Prevalence of organ impairment in Long COVID patients 6 and 12 months after initial symptoms

Authors:  Pooja Toshniwal PahariaMar 24 2022Reviewed by Danielle Ellis, B.Sc.

In a recent study posted to the medRxiv* preprint server, researchers assessed the prevalence of organ impairment in long coronavirus disease 2019 (COVID-19) six months and a year post-COVID-19 at London and Oxford.

Multi-organ impairment associated with long COVID-19 is a significant health burden. Standardized multi-organ evaluation is deficient, especially in non-hospitalized patients. Although the symptoms of long COVID-19, also known as post-acute sequelae of COVID-19 (PASC), are well-established, the natural history is poorly classified by symptoms, organ impairment, and function.

About the study

In the present prospective study, researchers assessed organ impairment in long COVID-19 patients six months and a year after the onset of early symptoms and correlated them to their clinical presentation.

The participants were recruited based on specialist referral or the response to advertisements in sites such as Mayo Clinic Healthcare, Perspectum, and Oxford from April 2020 to August 2021, based on their COVID-19 history.

The study was conducted on COVID-19 patients who recovered from the acute phase of the infection. Their health status, symptoms, and organ impairment were assessed. The symptoms assessed comprised cardiopulmonary, severe dyspnoea, and cognitive dysfunction. Biochemical and physiological parameters were analyzed at baseline and post-organ impairment. The radiological investigation comprised multi-organ magnetic resonance imaging (MRI) performed in the long COVID-19 patients and healthy controls.

Over a year, the team prospectively investigated the symptoms, organ impairment, and function, especially dyspnea, cognitive dysfunction, and health-related quality of life (HRQoL). They also evaluated the association between organ impairment and clinical symptoms.

Patients with symptoms of active pulmonary infections (body temperature >37.8°C or ≥3 coughing episodes in a day) and hospital discharges in the previous week or >4 months were excluded from the study. Asymptomatic patients and those with MRI contraindications such as defibrillators, pacemakers, devices with metal implants, and claustrophobia were removed.

Participants with impaired organs, as diagnosed by blood investigations, incidental findings, or MRI, were included in the follow-up assessments. Every visit comprised blood investigations, MRI scanning, and online questionnaire surveys, which were to be filled out beforehand. In addition, a sensitivity analysis was performed that excluded patients at risk of metabolic disorders (including body mass index (BMI) ≥30 kg/m2, diabetes, and hypertension)

Results

Related Stories

Out of 536 participants, the majority were middle-aged (mean age 45 years), female (73%), White (89%), and healthcare workers (32%). About 13% of the COVID-19 patients hospitalized during the acute phase of the infection completed the baseline evaluation. A total of 331 patients (62%) had incidental findings, organ impairment, or reduction in the symptoms from the baseline at both the time points.

Cognitive dysfunction (50% and 38%), poor HRQoL (EuroQOL <0.7 in 55% and 45%), and severe dyspnea (36% and 30%) were observed at six months and one year, respectively. On follow-up, the symptoms were reduced, especially cardiopulmonary and systemic symptoms, whereas fatigue, dyspnea, and cognitive dysfunction were consistently present. The greatest impact on quality of life was related to pain and difficulties performing routine activities. Almost every patient took time off work due to COVID-19. The symptoms were largely associated with obese women, young age, and impairment of a single organ.

At baseline, fibrous inflammation was observed in the pancreas (9%), heart (9%), liver (11%), and kidney (15%). Additionally, increased volumes of the spleen (8%), kidney (9%), and liver (7%) were observed. Moreover, reduced lung capacity (2%), excess adipose deposits in pancreatic tissues (15%) and liver (25%) were observed. High liver fibro-inflammation was associated with cognitive dysfunction at follow-up in 19% and 12% of patients with and without cognitive dysfunction, respectively. Low liver fat was more likely in those without severe dyspnoea at both time points. Increased liver volumes at follow-up were associated with lower HRQoL scores.

The prevalence of multi and single-organ impairment was 23% and 59% at baseline, respectively, and persisted in 27% and 59% of the participants on follow-up assessments. Most of the organ impairments were mild. However, they did not improve substantially between visits. Notably, participants without organ impairment had the lowest symptom burden.

Most biochemical parameters were normal except creatinine kinase (8% and 13%), lactate dehydrogenase (16% and 22%), mean cell hemoglobin concentration (21% and 15%), and cholesterol (46% and 48%), at six months and a year post-COVID-19, respectively. These biochemical markers increased from the baseline on follow-up assessments.

Conclusion

To summarize, organ impairment was detected in 59% of the patients at six months post-COVID-19 and persisted in 59% at one-year follow-up. This has significant implications on the quality of life, symptoms, and long-term health of the patients. These observations highlight the requirement for enhanced preventive measures and integrated patient care to decrease the long COVID-19 burden.

Journal reference:

Long COVID Could Be Linked to the Effects of SARS-CoV-2 on the Vagus Nerve

SciTechDaily

Authors: EUROPEAN SOCIETY OF CLINICAL MICROBIOLOGY AND INFECTIOUS DISEASES FEBRUARY 11, 2022

New research to be presented at this year’s European Congress of Clinical Microbiology and Infectious Diseases (ECCMID 2022, Lisbon, April 23-26) suggests that many of the symptoms connected to post-COVID syndrome (PCC, also known as long COVID) could be linked to the effect of the virus on the vagus nerve – one of the most important multi-functional nerves in the body. The study is by Dr. Gemma Lladós and Dr. Lourdes Mateu, University Hospital Germans Trias i Pujol, Badalona, Spain, and colleagues.

The vagus nerve extends from the brain down into the torso and into the heart, lungs, and intestines, as well as several muscles including those involved in swallowing. As such, this nerve is responsible for a wide variety of bodily functions including controlling heart rate, speech, the gag reflex, transferring food from the mouth to the stomach, moving food through the intestines, sweating, and many others.

Long COVID is a potentially disabling syndrome affecting an estimated 10-15% of subjects who survive COVID-19. The authors propose that SARS-CoV-2-mediated vagus nerve dysfunction (VND) could explain some long COVID symptoms, including dysphonia (persistent voice problems), dysphagia (difficulty in swallowing), dizziness, tachycardia (abnormally high heart rate), orthostatic hypotension (low blood pressure) and diarrhea.

The authors performed a pilot, extensive morphological and functional evaluation of the vagus nerve, using imaging and functional tests in a prospective observational cohort of long COVID subjects with symptoms suggestive of VND. In their total cohort of 348 patients, 228 (66%) had at least one symptom suggestive of VND. The current evaluation was performed in the first 22 subjects with VND symptoms (10% of the total) seen in the Long COVID Clinic of University Hospital Germans Trias i Pujol between March and June 2021. The study is ongoing and continues to recruit patients.

Of the 22 subjects analyzed, 20 (91%) were women with a median age of 44 years. The most frequent VND-related symptoms were: diarrhea (73%), tachycardia (59%), dizziness, dysphagia and dysphonia (45% each), and orthostatic hypotension (14%). Almost all (19 subjects, 86%) had at least 3 VND-related symptoms. The median prior duration of symptoms was 14 months. Six of 22 patients (27%) displayed alteration of the vagus nerve in the neck shown by ultrasound – including both thickening of the nerve and increased ‘echogenicity’ which indicates mild inflammatory reactive changes.

A thoracic ultrasound showed flattened ‘diaphragmatic curves’ in 10 out of 22 (46%) subjects (which translates a decrease in diaphragmatic mobility during breathing, or more simply abnormal breathing). A total of 10 of 16 (63%) assessed individuals showed reduced maximum inspiration pressures, showing weakness of breathing muscles.

Eating and digestive function was also affected in some patients, with 13 of 18 assessed (72%) having a positive screen for self-perceived oropharyngeal dysphagia (trouble swallowing). An assessment of gastric and bowel function performed in 19 patients revealed 8 (42%) had their ability to deliver food to the stomach (via the esophagus) impaired, with 2 of these 8 (25%) reporting difficulty in swallowing. Gastroesophageal reflux (acid reflux) was observed in 9 of 19 (47%) individuals; with 4 of these 9 (44%) again having difficulty delivering food to the stomach and 3 of these 9 (33%) with hiatal hernia – which occurs when the upper part of the stomach bulges through the diaphragm into the chest cavity.

A Voice Handicap Index 30 test (a standard way to measure voice function) was abnormal in 8/17 (47%) cases, with 7 of these 8 cases (88%) suffering dysphonia.

The authors say: “In this pilot evaluation, most long COVID subjects with vagus nerve dysfunction symptoms had a range of significant, clinically-relevant, structural and/or functional alterations in their vagus nerve, including nerve thickening, trouble swallowing, and symptoms of impaired breathing. Our findings so far thus point at vagus nerve dysfunction as a central pathophysiological feature of long COVID.”

Meeting: The European Congress of Clinical Microbiology & Infectious Diseases (ECCMID 2022)

1000 Peer Reviewed Studies Questioning Covid-19 Vaccine Safety

Peer Reviewed Medical Papers Submitted To Various Medical Journals, Evidencing A Multitude Of Adverse Events In Covid-19 Vaccine Recipients.

The list includes studies published as of January 20, 2022 concerning the potential adverse reaction from COVID-19 vaccines, such as myocarditis, thrombosis, thrombocytopenia, vasculitis, cardiac, Bell’s Palsy, immune-mediated disease, and many more.

  1. Myocarditis after mRNA vaccination against SARS-CoV-2, a case series: https://www.sciencedirect.com/science/article/pii/S2666602221000409
  2. Myocarditis after immunization with COVID-19 mRNA vaccines in members of the US military. This article reports that in “23 male patients, including 22 previously healthy military members, myocarditis was identified within 4 days after receipt of the vaccine”: https://jamanetwork.com/journals/jamacardiology/fullarticle/2781601
  3. Association of myocarditis with the BNT162b2 messenger RNA COVID-19 vaccine in a case series of children: https://pubmed.ncbi.nlm.nih.gov/34374740/
  4. Acute symptomatic myocarditis in seven adolescents after Pfizer-BioNTech COVID-19 vaccination: https://pediatrics.aappublications.org/content/early/2021/06/04/peds.2021-052478
  5. Myocarditis and pericarditis after vaccination with COVID-19 mRNA: practical considerations for care providers: https://www.sciencedirect.com/science/article/pii/S0828282X21006243
  6. Myocarditis, pericarditis and cardiomyopathy after COVID-19 vaccination: https://www.sciencedirect.com/science/article/pii/S1443950621011562
  7. Myocarditis with COVID-19 mRNA vaccines: https://www.ahajournals.org/doi/pdf/10.1161/CIRCULATIONAHA.121.056135
  8. Myocarditis and pericarditis after COVID-19 vaccination: https://jamanetwork.com/journals/jama/fullarticle/2782900
  9. Myocarditis temporally associated with COVID-19 vaccination: https://www.ahajournals.org/doi/pdf/10.1161/CIRCULATIONAHA.121.055891.
  10. COVID-19 Vaccination Associated with Myocarditis in Adolescents: https://pediatrics.aappublications.org/content/pediatrics/early/2021/08/12/peds.2021-053427.full.pdf
  11. Acute myocarditis after administration of BNT162b2 vaccine against COVID-19: https://pubmed.ncbi.nlm.nih.gov/33994339/
  12. Temporal association between COVID-19 vaccine Ad26.COV2.S and acute myocarditis: case report and review of the literature: https://www.sciencedirect.com/science/article/pii/S1553838921005789
  13. COVID-19 vaccine-induced myocarditis: a case report with review of the literature: https://www.sciencedirect.com/science/article/pii/S1871402121002253
  14. Potential association between COVID-19 vaccine and myocarditis: clinical and CMR findings: https://www.sciencedirect.com/science/article/pii/S1936878X2100485X
  15. Recurrence of acute myocarditis temporally associated with receipt of coronavirus mRNA disease vaccine 2019 (COVID-19) in a male adolescent: https://www.sciencedirect.com/science/article/pii/S002234762100617X
  16. Fulminant myocarditis and systemic hyper inflammation temporally associated with BNT162b2 COVID-19 mRNA vaccination in two patients: https://www.sciencedirect.com/science/article/pii/S0167527321012286.
  17. Acute myocarditis after administration of BNT162b2 vaccine: https://www.sciencedirect.com/science/article/pii/S2214250921001530
  18. Lymphohistocytic myocarditis after vaccination with COVID-19 Ad26.COV2.S viral vector: https://www.sciencedirect.com/science/article/pii/S2352906721001573
  19. Myocarditis following vaccination with BNT162b2 in a healthy male: https://www.sciencedirect.com/science/article/pii/S0735675721005362
  20. Acute myocarditis after Comirnaty (Pfizer) vaccination in a healthy male with previous SARS-CoV-2 infection: https://www.sciencedirect.com/science/article/pii/S1930043321005549
  21. Acute myocarditis after vaccination with SARS-CoV-2 mRNA-1273 mRNA: https://www.sciencedirect.com/science/article/pii/S2589790X21001931
  22. Acute myocarditis after SARS-CoV-2 vaccination in a 24-year-old man: https://www.sciencedirect.com/science/article/pii/S0870255121003243
  23. A series of patients with myocarditis after vaccination against SARS-CoV-2 with mRNA-1279 and BNT162b2: https://www.sciencedirect.com/science/article/pii/S1936878X21004861
  24. COVID-19 mRNA vaccination and myocarditis: https://pubmed.ncbi.nlm.nih.gov/34268277/
  25. COVID-19 vaccine and myocarditis: https://pubmed.ncbi.nlm.nih.gov/34399967/
  26. Epidemiology and clinical features of myocarditis/pericarditis before the introduction of COVID-19 mRNA vaccine in Korean children: a multicenter study https://search.bvsalud.org/global-literature-on-novel-coronavirus-2019-ncov/resourc e/en/covidwho-1360706.
  27. COVID-19 vaccines and myocarditis: https://pubmed.ncbi.nlm.nih.gov/34246566/
  28. Myocarditis and other cardiovascular complications of COVID-19 mRNA-based COVID-19 vaccines https://www.cureus.com/articles/61030-myocarditis-and-other-cardiovascular-complications-of-the-mrna-based-covid-19-vaccines
  29. Myocarditis and other cardiovascular complications of COVID-19 mRNA-based COVID-19 vaccines https://www.cureus.com/articles/61030-myocarditis-and-other-cardiovascular-complications-of-the-mrna-based-covid-19-vaccines
  30. Myocarditis, pericarditis, and cardiomyopathy after COVID-19 vaccination: https://pubmed.ncbi.nlm.nih.gov/34340927/
  31. Myocarditis with covid-19 mRNA vaccines: https://www.ahajournals.org/doi/10.1161/CIRCULATIONAHA.121.056135
  32. Association of myocarditis with COVID-19 mRNA vaccine in children: https://media.jamanetwork.com/news-item/association-of-myocarditis-with-mrna-co vid-19-vaccine-in-children/
  33. Association of myocarditis with COVID-19 messenger RNA vaccine BNT162b2 in a case series of children: https://jamanetwork.com/journals/jamacardiology/fullarticle/2783052
  34. Myocarditis after immunization with COVID-19 mRNA vaccines in members of the U.S. military: https://jamanetwork.com/journals/jamacardiology/fullarticle/2781601%5C
  35. Myocarditis occurring after immunization with COVID-19 mRNA-based COVID-19 vaccines: https://jamanetwork.com/journals/jamacardiology/fullarticle/2781600
  36. Myocarditis following immunization with Covid-19 mRNA: https://www.nejm.org/doi/full/10.1056/NEJMc2109975
  37. Patients with acute myocarditis after vaccination withCOVID-19 mRNA: https://jamanetwork.com/journals/jamacardiology/fullarticle/2781602
  38. Myocarditis associated with vaccination with COVID-19 mRNA: https://pubs.rsna.org/doi/10.1148/radiol.2021211430
  39. Symptomatic Acute Myocarditis in 7 Adolescents after Pfizer-BioNTech COVID-19 Vaccination: https://pediatrics.aappublications.org/content/148/3/e2021052478
  40. Cardiovascular magnetic resonance imaging findings in young adult patients with acute myocarditis after COVID-19 mRNA vaccination: a case series: https://jcmr-online.biomedcentral.com/articles/10.1186/s12968-021-00795-4
  41. Clinical Guidance for Young People with Myocarditis and Pericarditis after Vaccination with COVID-19 mRNA: https://www.cps.ca/en/documents/position/clinical-guidance-for-youth-with-myocarditis-and-pericarditis
  42. Cardiac imaging of acute myocarditis after vaccination with COVID-19 mRNA: https://pubmed.ncbi.nlm.nih.gov/34402228/
  43. Case report: acute myocarditis after second dose of mRNA-1273 SARS-CoV-2 mRNA vaccine: https://academic.oup.com/ehjcr/article/5/8/ytab319/6339567
  44. Myocarditis / pericarditis associated with COVID-19 vaccine: https://science.gc.ca/eic/site/063.nsf/eng/h_98291.html
  45. The new COVID-19 mRNA vaccine platform and myocarditis: clues to the possible underlying mechanism: https://pubmed.ncbi.nlm.nih.gov/34312010/
  46. Myocarditis associated with COVID-19 vaccination: echocardiographic, cardiac tomography, and magnetic resonance imaging findings: https://www.ahajournals.org/doi/10.1161/CIRCIMAGING.121.013236
  47. In-depth evaluation of a case of presumed myocarditis after the second dose of COVID-19 mRNA vaccine: https://www.ahajournals.org/doi/10.1161/CIRCULATIONAHA.121.056038
  48. Occurrence of acute infarct-like myocarditis after COVID-19 vaccination: just an accidental coincidence or rather a vaccination-associated autoimmune myocarditis?: https://pubmed.ncbi.nlm.nih.gov/34333695/

This list is not meant to be all inclusive of all peer-reviewed potential harms from mRNA vaccines. To access any of the 1,000 Vaccine Harms published in Medical journals Click The Link Below:

https://www.informedchoiceaustralia.com/post/1000-peer-reviewed-studies-questioning-covid-19-vaccine-safety

OR VIEW AND DOWNLOAD FULL LIST PDF

Updated_Peer_Reviewed_medical_papers_submitted_to_various_medical.pdfDownload PDF • 1.01MB

Pulmonary Vascular Endothelialitis, Thrombosis, and Angiogenesis in Covid-19

Authors.Maximilian Ackermann, M.D., Stijn E. Verleden, Ph.D., Mark Kuehnel, Ph.D., Axel Haverich, M.D., Tobias Welte, M.D., Florian Laenger, M.D., Arno Vanstapel, Ph.D., Christopher Werlein, M.D., Helge Stark, Ph.D., Alexandar Tzankov, M.D., William W. Li, M.D., Vincent W. Li, M.D., et al.

July 9, 2020 N Engl J Med 2020; 383:120-128 DOI: 10.1056/NEJMoa2015432

Abstract

BACKGROUND

Progressive respiratory failure is the primary cause of death in the coronavirus disease 2019 (Covid-19) pandemic. Despite widespread interest in the pathophysiology of the disease, relatively little is known about the associated morphologic and molecular changes in the peripheral lung of patients who die from Covid-19.

METHODS

We examined 7 lungs obtained during autopsy from patients who died from Covid-19 and compared them with 7 lungs obtained during autopsy from patients who died from acute respiratory distress syndrome (ARDS) secondary to influenza A(H1N1) infection and 10 age-matched, uninfected control lungs. The lungs were studied with the use of seven-color immunohistochemical analysis, micro–computed tomographic imaging, scanning electron microscopy, corrosion casting, and direct multiplexed measurement of gene expression.

RESULTS

In patients who died from Covid-19–associated or influenza-associated respiratory failure, the histologic pattern in the peripheral lung was diffuse alveolar damage with perivascular T-cell infiltration. The lungs from patients with Covid-19 also showed distinctive vascular features, consisting of severe endothelial injury associated with the presence of intracellular virus and disrupted cell membranes. Histologic analysis of pulmonary vessels in patients with Covid-19 showed widespread thrombosis with microangiopathy. Alveolar capillary microthrombi were 9 times as prevalent in patients with Covid-19 as in patients with influenza (P<0.001). In lungs from patients with Covid-19, the amount of new vessel growth — predominantly through a mechanism of intussusceptive angiogenesis — was 2.7 times as high as that in the lungs from patients with influenza (P<0.001).

CONCLUSIONS

In our small series, vascular angiogenesis distinguished the pulmonary pathobiology of Covid-19 from that of equally severe influenza virus infection. The universality and clinical implications of our observations require further research to define. (Funded by the National Institutes of Health and others.)

Infection with severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) in humans is associated with a broad spectrum of clinical respiratory syndromes, ranging from mild upper airway symptoms to progressive life-threatening viral pneumonia.1,2 Clinically, patients with severe coronavirus disease 2019 (Covid-19) have labored breathing and progressive hypoxemia and often receive mechanical ventilatory support. Radiographically, peripheral lung ground-glass opacities on computed tomographic (CT) imaging of the chest fulfill the Berlin criteria for acute respiratory distress syndrome (ARDS).3,4 Histologically, the hallmark of the early phase of ARDS is diffuse alveolar damage with edema, hemorrhage, and intraalveolar fibrin deposition, as described by Katzenstein et al.5 Diffuse alveolar damage is a nonspecific finding, since it may have noninfectious or infectious causes, including Middle East respiratory syndrome coronavirus (MERS-CoV),6 SARS-CoV,7 SARS-CoV-2,8-10 and influenza viruses.11

Among the distinctive features of Covid-19 are the vascular changes associated with the disease. With respect to diffuse alveolar damage in SARS-CoV7 and SARS-CoV-2 infection,8,12 the formation of fibrin thrombi has been observed anecdotally but not studied systematically. Clinically, many patients have elevated d-dimer levels, as well as cutaneous changes in their extremities suggesting thrombotic microangiopathy.13 Diffuse intravascular coagulation and large-vessel thrombosis have been linked to multisystem organ failure.14-16 Peripheral pulmonary vascular changes are less well characterized; however, vasculopathy in the gas-exchange networks, depending on its effect on the matching of ventilation and perfusion that results, could potentially contribute to hypoxemia and the effects of posture (e.g., prone positioning) on oxygenation.17

Despite previous experience with SARS-CoV18 and early experience with SARS-CoV-2, the morphologic and molecular changes associated with these infections in the peripheral lung are not well documented. Here, we examine the morphologic and molecular features of lungs obtained during autopsy from patients who died from Covid-19, as compared with those of lungs from patients who died from influenza and age-matched, uninfected control lungs.

Methods

PATIENT SELECTION AND WORKFLOW

We analyzed pulmonary autopsy specimens from seven patients who died from respiratory failure caused by SARS-CoV-2 infection and compared them with lungs from seven patients who died from pneumonia caused by influenza A virus subtype H1N1 (A[H1N1]) — a strain associated with the 1918 and 2009 influenza pandemics. The lungs from patients with influenza were archived tissue from the 2009 pandemic and were chosen for the best possible match with respect to age, sex, and disease severity from among the autopsies performed at the Hannover Medical School. Ten lungs that had been donated but not used for transplantation served as uninfected control specimens. The Covid-19 group consisted of lungs from two female and five male patients with mean (±SD) ages of 68±9.2 years and 80±11.5 years, respectively (clinical data are provided in Table S1A in the Supplementary Appendix, available with the full text of this article at NEJM.org). The influenza group consisted of lungs from two female and five male patients with mean ages of 62.5±4.9 years and 55.4±10.9 years, respectively. Five of the uninfected lungs were from female donors (mean age, 68.2±6.9 years), and five were from male donors (mean age, 79.2±3.3 years) (clinical data are provided in Table S1B). The study was approved by and conducted according to requirements of the ethics committees at the Hannover Medical School and the University of Leuven. There was no commercial support for this study.

All lungs were comprehensively analyzed with the use of microCT, histopathological, and multiplexed immunohistochemical analysis, transmission and scanning electron microscopy, corrosion casting, and direct multiplexed gene-expression analysis, as described in detail in the Methods section of the Supplementary Appendix.

STATISTICAL ANALYSIS

All comparisons of numeric variables (including those in the gene-expression analysis) were conducted with Student’s t-test familywise error rates due to multiplicity set at 0.05 with the use of the Benjamini–Hochberg method of controlling false discovery rates. Original P values are reported only for the tests that met the criteria for false discovery rates. All confidence intervals have been calculated on the basis of the t-distribution, as well. Additional details are provided in the Methods section of the Supplementary Appendix.

Results

GROSS EXAMINATION

The mean (±SE) weight of the lungs from patients with proven influenza pneumonia was significantly higher than that from patients with proven Covid-19 (2404±560 g vs. 1681±49 g; P=0.04). The mean weight of the uninfected control lungs (1045±91 g) was significantly lower than those in the influenza group (P=0.003) and the Covid-19 group (P<0.001).

ANGIOCENTRIC INFLAMMATION

Figure 1. Lymphocytic Inflammation in a Lung from a Patient Who Died from Covid-19.

All lung specimens from the Covid-19 group had diffuse alveolar damage with necrosis of alveolar lining cells, pneumocyte type 2 hyperplasia, and linear intraalveolar fibrin deposition (Figure 1). In four of seven cases, the changes were focal, with only mild interstitial edema. The remaining three cases had homogeneous fibrin deposits and marked interstitial edema with early intraalveolar organization. The specimens in the influenza group had florid diffuse alveolar damage with massive interstitial edema and extensive fibrin deposition in all cases. In addition, three specimens in the influenza group had focal organizing and resorptive inflammation (Fig. S2). These changes were reflected in the much higher weight of the lungs from patients with influenza.

Immunohistochemical analysis of angiotensin-converting enzyme 2 (ACE2) expression, measured as mean (±SD) relative counts of ACE2-positive cells per field of view, in uninfected control lungs showed scarce expression of ACE2 in alveolar epithelial cells (0.053±0.03) and capillary endothelial cells (0.066±0.03). In lungs from patients with Covid-19 and lungs from patients with influenza, the relative counts of ACE2-positive cells per field of view were 0.25±0.14 and 0.35±0.15, respectively, for alveolar epithelial cells and 0.49±0.28 and 0.55±0.11, respectively, for endothelial cells. Furthermore, ACE2-positive lymphocytes were not seen in perivascular tissue or in the alveoli of the control lungs but were present in the lungs in the Covid-19 group and the influenza group (relative counts of 0.22±0.18 and 0.15±0.09, respectively). (Details of counting are provided in Table S2.)

In the lungs from patients with Covid-19 and patients with influenza, similar mean (±SD) numbers of CD3-positive T cells were found within a 200-μm radius of precapillary and postcapillary vessel walls in 20 fields of examination per patient (26.2±13.1 for Covid-19 and 14.8±10.8 for influenza). With the same field size used for examination, CD4-positive T cells were more numerous in lungs from patients with Covid-19 than in lungs from patients with influenza (13.6±6.0 vs. 5.8±2.5, P=0.04), whereas CD8-positive T cells were less numerous (5.3±4.3 vs. 11.6±4.9, P=0.008). Neutrophils (CD15 positive) were significantly less numerous adjacent to the alveolar epithelial lining in the Covid-19 group than in the influenza group (0.4±0.5 vs. 4.8±5.2, P=0.002).

A multiplexed analysis of inflammation-related gene expression examining 249 genes from the nCounter Inflammation Panel (NanoString Technologies) revealed similarities and differences between the specimens in the Covid-19 group and those in the influenza group. A total of 79 inflammation-related genes were differentially regulated only in specimens from patients with Covid-19, whereas 2 genes were differentially regulated only in specimens from patients with influenza; a shared expression pattern was found for 7 genes (Fig. S1).

THROMBOSIS AND MICROANGIOPATHY

Figure 2. Microthrombi in the Interalveolar Septa of a Lung from a Patient Who Died from Covid-19.

The pulmonary vasculature of the lungs in the Covid-19 group and the influenza group was analyzed with hematoxylin–eosin, trichrome, and immunohistochemical staining (as described in the Methods section of the Supplementary Appendix). Analysis of precapillary vessels showed that in four of the seven lungs from patients with Covid-19 and four of the seven lungs from the patients with influenza, thrombi were consistently present in pulmonary arteries with a diameter of 1 mm to 2 mm, without complete luminal obstruction (Figs. S3 and S5). Fibrin thrombi of the alveolar capillaries could be seen in all the lungs from both groups of patients (Figure 2). Alveolar capillary microthrombi were 9 times as prevalent in patients with Covid-19 as in patients with influenza (mean [±SD] number of distinct thrombi per square centimeter of vascular lumen area, 159±73 and 16±16, respectively; P=0.002). Intravascular thrombi in postcapillary venules of less than 1 mm diameter were seen in lower numbers in the lungs from patients with Covid-19 than in those from patients with influenza (12±14 vs. 35±16, P=0.02). Two lungs in the Covid-19 group had involvement of all segments of the vasculature, as compared with four of the lungs in the influenza group; in three of the lungs in the Covid-19 group and three of the lungs in the influenza group, combined capillary and venous thrombi were found without arterial thrombi.

The histologic findings were supported by three-dimensional microCT of the pulmonary specimens: the lungs from patients with Covid-19 and from patients with influenza showed nearly total occlusions of precapillary and postcapillary vessels.

ANGIOGENESIS

Figure 3. Microvascular Alterations in Lungs from Patients Who Died from Covid-19

.

We examined the microvascular architecture of the lungs from patients with Covid-19, lungs from patients with influenza, and uninfected control lungs with the use of scanning electron microscopy and microvascular corrosion casting. The lungs in the Covid-19 group had a distorted vascularity with structurally deformed capillaries (Figure 3). Elongated capillaries in the lungs from patients with Covid-19 showed sudden changes in caliber and the presence of intussusceptive pillars within the capillaries (Figure 3C). Transmission electron microscopy of the Covid-19 endothelium showed ultrastructural damage to the endothelium, as well as the presence of intracellular SARS-CoV-2 (Figure 3D). The virus could also be identified in the extracellular space.

Figure 4.Numeric Density of Features of Intussusceptive and Sprouting Angiogenesis in Lungs from Patients Who Died from Covid-19 or Influenza A(H1N1).

In the lungs from patients with Covid-19, the density of intussusceptive angiogenic features (mean [±SE], 60.7±11.8 features per field) was significantly higher than that in lungs from patients with influenza (22.5±6.9) or in uninfected control lungs (2.1±0.6) (P<0.001 for both comparisons) (Figure 4A). The density of features of conventional sprouting angiogenesis was also higher in the Covid-19 group than in the influenza group (Figure 4B). When the pulmonary angiogenic feature count was plotted as a function of the length of hospital stay, the degree of intussusceptive angiogenesis was found to increase significantly with increasing duration of hospitalization (P<0.001) (Figure 4C). In contrast, the lungs from patients with influenza had less intussusceptive angiogenesis and no increase over time (Figure 4C). A similar pattern was seen for sprouting angiogenesis (Figure 4D).Figure 5.elative Expression Analysis of Angiogenesis-Associated Genes in Lungs from Patients Who Died from Covid-19 or Influenza A(H1N1).

A multiplexed analysis of angiogenesis-related gene expression examining 323 genes from the nCounter PanCancer Progression Panel (NanoString Technologies) revealed differences between the specimens from patients with Covid-19 and those from patients with influenza. A total of 69 angiogenesis-related genes were differentially regulated only in the Covid-19 group, as compared with 26 genes differentially regulated only in the influenza group; 45 genes had shared changes in expression (Figure 5).

Discussion

In this study, we examined the morphologic and molecular features of seven lungs obtained during autopsy from patients who died from SARS-CoV-2 infection. The lungs from these patients were compared with those obtained during autopsy from patients who had died from ARDS secondary to influenza A(H1N1) infection and from uninfected controls. The lungs from the patients with Covid-19 and the patients with influenza shared a common morphologic pattern of diffuse alveolar damage and infiltrating perivascular lymphocytes. There were three distinctive angiocentric features of Covid-19. The first feature was severe endothelial injury associated with intracellular SARS-CoV-2 virus and disrupted endothelial cell membranes. Second, the lungs from patients with Covid-19 had widespread vascular thrombosis with microangiopathy and occlusion of alveolar capillaries.12,19 Third, the lungs from patients with Covid-19 had significant new vessel growth through a mechanism of intussusceptive angiogenesis. Although our sample was small, the vascular features we identified are consistent with the presence of distinctive pulmonary vascular pathobiologic features in some cases of Covid-19.

Our finding of enhanced intussusceptive angiogenesis in the lungs from patients with Covid-19 as compared with the lungs from patients with influenza was unexpected. New vessel growth can occur by conventional sprouting or intussusceptive (nonsprouting) angiogenesis. The characteristic feature of intussusceptive angiogenesis is the presence of a pillar or post spanning the lumen of the vessel.20 Typically referred to as an intussusceptive pillar, this endothelial-lined intravascular structure is not seen by light microscopy but is readily identifiable by corrosion casting and scanning electron microscopy.21 Although tissue hypoxia was probably a common feature in the lungs from both these groups of patients, we speculate that the greater degree of endothelialitis and thrombosis in the lungs from patients with Covid-19 may contribute to the relative frequency of sprouting and intussusceptive angiogenesis observed in these patients. The relationship of these findings to the clinical course of Covid-19 requires further research to elucidate.

A major limitation of our study is that the sample was small; we studied only 7 patients among the more than 320,000 people who have died from Covid-19, and the autopsy data also represent static information. On the basis of the available data, we cannot reconstruct the timing of death in the context of an evolving disease process. Moreover, there could be other factors that account for the differences we observed between patients with Covid-19 and those with influenza. For example, none of the patients in our study who died from Covid-19 had been treated with standard mechanical ventilation, whereas five of the seven patients who died from influenza had received pressure-controlled ventilation. Similarly, it is possible that differences in detectable intussusceptive angiogenesis could be due to the different time courses of Covid-19 and influenza. These and other unknown factors must be considered when evaluating our data.22 Nonetheless, our analysis suggests that this possibility is unlikely, particularly since the degree of intussusceptive angiogenesis in the patients with Covid-19 increased significantly with increasing length of hospitalization, whereas in the patients with influenza it remained stable at a significantly lower level. Moreover, we have shown intussusceptive angiogenesis to be the predominant angiogenic mechanism even in late stages of chronic lung injury.21

ACE2 is an integral membrane protein that appears to be the host-cell receptor for SARS-CoV-2.23,24 Our data showed significantly greater numbers of ACE2-positive cells in the lungs from patients with Covid-19 and from patients with influenza than in those from uninfected controls. We found greater numbers of ACE2-positive endothelial cells and significant changes in endothelial morphology, a finding consistent with a central role of endothelial cells in the vascular phase of Covid-19. Endothelial cells in the specimens from patients with Covid-19 showed disruption of intercellular junctions, cell swelling, and a loss of contact with the basal membrane. The presence of SARS-CoV-2 virus within the endothelial cells, a finding consistent with other studies,25 suggests that direct viral effects as well as perivascular inflammation may contribute to the endothelial injury.

We report the presence of pulmonary intussusceptive angiogenesis and other pulmonary vascular features in the lungs of seven patients who died from Covid-19. Additional work is needed to relate our findings to the clinical course in these patients. To aid others in their research, our full data set is available on the Vivli platform (https://vivli.org/. opens in new tab) and can be requested with the use of the following digital object identifier: https://doi.org/10.25934/00005576. opens in new tab.

Supported by grants (HL94567 and HL134229, to Drs. Ackermann and Mentzer) from the National Institutes of Health, a grant from the Botnar Research Centre for Child Health (to Dr. Tzankov), a European Research Council Consolidator Grant (XHale) (771883, to Dr. Jonigk), and a grant (KFO311, to Dr. Jonigk) from Deutsche Forschungsgemeinschaft (Project Z2).

Disclosure forms provided by the authors are available with the full text of this article at NEJM.org.

This article was published on May 21, 2020, at NEJM.org.

We thank Kerstin Bahr, Jan Hinrich Braesen, Peter Braubach, Emily Brouwer, Annette Mueller Brechlin, Regina Engelhardt, Jasmin Haslbauer, Anne Hoefer, Nicole Kroenke, Thomas Menter, Mahtab Taleb Naghsh, Christina Petzold, Vincent Schmidt, and Pauline Tittmann for technical support; Peter Boor of the German Covid-19 registry; and Lynnette Sholl, Hans Kreipe, Hans Michael Kvasnicka, and Jean Connors for helpful comments. Dr. Jonigk thanks Anita Swiatlak for her continued support.

Author Affiliations

From the Institute of Pathology and Department of Molecular Pathology, Helios University Clinic Wuppertal, University of Witten–Herdecke, Wuppertal (M.A.), the Institute of Functional and Clinical Anatomy, University Medical Center of the Johannes Gutenberg University Mainz, Mainz (M.A.), the Institute of Pathology (M.K., F.L., C.W., H.S., D.J.), the Department of Cardiothoracic, Transplantation, and Vascular Surgery (A.H.), and the Clinic of Pneumology (T.W.), Hannover Medical School, and the German Center for Lung Research, Biomedical Research in Endstage and Obstructive Lung Disease Hannover (BREATH) (M.K., A.H., T.W., F.L., C.W., H.S., D.J.), Hannover — all in Germany; the Laboratory of Respiratory Diseases, BREATH, Department of Chronic Diseases, Metabolism, and Aging, KU Leuven, Leuven, Belgium (S.E.V., A.V.); the Institute of Pathology and Medical Genetics, University Hospital Basel, Basel, Switzerland (A.T.); and the Angiogenesis Foundation, Cambridge (W.W.L., V.W.L.), and the Laboratory of Adaptive and Regenerative Biology and the Division of Thoracic Surgery, Brigham and Women’s Hospital, Harvard Medical School, Boston (S.J.M.) — all in Massachusetts.

Address reprint requests to Dr. Mentzer at the Division of Thoracic Surgery, Brigham and Women’s Hospital, Harvard Medical School, 75 Francis St., Boston, MA 02115, or at smentzer@bwh.harvard.edu.

Supplementary Material

Supplementary AppendixPDF1523KB
Disclosure FormsPDF288KB

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

References

  1. 1.
    • Huang C 
    • Wang Y 
    • Li X 
    • et al.Clinical features of patients infected with 2019 novel coronavirus in Wuhan, China.Lancet. 2020; 395: 497-506View in Article 
  2. 2.
    • Klok FA 
    • Kruip M 
    • van der Meer NJM 
    • et al.Confirmation of the high cumulative incidence of thrombotic complications in critically ill ICU patients with COVID-19: An updated analysis.Thromb Res. 2020; 191: 148-150View in Article 
  3. 3.
    • Middeldorp S 
    • Coppens M 
    • van Haaps TF 
    • et al.Incidence of venous thromboembolism in hospitalized patients with COVID-19.J Thromb Haemost. 2020; 18: 1995-2002View in Article 
  4. 4.
    • Lodigiani C 
    • Iapichino G 
    • Carenzo L 
    • et al.Venous and arterial thromboembolic complications in COVID-19 patients admitted to an academic hospital in Milan, Italy.Thromb Res. 2020; 191: 9-14View in Article 
  5. 5.
    • Halushka MK 
    • Vander Heide RS.Myocarditis is rare in COVID-19 autopsies: cardiovascular findings across 277 postmortem examinations.Cardiovasc Pathol. 2021; 50107300View in Article 
  6. 6.
    • Tang N 
    • Li D 
    • Wang X 
    • Sun Z.Abnormal coagulation parameters are associated with poor prognosis in patients with novel coronavirus pneumonia.J Thromb Haemost. 2020; 18: 844-847View in Article 
  7. 7.
    • Ciceri F 
    • Beretta L 
    • Scandroglio AM 
    • et al.Microvascular COVID-19 lung vessels obstructive thromboinflammatory syndrome (MicroCLOTS): an atypical acute respiratory distress syndrome working hypothesis.Crit Care Resusc. 2020; 22: 95-97View in Article 
  8. 8.
    • Jin Y 
    • Ji W 
    • Yang H 
    • Chen S 
    • Zhang W 
    • Duan GEndothelial activation and dysfunction in COVID-19: from basic mechanisms to potential therapeutic approaches.Signal Transduct Target Ther. 2020; 5: 293View in Article 
  9. 9.
    • Wu KK 
    • Thiagarajan P.Role of endothelium in thrombosis and hemostasis.Annu Rev Med. 1996; 47: 315-331View in Article 
  10. 10.
    • Lenting PJ 
    • Christophe OD 
    • Denis CV.von Willebrand factor biosynthesis, secretion, and clearance: connecting the far ends.Blood. 2015; 125: 2019-2028View in Article 
  11. 11.
    • Conway EM.Thrombomodulin and its role in inflammation.Semin Immunopathol. 2012; 34: 107-125View in Article 
  12. 12.
    • Watanabe-Kusunoki K 
    • Nakazawa D 
    • Ishizu A 
    • Atsumi T.Thrombomodulin as a physiological modulator of intravascular injury.Front Immunol. 2020; 11575890View in Article 
  13. 13.
    • Goshua G 
    • Pine AB 
    • Meizlish ML 
    • et al.Endotheliopathy in COVID-19-associated coagulopathy: evidence from a single-centre, cross-sectional study.Lancet Haematol. 2020; 7: e575-ee82View in Article 
  14. 14.
    • Tong M 
    • Jiang Y 
    • Xia D 
    • et al.Elevated expression of serum endothelial cell adhesion molecules in COVID-19 patients.J Infect Dis. 2020; 222: 894-898View in Article 
  15. 15.
    • Bouck EG 
    • Denorme F 
    • Holle LA 
    • et al.COVID-19 and sepsis are associated with different abnormalities in plasma procoagulant and fibrinolytic activity.Arterioscler Thromb Vasc Biol. 2021; 41: 401-414View in Article 
  16. 16.
    • Jin X 
    • Duan Y 
    • Bao T 
    • et al.The values of coagulation function in COVID-19 patients.PLoS One. 2020; 15e0241329View in Article 
  17. 17.
    • Lowenstein CJ 
    • Solomon SD.Severe COVID-19 is a microvascular disease.Circulation. 2020; 142: 1609-1611View in Article 
  18. 18.
    • Teuwen LA 
    • Geldhof V 
    • Pasut A 
    • Carmeliet P.COVID-19: the vasculature unleashed.Nat Rev Immunol. 2020; 20: 389-391View in Article 
  19. 19.
    • Libby P 
    • Luscher T.COVID-19 is, in the end, an endothelial disease.Eur Heart J. 2020; 41: 3038-3044View in Article 
  20. 20.
    • Riollano-Cruz M 
    • Akkoyun E 
    • Briceno-Brito E 
    • et al.Multisystem inflammatory syndrome in children related to COVID-19: A New York City experience.J Med Virol. 2020;View in Article 
  21. 21.
    • Liao M 
    • Liu Y 
    • Yuan J 
    • et al.Single-cell landscape of bronchoalveolar immune cells in patients with COVID-19.Nat Med. 2020; 26: 842-844View in Article 
  22. 22.
    • Chua RL 
    • Lukassen S 
    • Trump S 
    • et al.COVID-19 severity correlates with airway epithelium-immune cell interactions identified by single-cell analysis.Nat Biotechnol. 2020; 38: 970-979View in Article 
  23. 23.
    • Xu G 
    • Qi F 
    • Li H 
    • et al.The differential immune responses to COVID-19 in peripheral and lung revealed by single-cell RNA sequencing.Cell Discov. 2020; 6: 73View in Article 
  24. 24.
    • Ren X 
    • Wen W 
    • Fan X 
    • et al.COVID-19 immune features revealed by a large-scale single-cell transcriptome atlas.Cell. 2021; 184 (e19): 1895-1913View in Article 
  25. 25.
    • Schulte-Schrepping J 
    • Reusch N 
    • Paclik D 
    • et al.Severe COVID-19 is marked by a dysregulated myeloid cell compartment.cell. 2020; 182 (e23): 1419-1440View in Article 
  26. 26.
    • Ziegler CGK 
    • Allon SJ 
    • Nyquist SK 
    • et al.SARS-CoV-2 receptor ACE2 is an interferon-stimulated gene in human airway epithelial cells and is detected in specific cell subsets across tissues.Cell. 2020; 181 (e19): 1016-1035View in Article 
  27. 27.
    • Wang A 
    • Chiou J 
    • Poirion OB 
    • et al.Single-cell multiomic profiling of human lungs reveals cell-type-specific and age-dynamic control of SARS-CoV2 host genes.Elife. 2020; 9View in Article 
  28. 28.
    • Singh M 
    • Bansal V 
    • Feschotte C.A single-cell RNA expression map of human coronavirus entry factors.Cell Rep. 2020; 32108175View in Article 
  29. 29.
    • Qi F 
    • Qian S 
    • Zhang S 
    • Zhang Z.Single cell RNA sequencing of 13 human tissues identify cell types and receptors of human coronaviruses.Biochem Biophys Res Commun. 2020; 526: 135-140View in Article 
  30. 30.
    • Muus C 
    • Luecken MD 
    • Eraslan G 
    • et al.Single-cell meta-analysis of SARS-CoV-2 entry genes across tissues and demographics.Nat Med. 2021; 27: 546-559View in Article 
  31. 31.
    • Zonneveld R 
    • Martinelli R 
    • Shapiro NI 
    • Kuijpers TW 
    • Plotz FB 
    • Carman CV.Soluble adhesion molecules as markers for sepsis and the potential pathophysiological discrepancy in neonates, children and adults.Crit Care. 2014; 18: 204View in Article 
  32. 32.
    • Hamming I 
    • Timens W 
    • Bulthuis ML 
    • Lely AT 
    • Navis G 
    • van Goor H.Tissue distribution of ACE2 protein, the functional receptor for SARS coronavirus. A first step in understanding SARS pathogenesis.J Pathol. 2004; 203: 631-637View in Article 
  33. 33.
    • Varga Z 
    • Flammer AJ 
    • Steiger P 
    • et al.Endothelial cell infection and endotheliitis in COVID-19.Lancet. 2020; 395: 1417-1418View in Article 
  34. 34.
    • Goldsmith CS 
    • Miller SE 
    • Martines RB 
    • Bullock HA 
    • Zaki SR.Electron microscopy of SARS-CoV-2: a challenging task.Lancet. 2020; 395: e99View in Article 
  35. 35.
    • McCracken IR 
    • Saginc G 
    • He L 
    • et al.Lack of evidence of angiotensin-converting enzyme 2 expression and replicative infection by SARS-CoV-2 in human endothelial cells.Circulation. 2021; 143: 865-868View in Article 
  36. 36.
    • Chen L 
    • Li X 
    • Chen M 
    • Feng Y 
    • Xiong C.The ACE2 expression in human heart indicates new potential mechanism of heart injury among patients infected with SARS-CoV-2.Cardiovasc Res. 2020; 116: 1097-1100View in Article 
  37. 37.
    • Deinhardt-Emmer S 
    • Bottcher S 
    • Haring C 
    • et al.SARS-CoV-2 causes severe epithelial inflammation and barrier dysfunction.J Virol. 2021;View in Article 
  38. 38.
    • Wang P 
    • Luo R 
    • Zhang M 
    • et al.A cross-talk between epithelium and endothelium mediates human alveolar-capillary injury during SARS-CoV-2 infection.Cell Death Dis. 2020; 11: 1042View in Article 
  39. 39.
    • Schaefer IM 
    • Padera RF 
    • Solomon IH 
    • et al.In situ detection of SARS-CoV-2 in lungs and airways of patients with COVID-19.Mod Pathol. 2020; 33: 2104-2114View in Article 
  40. 40.
    • Nascimento Conde J 
    • Schutt WR 
    • Gorbunova EE 
    • Mackow ERRecombinant ACE2 expression is required for SARS-CoV-2 to infect primary human endothelial cells and induce inflammatory and procoagulative responses.mBio. 2020; 11View in Article 
  41. 41.
    • Liu Y 
    • Garron TM 
    • Chang Q 
    • et al.Cell-Type Apoptosis in Lung during SARS-CoV-2 Infection.Pathogens. 2021; 10View in Article 
  42. 42.
    • Klein SL 
    • Pekosz A 
    • Park HS 
    • et al.Sex, age, and hospitalization drive antibody responses in a COVID-19 convalescent plasma donor population.J Clin Invest. 2020; 130: 6141-6150View in Article 
  43. 43.
    • Folsch H 
    • Pypaert M 
    • Schu P 
    • Mellman I.Distribution and function of AP-1 clathrin adaptor complexes in polarized epithelial cells.J Cell Biol. 2001; 152: 595-606View in Article 
  44. 44.
    • Angelini DJ 
    • Su Q 
    • Kolosova IA 
    • et al.Hypoxia-induced mitogenic factor (HIMF/FIZZ1/RELM alpha) recruits bone marrow-derived cells to the murine pulmonary vasculature.PLoS One. 2010; 5: e11251View in Article 
  45. 45.
    • Ackermann M 
    • Verleden SE 
    • Kuehnel M 
    • et al.Pulmonary vascular endothelialitis, thrombosis, and angiogenesis in Covid-19.N Engl J Med. 2020; 383: 120-128View in Article 
  46. 46.
    • Nicolai L 
    • Leunig A 
    • Brambs S 
    • et al.Immunothrombotic dysregulation in COVID-19 pneumonia is associated with respiratory failure and coagulopathy.Circulation. 2020; 142: 1176-1189View in Article 
  47. 47.
    • Bradley BT 
    • Maioli H 
    • Johnston R 
    • et al.Histopathology and ultrastructural findings of fatal COVID-19 infections in Washington State: a case series.Lancet. 2020; 396: 320-332View in Article 
  48. 48.
    • Hanley B 
    • Naresh KN 
    • Roufosse C 
    • et al.Histopathological findings and viral tropism in UK patients with severe fatal COVID-19: a post-mortem study.Lancet Microbe. 2020; 1: e245-ee53View in Article 
  49. 49.
    • Bois MC 
    • Boire NA 
    • Layman AJ 
    • et al.COVID-19-associated Non-Occlusive Fibrin Microthrombi in the Heart.Circulation. 2020;View in Article 
  50. 50.
    • Kattula S 
    • Byrnes JR 
    • Wolberg AS.Fibrinogen and fibrin in hemostasis and thrombosis.Arterioscler Thromb Vasc Biol. 2017; 37: e13-e21View in Article 
  51. 51.
    • Bongiovanni D 
    • Klug M 
    • Lazareva O 
    • et al.SARS-CoV-2 infection is associated with a pro-thrombotic platelet phenotype.Cell Death Dis. 2021; 12: 50View in Article 
  52. 52.
    • Manne BK 
    • Denorme F 
    • Middleton EA 
    • et al.Platelet gene expression and function in patients with COVID-19.Blood. 2020; 136: 1317-1329View in Article 
  53. 53.
    • Arepally GM 
    • Ortel TL.Heparin-induced thrombocytopenia.Annu Rev Med. 2010; 61: 77-90View in Article 
  54. 54.
    • Crikis S 
    • Zhang XM 
    • Dezfouli S 
    • et al.Anti-inflammatory and anticoagulant effects of transgenic expression of human thrombomodulin in mice.Am J Transplant. 2010; 10: 242-250View in Article 
  55. 55.
    • Ito T 
    • Thachil J 
    • Asakura H 
    • Levy JH 
    • Iba T.Thrombomodulin in disseminated intravascular coagulation and other critical conditions-a multi-faceted anticoagulant protein with therapeutic potential.Crit Care. 2019; 23: 280View in Article 
  56. 56.
    • Miller SE 
    • Goldsmith CS.Caution in identifying coronaviruses by electron microscopy.J Am Soc Nephrol. 2020; 31: 2223-2224View in Article 
  57. 57.
    • Smith CM 
    • Kulkarni H 
    • Radhakrishnan P 
    • et al.Ciliary dyskinesia is an early feature of respiratory syncytial virus infection.Eur Respir J. 2014; 43: 485-496View in Article 
  58. 58.
    • Sungnak W 
    • Huang N 
    • Becavin C 
    • et al.SARS-CoV-2 entry factors are highly expressed in nasal epithelial cells together with innate immune genes.Nat Med. 2020; 26: 681-687View in Article 
  59. 59.
    • Mason RJ.Pathogenesis of COVID-19 from a cell biology perspective.Eur Respir J. 2020; 55View in Article 
  60. 60.
    • Jahani M 
    • Dokaneheifard S 
    • Mansouri K.Hypoxia: A key feature of COVID-19 launching activation of HIF-1 and cytokine storm.J Inflamm (Lond). 2020; 17: 33View in Article 
  61. 61.
    • Dickson RP.Lung microbiota and COVID-19 severity.Nat Microbiol. 2021; 6: 1217-1218View in Article 
  62. 62.
    • Thiyagarajan M 
    • Cheng T 
    • Zlokovic BV.Endothelial cell protein C receptor: role beyond endothelium?.Circ Res. 2007; 100: 155-157View in Article 
  63. 63.
    • Faust SN 
    • Levin M 
    • Harrison OB 
    • et al.Dysfunction of endothelial protein C activation in severe meningococcal sepsis.N Engl J Med. 2001; 345: 408-416View in Article 
  64. 64.
    • Lerolle N 
    • Carlotti A 
    • Melican K 
    • et al.Assessment of the interplay between blood and skin vascular abnormalities in adult purpura fulminans.Am J Respir Crit Care Med. 2013; 188: 684-692View in Article 
  65. 65.
    • Moxon CA 
    • Wassmer SC 
    • Milner Jr., DA 
    • et al.Loss of endothelial protein C receptors links coagulation and inflammation to parasite sequestration in cerebral malaria in African children.Blood. 2013; 122: 842-851View in Article 
  66. 66.
    • Isermann B 
    • Hendrickson SB 
    • Zogg M 
    • et al.Endothelium-specific loss of murine thrombomodulin disrupts the protein C anticoagulant pathway and causes juvenile-onset thrombosis.J Clin Invest. 2001; 108: 537-546View in Article 
  67. 67.
    • Francischetti IMB 
    • Toomer K 
    • Zhang Y 
    • et al.Upregulation of pulmonary tissue factor, loss of thrombomodulin and immunothrombosis in SARS-CoV-2 infection.EClinicalMedicine. 2021; 39101069View in Article 
  68. 68.
    • Wang C 
    • Xie J 
    • Zhao L 
    • et al.Alveolar macrophage dysfunction and cytokine storm in the pathogenesis of two severe COVID-19 patients.EBioMedicine. 2020; 57102833View in Article 
  69. 69.
    • Dorward DA 
    • Russell CD 
    • Um IH 
    • et al.Tissue-specific immunopathology in fatal COVID-19.Am J Respir Crit Care Med. 2021; 203: 192-201View in Article 
  70. 70.
    • Melms JC 
    • Biermann J 
    • Huang H 
    • et al.A molecular single-cell lung atlas of lethal COVID-19.Nature. 2021; 595: 114-119View in Article 
  71. 71.
    • Silvin A 
    • Chapuis N 
    • Dunsmore G 
    • et al.Elevated calprotectin and abnormal myeloid cell subsets discriminate severe from Mild COVID-19.Cell. 2020; 182 (e18): 1401-1418View in Article 
  72. 72.
    • Giamarellos-Bourboulis EJ 
    • Netea MG 
    • Rovina N 
    • et al.Complex immune dysregulation in COVID-19 patients with severe respiratory failure.Cell Host Microbe. 2020; 27 (e3): 992-1000View in Article 
  73. 73.
    • Szabo PA 
    • Dogra P 
    • Gray JI 
    • et al.Longitudinal profiling of respiratory and systemic immune responses reveals myeloid cell-driven lung inflammation in severe COVID-19.Immunity. 2021; 54 (e6): 797-814View in Article 
  74. 74.
    • Niles MA 
    • Gogesch P 
    • Kronhart S 
    • et al.Macrophages and dendritic cells are not the major source of Pro-inflammatory Cytokines upon SARS-CoV-2 infection.Front Immunol. 2021; 12647824View in Article 
  75. 75.
    • Zhang D 
    • Guo R 
    • Lei L 
    • et al.Frontline Science: COVID-19 infection induces readily detectable morphologic and inflammation-related phenotypic changes in peripheral blood monocytes.J Leukoc Biol. 2021; 109: 13-22View in Article 
  76. 76.
    • Zhou YG 
    • Fu BQ 
    • Zheng XH 
    • et al.Pathogenic T-cells and inflammatory monocytes incite inflammatory storms in severe COVID-19 patients.Natl Sci Rev. 2020; 7: 998-1002View in Article 
  77. 77.
    • Kuri-Cervantes L 
    • Pampena MB 
    • Meng W 
    • et al.Comprehensive mapping of immune perturbations associated with severe COVID-19.Sci Immunol. 2020; 5View in Article 
  78. 78.
    • Thevarajan I 
    • Nguyen THO 
    • Koutsakos M 
    • et al.Breadth of concomitant immune responses prior to patient recovery: a case report of non-severe COVID-19.Nat Med. 2020; 26: 453-455View in Article 
  79. 79.
    • Rahman A 
    • Fazal F.Hug tightly and say goodbye: role of endothelial ICAM-1 in leukocyte transmigration.Antioxid Redox Signal. 2009; 11: 823-839View in Article 
  80. 80.
    • Jansen F 
    • Yang X 
    • Baumann K 
    • et al.Endothelial microparticles reduce ICAM-1 expression in a microRNA-222-dependent mechanism.J Cell Mol Med. 2015; 19: 2202-2214View in Article 
  81. 81.
    • Abers MS 
    • Delmonte OM 
    • Ricotta EE 
    • et al.An immune-based biomarker signature is associated with mortality in COVID-19 patients.JCI Insight. 2021; 6View in Article 
  82. 82.
    • Ueland T 
    • Holter JC 
    • Holten AR 
    • et al.Distinct and early increase in circulating MMP-9 in COVID-19 patients with respiratory failure.J Infect. 2020; 81: e41-ee3View in Article 
  83. 83.
    • Syed F 
    • Li W 
    • Relich RF 
    • et al.Excessive matrix metalloproteinase-1 and hyperactivation of endothelial cells occurred in COVID-19 patients and were associated with the severity of COVID-19.J Infect Dis. 2021; 224: 60-69View in Article 
  84. 84.
    • Dittmayer C 
    • Meinhardt J 
    • Radbruch H 
    • et al.Why misinterpretation of electron micrographs in SARS-CoV-2-infected tissue goes viral.Lancet. 2020; 396: e64-ee5View in Article 
  85. 85.
    • Mackman N.New insights into the mechanisms of venous thrombosis.J Clin Invest. 2012; 122: 2331-2336View in Article 
  86. 86.
    • Bovill EG 
    • van der Vliet A.Venous valvular stasis-associated hypoxia and thrombosis: what is the link?.Annu Rev Physiol. 2011; 73: 527-545View in Article 
  87. 87.
    • Owens 3rd, AP 
    • Mackman N.Microparticles in hemostasis and thrombosis.Circ Res. 2011; 108: 1284-1297View in Article 
  88. 88.
    • Morel O 
    • Marchandot B 
    • Jesel L 
    • et al.Microparticles in COVID-19 as a link between lung injury extension and thrombosis.ERJ Open Res. 2021; 7View in Article 
  89. 89.
    • Beristain-Covarrubias N 
    • Perez-Toledo M 
    • Thomas MR 
    • Henderson IR 
    • Watson SP 
    • Cunningham AF.Understanding infection-induced thrombosis: lessons learned from animal models.Front Immunol. 2019; 10: 2569View in Article 
  90. 90.
    • Connors JM 
    • Levy JH.COVID-19 and its implications for thrombosis and anticoagulation.Blood. 2020; 135: 2033-2040View in Article 
  91. 91.
    • Wu Z 
    • Liu MC 
    • Liang M 
    • Fu J.Sirt1 protects against thrombomodulin down-regulation and lung coagulation after particulate matter exposure.Blood. 2012; 119: 2422-2429View in Article 
  92. 92.
    • Su EJ 
    • Geyer M 
    • Wahl M 
    • et al.The thrombomodulin analog Solulin promotes reperfusion and reduces infarct volume in a thrombotic stroke model.J Thromb Haemost. 2011; 9: 1174-1182View in Article 
  93. 93.
    • Nakamura Y 
    • Nakano T 
    • Irie K 
    • et al.Recombinant human soluble thrombomodulin ameliorates cerebral ischemic injury through a high-mobility group box 1 inhibitory mechanism without hemorrhagic complications in mice.J Neurol Sci. 2016; 362: 278-282 View in Article 
  94. 94.
    • Yamakawa K 
    • Aihara M 
    • Ogura H 
    • Yuhara H 
    • Hamasaki T 
    • Shimazu T.Recombinant human soluble thrombomodulin in severe sepsis: a systematic review and meta-analysis.J Thromb Haemost. 2015; 13: 508-519View in Article 

Article Info

Publication History

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

Identification

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

Copyright

© 2022 Published by Elsevier B.V.

User License

Creative Commons Attribution – NonCommercial – NoDerivs (CC BY-NC-ND 4.0) | How you can reuseInformation Icon

ScienceDirect

Access this article on ScienceDirect

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

Study reveals mouth as primary source of COVID-19 infection

While most COVID-19 research has focused on the nose and lungs, this is the first study to identify the mouth as a primary site for coronavirus infection and underscores the importance of wearing a face covering and physical distancing.

By University Communications, Thursday, October 29th, 2020

A team of researchers led by the University of North Carolina at Chapel Hill and the National Institute of Dental and Craniofacial Research reveals coronavirus can take hold in the salivary glands where it replicates, and in some cases, leads to prolonged disease when infected saliva is swallowed into the gastrointestinal tract or aspirated to the lungs where it can lead to pneumonia.

While most COVID-19 research has focused on the nose and lungs, this is the first study to identify the mouth as a primary site for coronavirus infection and underscores the importance of wearing a face covering and physical distancing. The results have not been peer-reviewed.

“Our results show oral infection of COVID-19 may be underappreciated,” said senior study author Kevin M. Byrd, research instructor at the UNC Adams School of Dentistry and the Anthony R. Volpe Research Scholar at the American Dental Association Science and Research Institute. “Like nasal infection, oral infection could underlie the asymptomatic spread that makes this disease so hard to contain.”

Byrd along with Blake Warner, chief of the Salivary Disorders Unit at the National Institute of Dental and Craniofacial Research, coordinated the research conducted at the National Institutes of Health, Wellcome Sanger Institute, UNC Marsico Lung Institute and the J. Craig Venter Institute.

Researchers are just beginning to explore the oral symptoms patients experience during COVID-19, such as loss of taste or smell and persistent dry mouth.

In the study, researchers report preliminary results from a clinical trial of 40 subjects with COVID-19 which showed sloughed epithelial cells lining the mouth can be infected with SARS-CoV-2, the coronavirus that causes COVID-19. The amount of virus in patient saliva was positively correlated with taste and smell changes, according to the study.

Relying on oral cell identity maps, researchers also looked at where in the mouth the virus infects. They surveyed oral tissues with the highest levels of ACE2, the receptor that helps coronavirus grab and invade human cells.

Based on ACE2 expression and analysis of cadaver tissue, the most likely sites of infection in the mouth are the salivary glands, tongue and tonsil, the study showed.

The findings provide more evidence of the role of saliva in COVID-19. COVID-19 infection, specifically in the mouth, can allow the virus to spread internally and to others as the infected person breathes, speaks and coughs.

For More Information: https://www.unc.edu/posts/2020/10/29/study-reveals-mouth-as-primary-source-of-covid-19-infection/

Pathological findings in organs and tissues of patients with COVID-19: A systematic review

Authors: Sasha Peiris 1 2Hector Mesa 3Agnes Aysola 4Juan Manivel 5Joao Toledo 1 2Marcio Borges-Sa 6Sylvain Aldighieri 1 2Ludovic Reveiz 2 7

Abstract

Background: Coronavirus disease (COVID-19) is the pandemic caused by SARS-CoV-2 that has caused more than 2.2 million deaths worldwide. We summarize the reported pathologic findings on biopsy and autopsy in patients with severe/fatal COVID-19 and documented the presence and/or effect of SARS-CoV-2 in all organs.

Methods and findings: A systematic search of the PubMed, Embase, MedRxiv, Lilacs and Epistemonikos databases from January to August 2020 for all case reports and case series that reported histopathologic findings of COVID-19 infection at autopsy or tissue biopsy was performed. 603 COVID-19 cases from 75 of 451 screened studies met inclusion criteria. The most common pathologic findings were lungs: diffuse alveolar damage (DAD) (92%) and superimposed acute bronchopneumonia (27%); liver: hepatitis (21%), heart: myocarditis (11.4%). Vasculitis was common only in skin biopsies (25%). Microthrombi were described in the placenta (57.9%), lung (38%), kidney (20%), Central Nervous System (CNS) (18%), and gastrointestinal (GI) tract (2%). Injury of endothelial cells was common in the lung (18%) and heart (4%). Hemodynamic changes such as necrosis due to hypoxia/hypoperfusion, edema and congestion were common in kidney (53%), liver (48%), CNS (31%) and GI tract (18%). SARS-CoV-2 viral particles were demonstrated within organ-specific cells in the trachea, lung, liver, large intestine, kidney, CNS either by electron microscopy, immunofluorescence, or immunohistochemistry. Additional tissues were positive by Polymerase Chain Reaction (PCR) tests only. The included studies were from numerous countries, some were not peer reviewed, and some studies were performed by subspecialists, resulting in variable and inconsistent reporting or over statement of the reported findings.

Conclusions: The main pathologic findings of severe/fatal COVID-19 infection are DAD, changes related to coagulopathy and/or hemodynamic compromise. In addition, according to the observed organ damage myocarditis may be associated with sequelae.

For More Information: https://pubmed.ncbi.nlm.nih.gov/33909679/

Long covid: Damage to multiple organs presents in young, low risk patients

Authors: Gareth Iacobucci BMJ 2020; 371 doi: https://doi.org/10.1136/bmj.m4470 (Published 17 November 2020)Cite this as: BMJ 2020;371:m4470

Young, low risk patients with ongoing symptoms of covid-19 had signs of damage to multiple organs four months after initially being infected, a preprint study has suggested.1

Initial data from 201 patients suggest that almost 70% had impairments in one or more organs four months after their initial symptoms of SARS-CoV-2 infection.

The results emerged as the NHS announced plans to establish a network of more than 40 long covid specialist clinics across England this month to help patients with long term symptoms of infection.

The prospective Coverscan study examined the impact of long covid (persistent symptoms three months post infection) across multiple organs in low risk people who are relatively young and had no major underlying health problems. Assessment was done using results from magnetic resonance image scans, blood tests, and online questionnaires.

The research has not yet been peer reviewed and could not establish a causal link between organ impairment and infection. But the authors said the results had “implications not only for [the] burden of long covid but also public health approaches which have assumed low risk in young people with no comorbidities.”

The study enrolled participants at two UK sites in Oxford and London between April and August 2020. Two hundred and one individuals (mean age 44 (standard deviation 11.0) years) completed assessments after SARS-CoV-2 infection a median of 140 days after initial symptoms.

Participants were eligible if they tested positive for SARS-CoV-2 by random polymerase chain reaction swab (n=62), a positive antibody test (n=63), or had typical symptoms and were determined to have covid-19 by two independent clinicians (n=73).

The prevalence of pre-existing conditions was low (obesity: 20%, hypertension: 6%, diabetes: 2%, heart disease: 4%), and less than a fifth (18%) of individuals had been hospitalised with covid-19.

The most commonly reported ongoing symptoms—regardless of hospitalization status—were fatigue (98%), muscle ache (88%), shortness of breath (87%), and headache (83%). There was evidence of mild organ impairment in the heart (32% of patients), lungs (33%), kidneys (12%), liver (10%), pancreas (17%), and spleen (6%).

For More Information: https://www.bmj.com/content/371/bmj.m4470

Pathological findings in organs and tissues of patients with COVID-19: A systematic review

  1. Authors: Sasha Peiris, Hector Mesa, Agnes Aysola, Juan Manivel, Joao Toledo, Marcio Borges-Sa, Sylvain Aldighieri, Ludovic Reveiz

Abstract

Background

Coronavirus disease (COVID-19) is the pandemic caused by SARS-CoV-2 that has caused more than 2.2 million deaths worldwide. We summarize the reported pathologic findings on biopsy and autopsy in patients with severe/fatal COVID-19 and documented the presence and/or effect of SARS-CoV-2 in all organs.

Methods and findings

A systematic search of the PubMed, Embase, MedRxiv, Lilacs and Epistemonikos databases from January to August 2020 for all case reports and case series that reported histopathologic findings of COVID-19 infection at autopsy or tissue biopsy was performed. 603 COVID-19 cases from 75 of 451 screened studies met inclusion criteria. The most common pathologic findings were lungs: diffuse alveolar damage (DAD) (92%) and superimposed acute bronchopneumonia (27%); liver: hepatitis (21%), heart: myocarditis (11.4%). Vasculitis was common only in skin biopsies (25%). Microthrombi were described in the placenta (57.9%), lung (38%), kidney (20%), Central Nervous System (CNS) (18%), and gastrointestinal (GI) tract (2%). Injury of endothelial cells was common in the lung (18%) and heart (4%). Hemodynamic changes such as necrosis due to hypoxia/hypoperfusion, edema and congestion were common in kidney (53%), liver (48%), CNS (31%) and GI tract (18%). SARS-CoV-2 viral particles were demonstrated within organ-specific cells in the trachea, lung, liver, large intestine, kidney, CNS either by electron microscopy, immunofluorescence, or immunohistochemistry. Additional tissues were positive by Polymerase Chain Reaction (PCR) tests only. The included studies were from numerous countries, some were not peer reviewed, and some studies were performed by subspecialists, resulting in variable and inconsistent reporting or over statement of the reported findings.

Conclusions

The main pathologic findings of severe/fatal COVID-19 infection are DAD, changes related to coagulopathy and/or hemodynamic compromise. In addition, according to the observed organ damage myocarditis may be associated with sequelae.

For More Information: https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0250708

Late Complications of COVID-19; a Systematic Review of Current Evidence

Authors: SeyedAhmad SeyedAlinaghi,1Amir Masoud Afsahi,2Mehrzad MohsseniPour,1Farzane Behnezhad,3Mohammad Amin Salehi,1Alireza Barzegary,4Pegah Mirzapour,1Esmaeil Mehraeen,5,* and Omid Dadras6

Introduction

Introduction:

COVID-19 is a new rapidly spreading epidemic. The symptoms of this disease could be diverse as the virus can affect any organ in the body of an infected person. This study aimed to investigate the available evidence for long-term complications of COVID-19.

Methods:

This study was a systematic review of current evidence conducted in November 2020 to investigate probable late and long-term complications of COVID-19. We performed a systematic search, using the keywords, in online databases including PubMed, Scopus, Science Direct, Up to Date, and Web of Science, to find papers published from December 2019 to October 2020. Peer-reviewed original papers published in English, which met the eligibility criteria were included in the final report. Addressing non-human studies, unavailability of the full-text document, and duplicated results in databases, were characteristics that led to exclusion of the papers from review.

Results:

The full-texts of 65 articles have been reviewed. We identified 10 potential late complications of COVID-19. A review of studies showed that lung injuries (n=31), venous/arterial thrombosis (n=28), heart injuries (n=26), cardiac/brain stroke (n=23), and neurological injuries (n=20) are the most frequent late complications of COVID-19.

For More Information: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7927752/