. Slightly less than half of the U.S. population is fully vaccinated.

According to the Centers for Disease Control (CDC) and Food and Drug Administration (FDA), the benefits of Covid-19 vaccine outweigh the risks for all groups and age categories authorized to receive it.

Watch: CDC disinformation re: studies on Covid-19 vaccine effectiveness in people who have had Covid-19.

The following is a summary of some of the most frequent adverse events reported to VAERS after Covid-19 vaccination. (It is not the entire list.)

Most common Covid-19 vaccine adverse events reported as of July 19, 2021

Yellow highlighted adverse events are subjects of investigations, warnings or stated concerns by public health officials. For details, click here.

128,370 Muscle, bone, joint pain and swelling including:

  • 39,902 Pain in extremity
  • 37,819 Myalgia, muscle pain, weakness, fatigue, spasms, disorders, related
  • 30,138 Arthralgia, joint pain or arthritis, swelling, joint disease, bone pain, spinal osteoarthritis
  • 14,682 Back pain, neck pain
  • 5,829 Muscle and skeletal pain, stiffness, weakness

119,866 Injection site pain, bleeding, hardening, bruising, etc.

105,332 Skin reddening, at injection site or elsewhere, rash, hives

100,564 Fatigue, lethargy, malaise, asthenia, abnormal weakness, loss of energy

89,302 Headache, incl. migraine, sinus

68,252 Vomiting, nausea

68,064 Fever

63,133 Chills

60,913 Pain

49,574 Dizziness

34,076 Flushing, hot flush, feeling hot, abnormally warm skin

31,785 Lung pain or abnormalities, fluid in lung, respiratory tract or lung congestion or infection, wheezing, acute respiratory failure including:

  • 23,005 Dyspnoea, difficulty breathing
  • 1,398 Pneumonia
  • 1,128 Respiratory arrest, failure, stopped or inefficient breathing, abnormal breathing
  • 563 Covid-19 pneumonia
  • 265 Mechanical ventilation
  • 217 Bronchitis

30,909 Skin swelling, pain, tightness, face swelling, swelling under skin, hives, angioedema including:

  • 7,579 Skin pain, sensitivity, burning, discoloration, tenderness

25,319 Heart failure, heart rhythm and rate abnormalities, atrial fibrillation, palpitations, flutter, murmur, pacemaker added, fluid in heart, abnormal echocardiogram including:

  • 3,105 Heart attack or cardiac arrest, sudden loss of blood flow from failure to pump to heart effectively, cardiac failure, disorder

22,085 Itchiness

29,861 Sensory disturbance including:

  • 8,236 Tinnitus, hearing noise
  • 7,951 Abnormal vision, blindness
  • 6,349 Ageusia, loss of taste, altered taste, disorders
  • 2,249 Anosmia, loss of smell, parosmia (rotten smell)
  • 2,075 Hypersensitivity
  • 1,560 Sensitivity or reaction to light 
  • 890 Hearing loss, deafness

Characteristics of SARS-CoV-2 and COVID-19

Abstract

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) is a highly transmissible and pathogenic coronavirus that emerged in late 2019 and has caused a pandemic of acute respiratory disease, named ‘coronavirus disease 2019’ (COVID-19), which threatens human health and public safety. In this Review, we describe the basic virology of SARS-CoV-2, including genomic characteristics and receptor use, highlighting its key difference from previously known coronaviruses. We summarize current knowledge of clinical, epidemiological and pathological features of COVID-19, as well as recent progress in animal models and antiviral treatment approaches for SARS-CoV-2 infection. We also discuss the potential wildlife hosts and zoonotic origin of this emerging virus in detail.

Introduction

Coronaviruses are a diverse group of viruses infecting many different animals, and they can cause mild to severe respiratory infections in humans. In 2002 and 2012, respectively, two highly pathogenic coronaviruses with zoonotic origin, severe acute respiratory syndrome coronavirus (SARS-CoV) and Middle East respiratory syndrome coronavirus (MERS-CoV), emerged in humans and caused fatal respiratory illness, making emerging coronaviruses a new public health concern in the twenty-first century1. At the end of 2019, a novel coronavirus designated as SARS-CoV-2 emerged in the city of Wuhan, China, and caused an outbreak of unusual viral pneumonia. Being highly transmissible, this novel coronavirus disease, also known as coronavirus disease 2019 (COVID-19), has spread fast all over the world2,3. It has overwhelmingly surpassed SARS and MERS in terms of both the number of infected people and the spatial range of epidemic areas. The ongoing outbreak of COVID-19 has posed an extraordinary threat to global public health4,5. In this Review, we summarize the current understanding of the nature of SARS-CoV-2 and COVID-19. On the basis of recently published findings, this comprehensive Review covers the basic biology of SARS-CoV-2, including the genetic characteristics, the potential zoonotic origin and its receptor binding. Furthermore, we will discuss the clinical and epidemiological features, diagnosis of and countermeasures against COVID-19.

Emergence and spread

In late December 2019, several health facilities in Wuhan, in Hubei province in China, reported clusters of patients with pneumonia of unknown cause6. Similarly to patients with SARS and MERS, these patients showed symptoms of viral pneumonia, including fever, cough and chest discomfort, and in severe cases dyspnea and bilateral lung infiltration6,7. Among the first 27 documented hospitalized patients, most cases were epidemiologically linked to Huanan Seafood Wholesale Market, a wet market located in downtown Wuhan, which sells not only seafood but also live animals, including poultry and wildlife4,8. According to a retrospective study, the onset of the first known case dates back to 8 December 2019 (ref.9). On 31 December, Wuhan Municipal Health Commission notified the public of a pneumonia outbreak of unidentified cause and informed the World Health Organization (WHO)9 (Fig. 1).

figure1
Fig. 1: Timeline of the key events of the COVID-19 outbreak.

By metagenomic RNA sequencing and virus isolation from bronchoalveolar lavage fluid samples from patients with severe pneumonia, independent teams of Chinese scientists identified that the causative agent of this emerging disease is a betacoronavirus that had never been seen before6,10,11. On 9 January 2020, the result of this etiological identification was publicly announced (Fig. 1). The first genome sequence of the novel coronavirus was published on the Virological website on 10 January, and more nearly complete genome sequences determined by different research institutes were then released via the GISAID database on 12 January7. Later, more patients with no history of exposure to Huanan Seafood Wholesale Market were identified. Several familial clusters of infection were reported, and nosocomial infection also occurred in health-care facilities. All these cases provided clear evidence for human-to-human transmission of the new virus4,12,13,14. As the outbreak coincided with the approach of the lunar New Year, travel between cities before the festival facilitated virus transmission in China. This novel coronavirus pneumonia soon spread to other cities in Hubei province and to other parts of China. Within 1 month, it had spread massively to all 34 provinces of China. The number of confirmed cases suddenly increased, with thousands of new cases diagnosed daily during late January15. On 30 January, the WHO declared the novel coronavirus outbreak a public health emergency of international concern16. On 11 February, the International Committee on Taxonomy of Viruses named the novel coronavirus ‘SARS-CoV-2’, and the WHO named the disease ‘COVID-19’ (ref.17).

The outbreak of COVID-19 in China reached an epidemic peak in February. According to the National Health Commission of China, the total number of cases continued to rise sharply in early February at an average rate of more than 3,000 newly confirmed cases per day. To control COVID-19, China implemented unprecedentedly strict public health measures. The city of Wuhan was shut down on 23 January, and all travel and transportation connecting the city was blocked. In the following couple of weeks, all outdoor activities and gatherings were restricted, and public facilities were closed in most cities as well as in countryside18. Owing to these measures, the daily number of new cases in China started to decrease steadily19.

However, despite the declining trend in China, the international spread of COVID-19 accelerated from late February. Large clusters of infection have been reported from an increasing number of countries18. The high transmission efficiency of SARS-CoV-2 and the abundance of international travel enabled rapid worldwide spread of COVID-19. On 11 March 2020, the WHO officially characterized the global COVID-19 outbreak as a pandemic20. Since March, while COVID-19 in China has become effectively controlled, the case numbers in Europe, the USA and other regions have jumped sharply. According to the COVID-19 dashboard of the Center for System Science and Engineering at Johns Hopkins University, as of 11 August 2020, 216 countries and regions from all six continents had reported more than 20 million cases of COVID-19, and more than 733,000 patients had died21. High mortality occurred especially when health-care resources were overwhelmed. The USA is the country with the largest number of cases so far.

Although genetic evidence suggests that SARS-CoV-2 is a natural virus that likely originated in animals, there is no conclusion yet about when and where the virus first entered humans. As some of the first reported cases in Wuhan had no epidemiological link to the seafood market22, it has been suggested that the market may not be the initial source of human infection with SARS-CoV-2. One study from France detected SARS-CoV-2 by PCR in a stored sample from a patient who had pneumonia at the end of 2019, suggesting SARS-CoV-2 might have spread there much earlier than the generally known starting time of the outbreak in France23. However, this individual early report cannot give a solid answer to the origin of SARS-CoV-2 and contamination, and thus a false positive result cannot be excluded. To address this highly controversial issue, further retrospective investigations involving a larger number of banked samples from patients, animals and environments need to be conducted worldwide with well-validated assays.

For More Information: https://www.nature.com/articles/s41579-020-00459-7

The characteristics and evolution of pulmonary fibrosis in COVID-19 patients as assessed by AI-assisted chest HRCT

PLOS

Abstract

The characteristics and evolution of pulmonary fibrosis in patients with coronavirus disease 2019 (COVID-19) have not been adequately studied. AI-assisted chest high-resolution computed tomography (HRCT) was used to investigate the proportion of COVID-19 patients with pulmonary fibrosis, the relationship between the degree of fibrosis and the clinical classification of COVID-19, the characteristics of and risk factors for pulmonary fibrosis, and the evolution of pulmonary fibrosis after discharge. The incidence of pulmonary fibrosis in patients with severe or critical COVID-19 was significantly higher than that in patients with moderate COVID-19. There were significant differences in the degree of pulmonary inflammation and the extent of the affected area among patients with mild, moderate and severe pulmonary fibrosis. The IL-6 level in the acute stage and albumin level were independent risk factors for pulmonary fibrosis. Ground-glass opacities, linear opacities, interlobular septal thickening, reticulation, honeycombing, bronchiectasis and the extent of the affected area were significantly improved 30, 60 and 90 days after discharge compared with at discharge. The more severe the clinical classification of COVID-19, the more severe the residual pulmonary fibrosis was; however, in most patients, pulmonary fibrosis was improved or even resolved within 90 days after discharge.

Introduction

Pulmonary fibrosis can occur as a serious complication of viral pneumonia, which often leads to dyspnea and impaired lung function. It significantly affects quality of life and is associated with increased mortality in severe cases [12]. Patients with confirmed severe acute respiratory syndrome coronavirus (SARS‐CoV) or Middle East respiratory syndrome coronavirus (MERS‐CoV) infections were found to have different degrees of pulmonary fibrosis after hospital discharge, and some still had residual pulmonary fibrosis and impaired lung function two years later. In addition, wheezing and dyspnea have also been reported in critically ill patients [35].

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) is a novel Betacoronavirus that is responsible for an outbreak of acute respiratory illness known as coronavirus disease 2019 (COVID-19). SARS-CoV-2 shares 85% of its genome with the bat coronavirus bat-SL-CoVZC45 [6]. However, there are still some considerable differences between SARS-CoV-2 and SARS‐CoV or MERS‐CoV. Whether COVID-19 can trigger irreversible pulmonary fibrosis deserves more investigation. George reported that COVID-19 was associated with extensive respiratory deterioration, especially acute respiratory distress syndrome (ARDS), which suggested that there could be substantial fibrotic consequences of infection with SARS-CoV-2 [7]. Moreover, it has also been shown that the pathological manifestations of COVID-19 strongly resemble those of SARS and MERS [8], with pulmonary carnification and pulmonary fibrosis in the late stages.

Chest X-rays and high-resolution computed tomography (HRCT) of the chest play important auxiliary roles in the diagnosis and management of patients with suspected cases of COVID-19 [910]. The newly applied artificial intelligence (AI)-assisted pneumonia diagnosis system has been described as an objective tool that can be used to qualitatively and quantitatively assess the progression of pulmonary inflammation [11]. At present, although COVID-19 has been classified as a global epidemic for months, the risk factors for and severity and evolution of pulmonary fibrosis have not yet been reported. In this study, this new technology was applied to investigate the pulmonary imaging characteristics and related risk factors in COVID-19 patients at the time of hospital discharge, as well as the evolution of pulmonary fibrosis 30, 60 and 90 days after discharge, with the aim of providing an important basis for the clinical diagnosis, treatment and prognostic prediction of COVID-19-related pulmonary fibrosis.

For More Information: https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0248957

Post-acute COVID-19 syndrome

Abstract

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) is the pathogen responsible for the coronavirus disease 2019 (COVID-19) pandemic, which has resulted in global healthcare crises and strained health resources. As the population of patients recovering from COVID-19 grows, it is paramount to establish an understanding of the healthcare issues surrounding them. COVID-19 is now recognized as a multi-organ disease with a broad spectrum of manifestations. Similarly to post-acute viral syndromes described in survivors of other virulent coronavirus epidemics, there are increasing reports of persistent and prolonged effects after acute COVID-19. Patient advocacy groups, many members of which identify themselves as long haulers, have helped contribute to the recognition of post-acute COVID-19, a syndrome characterized by persistent symptoms and/or delayed or long-term complications beyond 4 weeks from the onset of symptoms. Here, we provide a comprehensive review of the current literature on post-acute COVID-19, its pathophysiology and its organ-specific sequelae. Finally, we discuss relevant considerations for the multidisciplinary care of COVID-19 survivors and propose a framework for the identification of those at high risk for post-acute COVID-19 and their coordinated management through dedicated COVID-19 clinics.

Main

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), the pathogen responsible for coronavirus disease 2019 (COVID-19), has caused morbidity and mortality at an unprecedented scale globally1. Scientific and clinical evidence is evolving on the subacute and long-term effects of COVID-19, which can affect multiple organ systems2. Early reports suggest residual effects of SARS-CoV-2 infection, such as fatigue, dyspnea, chest pain, cognitive disturbances, arthralgia and decline in quality of life3,4,5. Cellular damage, a robust innate immune response with inflammatory cytokine production, and a pro-coagulant state induced by SARS-CoV-2 infection may contribute to these sequelae6,7,8. Survivors of previous coronavirus infections, including the SARS epidemic of 2003 and the Middle East respiratory syndrome (MERS) outbreak of 2012, have demonstrated a similar constellation of persistent symptoms, reinforcing concern for clinically significant sequelae of COVID-19 (refs. 9,10,11,12,13,14,15).

Systematic study of sequelae after recovery from acute COVID-19 is needed to develop an evidence-based multidisciplinary team approach for caring for these patients, and to inform research priorities. A comprehensive understanding of patient care needs beyond the acute phase will help in the development of infrastructure for COVID-19 clinics that will be equipped to provide integrated multispecialty care in the outpatient setting. While the definition of the post-acute COVID-19 timeline is evolving, it has been suggested to include persistence of symptoms or development of sequelae beyond 3 or 4 weeks from the onset of acute symptoms of COVID-19 (refs. 16,17), as replication-competent SARS-CoV-2 has not been isolated after 3 weeks18. For the purpose of this review, we defined post-acute COVID-19 as persistent symptoms and/or delayed or long-term complications of SARS-CoV-2 infection beyond 4 weeks from the onset of symptoms (Fig. 1). Based on recent literature, it is further divided into two categories: (1) subacute or ongoing symptomatic COVID-19, which includes symptoms and abnormalities present from 4–12 weeks beyond acute COVID-19; and (2) chronic or post-COVID-19 syndrome, which includes symptoms and abnormalities persisting or present beyond 12 weeks of the onset of acute COVID-19 and not attributable to alternative diagnoses17,19. Herein, we summarize the epidemiology and organ-specific sequelae of post-acute COVID-19 and address management considerations for the interdisciplinary comprehensive care of these patients in COVID-19 clinics 

For More Information: https://www.nature.com/articles/s41591-021-01283-z

Taking a Closer Look at COVID-19’s Effects on the Brain

Authors: Dr. Francis Collins

While primarily a respiratory disease, COVID-19 can also lead to neurological problems. The first of these symptoms might be the loss of smell and taste, while some people also may later battle headaches, debilitating fatigue, and trouble thinking clearly, sometimes referred to as “brain fog.” All of these symptoms have researchers wondering how exactly the coronavirus that causes COVID-19, SARS-CoV-2, affects the human brain.

In search of clues, researchers at NIH’s National Institute of Neurological Disorders and Stroke (NINDS) have now conducted the first in-depth examinations of human brain tissue samples from people who died after contracting COVID-19. Their findings, published in the New England Journal of Medicine, suggest that COVID-19’s many neurological symptoms are likely explained by the body’s widespread inflammatory response to infection and associated blood vessel injury—not by infection of the brain tissue itself [1].

The NIH team, led by Avindra Nath, used a high-powered magnetic resonance imaging (MRI) scanner (up to 10 times as sensitive as a typical MRI) to examine postmortem brain tissue from 19 patients. They ranged in age from 5 to 73, and some had preexisting conditions, such as diabetes, obesity, and cardiovascular disease.
The team focused on the brain’s olfactory bulb that controls our ability to smell and the brainstem, which regulates breathing and heart rate. Based on earlier evidence, both areas are thought to be highly susceptible to COVID-19.

For More Information: https://directorsblog.nih.gov/2021/01/14/taking-a-closer-look-at-the-effects-of-covid-19-on-the-brain/