Colchicine: A Possible COVID-19 Long haul Cardiac Therapy

Last Updated: December 16, 2021

Last Updated: December 16, 2021

Colchicine is an anti-inflammatory drug that is used to treat a variety of conditions, including gout, recurrent pericarditis, and familial Mediterranean fever.1 Recently, the drug has been shown to potentially reduce the risk of cardiovascular events in those with coronary artery disease.2 Colchicine has several potential mechanisms of action, including reducing the chemotaxis of neutrophils, inhibiting inflammasome signaling, and decreasing the production of cytokines, such as interleukin-1 beta.3 When colchicine is administered early in the course of COVID-19, these mechanisms could potentially mitigate or prevent inflammation-associated manifestations of the disease. These anti-inflammatory properties coupled with the drug’s limited immunosuppressive potential, favorable safety profile, and widespread availability have prompted investigation of colchicine for the treatment of COVID-19.

Recommendations

  • The COVID-19 Treatment Guidelines Panel (the Panel) recommends against the use of colchicine for the treatment of nonhospitalized patients with COVID-19, except in a clinical trial (BIIa).
  • The Panel recommends against the use of colchicine for the treatment of hospitalized patients with COVID-19 (AI).

Rationale

For Nonhospitalized Patients With COVID-19

COLCORONA, a large randomized placebo-controlled trial that evaluated colchicine in outpatients with COVID-19, did not reach its primary efficacy endpoint of reducing hospitalizations and death.4 However, in the subset of patients whose diagnosis was confirmed by a positive SARS-CoV-2 polymerase chain reaction (PCR) result from a nasopharyngeal (NP) swab, a slight reduction in hospitalizations was observed among those who received colchicine.

PRINCIPLE, another randomized, open-label, adaptive-platform trial that evaluated colchicine versus usual care, was stopped for futility when no significant difference in time to first self-reported recovery from COVID-19 between the colchicine and usual care recipients was found.5

The PRINCIPLE trial showed no benefit of colchicine, and the larger COLCORONA trial failed to reach its primary endpoint, found only a very modest effect of colchicine in the subgroup of patients with positive SARS-CoV-2 PCR results, and reported more gastrointestinal adverse events in those receiving colchicine. Therefore, the Panel recommends against the use of colchicine for the treatment of COVID-19 in nonhospitalized patients, except in a clinical trial (BIIa).

For Hospitalized Patients With COVID-19

In the RECOVERY trial, a large randomized trial in hospitalized patients with COVID-19, colchicine demonstrated no benefit with regard to 28-day mortality or any secondary outcomes.6 Based on the results from this large trial, the Panel recommends against the use of colchicine for the treatment of COVID-19 in hospitalized patients (AI).

Clinical Data for COVID-19

Colchicine in Nonhospitalized Patients With COVID-19

The COLCORONA Trial

The COLCORONA trial was a contactless, double-blind, placebo-controlled, randomized trial in outpatients who received a diagnosis of COVID-19 within 24 hours of enrollment. Participants were aged ≥70 years or aged ≥40 years with at least 1 of the following risk factors for COVID-19 complications: body mass index ≥30, diabetes mellitus, uncontrolled hypertension, known respiratory disease, heart failure or coronary disease, fever ≥38.4°C within the last 48 hours, dyspnea at presentation, bicytopenia, pancytopenia, or the combination of high neutrophil count and low lymphocyte count. Participants were randomized 1:1 to receive colchicine 0.5 mg twice daily for 3 days and then once daily for 27 days or placebo. The primary endpoint was a composite of death or hospitalization by Day 30; secondary endpoints included components of the primary endpoint, as well as the need for mechanical ventilation by Day 30. Participants reported by telephone the occurrence of any study endpoints at 15 and 30 days after randomization; in some cases, clinical data were confirmed or obtained by medical chart reviews.4

Results

  • The study enrolled 4,488 participants.
  • The primary endpoint occurred in 104 of 2,235 participants (4.7%) in the colchicine arm and 131 of 2,253 participants (5.8%) in the placebo arm (OR 0.79; 95% CI, 0.61–1.03; P = 0.08).
  • There were no statistically significant differences in the secondary outcomes between the arms.
  • In a prespecified analysis of 4,159 participants who had a SARS-CoV-2 diagnosis confirmed by PCR testing of an NP specimen (93% of those enrolled), those in the colchicine arm were less likely to reach the primary endpoint (96 of 2,075 participants [4.6%]) than those in the placebo arm (126 of 2,084 participants [6.0%]; OR 0.75; 95% CI, 0.57–0.99; P = 0.04). In this subgroup of patients with PCR-confirmed SARS-CoV-2 infection, there were fewer hospitalizations (a secondary outcome) in the colchicine arm (4.5% of patients) than in the placebo arm (5.9% of patients; OR 0.75; 95% CI, 0.57–0.99).
  • More participants in the colchicine arm experienced gastrointestinal adverse events, including diarrhea which occurred in 13.7% of colchicine recipients versus 7.3% of placebo recipients (P < 0.0001). Unexpectedly, more pulmonary emboli were reported in the colchicine arm than in the placebo arm (11 events [0.5% of patients] vs. 2 events [0.1% of patients]; P= 0.01).

Limitations

  • Due to logistical difficulties with staffing, the trial was stopped at approximately 75% of the target enrollment, which may have limited the study’s power to detect differences for the primary outcome.
  • There was uncertainty as to the accuracy of COVID-19 diagnoses in presumptive cases.
  • Some patient-reported clinical outcomes were potentially misclassified.

The PRINCIPLE Trial

PRINCIPLE is a randomized, open-label, platform trial that evaluated colchicine in symptomatic, nonhospitalized patients with COVID-19 who were aged ≥65 years or aged ≥18 years with comorbidities or shortness of breath, and who had symptoms for ≤14 days. Participants were randomized to receive colchicine 0.5 mg daily for 14 days or usual care. The coprimary endpoints, which included time to first self-reported recovery or hospitalization or death due to COVID-19 by Day 28, were analyzed using a Bayesian model. Participants were followed through symptom diaries that they completed online daily; those who did not complete the diaries were contacted by telephone on Days 7, 14, and 29. The investigators developed a prespecified criterion for futility, specifying a clinically meaningful benefit in time to first self-reported recovery as a hazard ratio ≥1.2, corresponding to about 1.5 days of faster recovery in the colchicine arm.

Results

  • The study enrolled 4,997 participants: 212 participants were randomized to receive colchicine; 2,081 to receive usual care alone; and 2,704 to receive other treatments.
  • The prespecified primary analysis included participants with SARS-CoV-2 positive test results (156 in the colchicine arm; 1,145 in the usual care arm; and 1,454 in the other treatments arm).
  • The trial was stopped early because the criterion for futility was met; the median time to self-reported recovery was similar in the colchicine arm and the usual care arm (HR 0.92; 95% CrI, 0.72–1.16).
  • Analyses of self-reported time to recovery and hospitalizations or death due to COVID-19 among concurrent controls also showed no significant differences between the colchicine and usual care arms.
  • There were no statistically significant differences in the secondary outcomes between the colchicine and usual care arms in both the primary analysis population and in subgroups, including subgroups based on symptom duration, baseline disease severity, age, or comorbidities.
  • The occurrence of adverse events was similar in the colchicine and usual care arms.

Limitations

  • The design of the study was open-label treatment.
  • The sample size of the colchicine arm was small.

Colchicine in Hospitalized Patients With COVID-19

The RECOVERY Trial

In the RECOVERY trial, hospitalized patients with COVID-19 were randomized to receive colchicine (1 mg loading dose, followed by 0.5 mg 12 hours later, and then 0.5 mg twice daily for 10 days or until discharge) or usual care.6

Results

  • The study enrolled 11,340 participants.
  • At randomization, 10,603 patients (94%) were receiving corticosteroids.
  • The primary endpoint of all-cause mortality at Day 28 occurred in 1,173 of 5,610 participants (21%) in the colchicine arm and 1,190 of 5,730 participants (21%) in the placebo arm (rate ratio 1.01; 95% CI, 0.93–1.10; P = 0.77).
  • There were no statistically significant differences between the arms for the secondary outcomes of median time to being discharged alive, discharge from the hospital within 28 days, and receipt of mechanical ventilation or death.
  • The incidence of new cardiac arrhythmias, bleeding events, and thrombotic events was similar in the 2 arms. Two serious adverse events were attributed to colchicine: 1 case of severe acute kidney injury and one case of rhabdomyolysis.

Limitations

  • The trial’s open-label design may have introduced bias for assessing some of the secondary endpoints.

The GRECCO-19 Trial

GRECCO-19 was a small, prospective, open-label randomized clinical trial in 105 patients hospitalized with COVID-19 across 16 hospitals in Greece. Patients were assigned 1:1 to receive standard of care with colchicine (1.5 mg loading dose, followed by 0.5 mg after 60 minutes and then 0.5 mg twice daily until hospital discharge or for up to 3 weeks) or standard of care alone.7

Results

  • Fewer patients in the colchicine arm (1 of 55 patients) than in the standard of care arm (7 of 50 patients) reached the primary clinical endpoint of deterioration in clinical status from baseline by 2 points on a 7-point clinical status scale (OR 0.11; 95% CI, 0.01–0.96).
  • Participants in the colchicine group were significantly more likely to experience diarrhea (occurred in 45.5% of participants in the colchicine arm vs. 18.0% in the standard of care arm; P = 0.003).

Limitations

  • The overall sample size and the number of clinical events reported were small.
  • The study design was open-label treatment assignment.

The results of several small randomized trials and retrospective cohort studies that have evaluated various doses and durations of colchicine in hospitalized patients with COVID-19 have been published in peer-reviewed journals or made available as preliminary, non-peer-reviewed reports.8-11 Some have shown benefits of colchicine use, including less need for supplemental oxygen, improvements in clinical status on an ordinal clinical scale, and reductions in certain inflammatory markers. In addition, some studies have reported higher discharge rates or fewer deaths among patients who received colchicine than among those who received comparator drugs or placebo. However, the findings of these studies are difficult to interpret due to significant design or methodological limitations, including small sample sizes, open-label designs, and differences in the clinical and demographic characteristics of participants and permitted use of various cotreatments (e.g., remdesivir, corticosteroids) in the treatment arms.

Adverse Effects, Monitoring, and Drug-Drug Interactions

Common adverse effects of colchicine include diarrhea, nausea, vomiting, abdominal cramping and pain, bloating, and loss of appetite. In rare cases, colchicine is associated with serious adverse events, such as neuromyotoxicity and blood dyscrasias. Use of colchicine should be avoided in patients with severe renal insufficiency, and patients with moderate renal insufficiency who receive the drug should be monitored for adverse effects. Caution should be used when colchicine is coadministered with drugs that inhibit cytochrome P450 (CYP) 3A4 and/or P-glycoprotein (P-gp) because such use may increase the risk of colchicine-induced adverse effects due to significant increases in colchicine plasma levels. The risk of myopathy may be increased with the concomitant use of certain HMG-CoA reductase inhibitors (e.g., atorvastatin, lovastatin, simvastatin) due to potential competitive interactions mediated by CYP3A4 and P-gp pathways.12,13 Fatal colchicine toxicity has been reported in individuals with renal or hepatic impairment who received colchicine in conjunction with P-gp inhibitors or strong CYP3A4 inhibitors.

Considerations in Pregnancy

There are limited data on the use of colchicine in pregnancy. Fetal risk cannot be ruled out based on data from animal studies and the drug’s mechanism of action. Colchicine crosses the placenta and has antimitotic properties, which raises a theoretical concern for teratogenicity. However, a recent meta-analysis did not find that colchicine exposure during pregnancy increased the rates of miscarriage or major fetal malformations. There are no data for colchicine use in pregnant women with acute COVID-19. Risks of use should be balanced against potential benefits.12,14

Considerations in Children

Colchicine is most commonly used in children to treat periodic fever syndromes and autoinflammatory conditions. Although colchicine is generally considered safe and well tolerated in children, there are no data on the use of the drug to treat pediatric acute COVID-19 or multisystem inflammatory syndrome in children (MIS-C).

References

  1. van Echteld I, Wechalekar MD, Schlesinger N, Buchbinder R, Aletaha D. Colchicine for acute gout. Cochrane Database Syst Rev. 2014(8):CD006190. Available at: https://www.ncbi.nlm.nih.gov/pubmed/25123076.
  2. Xia M, Yang X, Qian C. Meta-analysis evaluating the utility of colchicine in secondary prevention of coronary artery disease. Am J Cardiol. 2021;140:33-38. Available at: https://www.ncbi.nlm.nih.gov/pubmed/33137319.
  3. Reyes AZ, Hu KA, Teperman J, et al. Anti-inflammatory therapy for COVID-19 infection: the case for colchicine. Ann Rheum Dis. 2021 May;80(5):550-557. Available at: https://www.ncbi.nlm.nih.gov/pubmed/33293273.
  4. Tardif JC, Bouabdallaoui N, L’Allier PL, et al. Colchicine for community-treated patients with COVID-19 (COLCORONA): a phase 3, randomised, double-blinded, adaptive, placebo-controlled, multicentre trial. Lancet Respir Med. 2021;9(8):924-932. Available at: https://www.ncbi.nlm.nih.gov/pubmed/34051877.
  5. PRINCIPLE Trial Collaborative Group, Dorward J, Yu L, et al. Colchicine for COVID-19 in adults in the community (PRINCIPLE): a randomised, controlled, adaptive platform trial. medRxiv. 2021;Preprint. Available at: https://www.medrxiv.org/content/10.1101/2021.09.20.21263828v1.
  6. RECOVERY Collaborative Group. Colchicine in patients admitted to hospital with COVID-19 (RECOVERY): a randomised, controlled, open-label, platform trial. Lancet Respir Med. 2021;Published online ahead of print. Available at: https://www.ncbi.nlm.nih.gov/pubmed/34672950.
  7. Deftereos SG, Giannopoulos G, Vrachatis DA, et al. Effect of colchicine vs standard care on cardiac and inflammatory biomarkers and clinical outcomes in patients hospitalized with coronavirus disease 2019: the GRECCO-19 randomized clinical trial. JAMA Netw Open. 2020;3(6):e2013136. Available at: https://www.ncbi.nlm.nih.gov/pubmed/32579195.
  8. Brunetti L, Diawara O, Tsai A, et al. Colchicine to weather the cytokine storm in hospitalized patients with COVID-19. J Clin Med. 2020;9(9). Available at: https://www.ncbi.nlm.nih.gov/pubmed/32937800.
  9. Sandhu T, Tieng A, Chilimuri S, Franchin G. A case control study to evaluate the impact of colchicine on patients admitted to the hospital with moderate to severe COVID-19 infection. Can J Infect Dis Med Microbiol. 2020. Available at: https://www.ncbi.nlm.nih.gov/pubmed/33133323.
  10. Lopes MI, Bonjorno LP, Giannini MC, et al. Beneficial effects of colchicine for moderate to severe COVID-19: a randomised, double-blinded, placebo-controlled clinical trial. RMD Open. 2021;7(1). Available at: https://www.ncbi.nlm.nih.gov/pubmed/33542047.
  11. Salehzadeh F, Pourfarzi F, Ataei S. The impact of colchicine on the COVID-19 patients; a clinical trial. Research Square. 2020;Preprint. Available at: https://www.researchsquare.com/article/rs-69374/v1.
  12. Colchicine (Colcrys) [package insert]. Food and Drug Administration. 2012. Available at: https://www.accessdata.fda.gov/drugsatfda_docs/label/2014/022352s017lbl.pdf.
  13. American College of Cardiology. AHA statement on drug-drug interactions with statins. 2016. Available at: https://www.acc.org/latest-in-cardiology/ten-points-to-remember/2016/10/20/21/53/recommendations-for-management-of-clinically-significant-drug. Accessed November 2, 2021.
  14. Indraratna PL, Virk S, Gurram D, Day RO. Use of colchicine in pregnancy: a systematic review and meta-analysis. Rheumatology (Oxford). 2018;57(2):382-387. Available at: https://www.ncbi.nlm.nih.gov/pubmed/29029311.

www.covid19treatmentguidelines.nih.govAn official website of the National Institutes of Health

Heart-disease risk soars after COVID — even with a mild case

Authors: Saima May Sidik 10 February 2022

Nature

Massive study shows a long-term, substantial rise in risk of cardiovascular disease, including heart attack and stroke, after a SARS-CoV-2 infection.

Even a mild case of COVID-19 can increase a person’s risk of cardiovascular problems for at least a year after diagnosis, a new study1 shows. Researchers found that rates of many conditions, such as heart failure and stroke, were substantially higher in people who had recovered from COVID-19 than in similar people who hadn’t had the disease.

What’s more, the risk was elevated even for those who were under 65 years of age and lacked risk factors, such as obesity or diabetes.

“It doesn’t matter if you are young or old, it doesn’t matter if you smoked, or you didn’t,” says study co-author Ziyad Al-Aly at Washington University in St. Louis, Missouri, and the chief of research and development for the Veterans Affairs (VA) St. Louis Health Care System. “The risk was there.”

Al-Aly and his colleagues based their research on an extensive health-record database curated by the United States Department of Veterans Affairs. The researchers compared more than 150,000 veterans who survived for at least 30 days after contracting COVID-19 with two groups of uninfected people: a group of more than five million people who used the VA medical system during the pandemic, and a similarly sized group that used the system in 2017, before SARS-CoV-2 was circulating.

Troubled hearts

People who had recovered from COVID-19 showed stark increases in 20 cardiovascular problems over the year after infection. For example, they were 52% more likely to have had a stroke than the contemporary control group, meaning that, out of every 1,000 people studied, there were around 4 more people in the COVID-19 group than in the control group who experienced stroke.

The risk of heart failure increased by 72%, or around 12 more people in the COVID-19 group per 1,000 studied. Hospitalization increased the likelihood of future cardiovascular complications, but even people who avoided hospitalization were at higher risk for many conditions.

“I am actually surprised by these findings that cardiovascular complications of COVID can last so long,” Hossein Ardehali, a cardiologist at Northwestern University in Chicago, Illinois, wrote in an e-mail to Nature. Because severe disease increased the risk of complications much more than mild disease, Ardehali wrote, “it is important that those who are not vaccinated get their vaccine immediately”.COVID’s cardiac connection

Ardehali cautions that the study’s observational nature comes with some limitations. For example, people in the contemporary control group weren’t tested for COVID-19, so it’s possible that some of them actually had mild infections. And because the authors considered only VA patients — a group that’s predominantly white and male — their results might not translate to all populations.

Ardehali and Al-Aly agree that health-care providers around the world should be prepared to address an increase in cardiovascular conditions. But with high COVID-19 case counts still straining medical resources, Al-Aly worries that health authorities will delay preparing for the pandemic’s aftermath for too long. “We collectively dropped the ball on COVID,” he said. “And I feel we’re about to drop the ball on long COVID.”

doi: https://doi.org/10.1038/d41586-022-00403-0

References

  1. Xie, Y., Xu, E., Bowe, B. & Al-Aly, Z. Nature Med. https://www.nature.com/articles/s41591-022-01689-3 (2022).PubMed Article Google Scholar 

Former Senior FDA Official: Manufacturers, FDA Negligent In Not Investigating Covid-19 Vaccine Risks To Heart Health

Authors: DAVID GORTLER FEBRUARY 10, 2022

Manufacturers, FDA, and CDC must investigate serious cardiovascular incidents related to the Pfizer and Moderna Covid vaccines.

From day ,one the U.S. Food and Drug Administration knew the Covid-19 vaccine was linked to serious heart trouble in recipients. The FDA medical officer review of Pfizer’s original Covid-19 application notes “clinically important serious adverse reactions [included] anaphylaxis and myocarditis/pericarditis”—that is, severe allergic reactions and inflammation of the heart and or the sac containing the heart, respectively. As of this writing, FDA has not released its review of the Moderna “Spikevax” mRNA vaccine application despite having granted emergency use authorization well more than a year ago and full approval late last month.

The Vaccine Adverse Event Reporting System (VAERS), jointly run by FDA and the Centers for Disease Control, lists a long and impersonal number of cardiovascular-related events in young, healthy people. Without reading the underlying narratives submitted with the reports, it’s hard to establish the precise causal links regarding these adverse events. Still, there are thousands of reports of heart attacks, myocarditis, and pericarditis in the United States alone, which should have spurred manufacturers and the FDA into full investigation mode.

Studies acknowledged by FDA officials show that the FDA’s various safety databases only collect an estimated 1 to 13 percent of all adverse events that occur. Multiple FDA drug safety epidemiologists have stated during official FDA presentation that it only takes a single well-documented adverse event to justify a safety signal investigation and in turn to warn the American public of the potential risk.

Historically, the FDA has sought safety warnings on labels, up to and including a “black boxed warning” and a prescribing restriction known as a Risk Evaluation and Mitigation Strategy (REMS) for much less. For instance, in 2008, after fewer than 200 spontaneous VAERS reports of tendon rupture following administration of the class of antibiotics known as fluoroquinolones, FDA added a “black box warning” and REMS prescribing restrictions.

Yet thousands of serious, debilitating, and deadly safety VAERS reports following Covid vaccines and boosters are not being held to the same regulatory standards. If approximately 1 to 13 percent of adverse events are reported, extrapolating those numbers means the actual number of adverse health events could easily be in the hundreds of thousands in the United States and many millions worldwide.

In addition to VAERS, the CDC’s Vaccine Safety Datalink indicates an excess risk of myocarditis and pericarditis in recipients following the Pfizer and Moderna vaccines. The cardiovascular risk after any mRNA vaccine is high, but with Moderna it’s approximately four times higher than Pfizer’s.

Other public health agencies with much tinier budgets and staff compared to our FDA’s took action on this months ago. In October, DenmarkFinlandNorway, and Sweden suspended the use of the Moderna vaccine for young people, but it’s still full speed ahead here in the United States.

Since then, more data has been released affirming the same: On Jan. 25, 2022, a CDC and FDA study published in JAMA shows the risk of myocarditis following any kind of mRNA Covid vaccination is greater than the background risk in the population, with the largest proportions of cases of myocarditis occurring among white males.

A comprehensive study out of Britain from December 2021 examined data from more than 42 million people who have taken a Covid-19 shot found a noteworthy increase in myocarditis with mRNA vaccines that persisted and increased with every dose and booster. “An association between Covid-19 infection and myocarditis was observed in all ages for both sexes,” the study’s abstract states. “These findings have important implications for public health and vaccination policy.” Indeed they do—especially in light of the questionable way the FDA approved vaccines in kids from 5 to 13 years old, and the pending FDA applications to approve vaccination in babies starting at 6 months old.

The FDA, CDC, and manufacturers have access to VAERS and additional high-quality denominator-based vaccine safety systems including the Biologics Effectiveness and Safety Initiative (BEST) and the Vaccine Safety Datalink (VSD), respectively. Have manufacturers and our health agencies used these tools and others to fully investigate the cardiovascular health risks of the vaccine? There is reason to doubt, given the political pressure the Biden administration has put on the agencies to advocate for taking the vaccine while almost never mentioning safety.

Myocarditis and pericarditis have historically been rare. They are defined as inflammation of the heart muscle or layers of the pericardial sac, respectively. Both conditions cause easily recognizable ECG changes and have ambiguous symptoms that include shortness of breath and chest pain. Myocarditis and pericarditis can easily be diagnosed clinically with echocardiograms and can be treated by inexpensive pharmacology and bedrest, but for that to happen, people need to know to seek medical diagnosis and care. 

Therein is the problem: providers and patients are not being adequately warned to monitor for cardiovascular symptoms despite the increased incidence. Since there is a failure of manufacturers and the FDA to address this and other untoward effects of mRNA utility and mandates, outside drug safety experts need to publicly address mRNA Covid vaccine safety immediately.

On February 4, 2022, a CDC advisory committee proposed extending the gap between Covid-19 shots to mitigate the cardiovascular damage of the vaccine. This indicates the federal government is aware of the serious risk. Yet rather than addressing the risk head-on by communicating the facts to the public, they seem to be taking a “half measure” of changing the interval and hoping to mitigate risk without evidence it will have any effect on outcome.

In the very recent past, anyone warning about the exact same cardiovascular risk that this advisory panel spoke about less than a week ago were shamed and banned on social media by “big tech” “fact checkers.”

Vaccines are one of the most important inventions in human history, having saved millions of lives. That does not mean every person should get every vaccine. Also, like every drug out there, it is critically important to quickly detect and report safety problems. Now we have a federally mandated vaccine that is clearly no longer effective, and potentially causing additional illness and death.

The failure to adequately monitor and warn for Covid vaccine adverse events has served to harden not only Covid vaccine hesitancy but has shredded the credibility of public health authorities. The failure to openly talk about known adverse reactions erodes trust.

In the 1950s physicians used to not tell patients when they had terminal cancer because they thought it was for their own good. We are long past the day when hiding information from the public is considered good for public health. It never is. It is not only unethical and insulting, it’s dangerous.

To See 1,000 Peer Reviewed Articles on COVID-19 Adverse Events Click Below

http://wp.cov19longhaulfoundation.org/?s=1000

Multiple Early Factors Anticipate Post-Acute COVID-19 Sequelae

Highlights

  • Longitudinal multiomics associate PASC with autoantibodies, viremia and comorbidities
  • Reactivation of latent viruses during initial infection may contribute to PASC
  • Subclinical autoantibodies negatively correlate with anti-SARS-CoV-2 antibodies
  • Gastrointestinal PASC uniquely present with post-acute expansion of cytotoxic T cells

SUMMARY

Post-acute sequelae of COVID-19 (PASC) represent an emerging global crisis. However, quantifiable risk-factors for PASC and their biological associations are poorly resolved. We executed a deep multi-omic, longitudinal investigation of 309 COVID-19 patients from initial diagnosis to convalescence (2-3 months later), integrated with clinical data, and patient-reported symptoms. We resolved four PASC-anticipating risk factors at the time of initial COVID-19 diagnosis: type 2 diabetes, SARS-CoV-2 RNAemia, Epstein-Barr virus viremia, and specific autoantibodies. In patients with gastrointestinal PASC, SARS-CoV-2-specific and CMV-specific CD8+ T cells exhibited unique dynamics during recovery from COVID-19. Analysis of symptom-associated immunological signatures revealed coordinated immunity polarization into four endotypes exhibiting divergent acute severity and PASC. We find that immunological associations between PASC factors diminish over time leading to distinct convalescent immune states. Detectability of most PASC factors at COVID-19 diagnosis emphasizes the importance of early disease measurements for understanding emergent chronic conditions and suggests PASC treatment strategies.

Article Info

Publication History

Accepted: January 19, 2022Received in revised form: December 14, 2021Received: September 29, 2021

Click on Link Below for Complete Study:

https://www.cell.com/cell/pdf/S0092-8674(22)00072-1.pdf?_returnURL=https%3A%2F%2Flinkinghub.elsevier.com%2Fretrieve%2Fpii%2FS0092867422000721%3Fshowall%3Dtrue

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

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

Background

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

Methods

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

Findings

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

Interpretation

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

Funding

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

Keywords

 Evidence before this study

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

 Added value of this study

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

 Implications of all the available evidence

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

Introduction

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

Methods

 Human samples

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

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

 Ethics

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

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

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

 Histology study

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

 Immunohistochemistry staining

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

 Cell isolation from autopsy tissue samples

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

 Flow cytometry

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

 Real-time RT-PCR

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

 SARS-CoV-2 RNA detection

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

 Electron microscopy

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

 Mice

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

 Statistics

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

 Role of the funding source

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

Results

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

Discussion

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

Contributors

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

Declaration of interests

None exist.

Acknowledgments

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

Data sharing statement

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

Appendix. Supplementary materials

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

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

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Figures

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

13 ways that the SARS-CoV-2 spike protein causes damage

Authors: Posted on January 13, 2022 by Jesse Santiano, M.D.

The SARS-CoV-2 virus has four structural proteins. The spike, membrane, envelop, and nucleocapsid proteins. The spike protein protrudes from the middle of the coronavirus and attaches to the ACE2 receptor of cells to start the process of cell entry, replication, and infection. The two major parts of the spike protein are the S1 and S2 subunit. The S1 has the receptor-binding domain.

For easier reading, this review starts with what happens after the COVID jabs, soluble spike proteins, and what it takes to have a normal blood vessel. Then I will enumerate how the spike protein damages the body.

What happens after the COVID shots?

COVID vaccination aims to produce an immune response against the spike protein in the form of neutralizing antibodies so that in future SARS-CoV-2 exposures, COVID-19 will be prevented.

The injected messenger RNA provides instructions to the cells on making the spike proteins. Once the spike protein is produced, it migrates to the outside of the cell to be anchored on the cells’ outer surface, where the immune system will recognize it and develop an immune response to it. (antibodies, T cells, B cells).

Soluble spike proteins

Ideally, the whole spike protein should stay attached to the outside of the cells. Sometimes incomplete spike proteins are produced in the form of spike peptides. As shown below, they are also presented to the immune system by cells outside the surface with an anchoring protein called the Major Histocompatibility Complex (MHC).

Anchoring to the cells is critical because once the spike protein or its pieces in the form of peptides become soluble or float in the bloodstream, they induce inflammation and clot formation in the arteries and capillaries. Scientists have found several ways that it happens.

First is that enzymes called metalloproteinases can cut the MHC1 at their bases.[1] Free-floating MHCs are found in patients with systemic lupus erythematosus SLE and cancers.[4][5]

The second is that errors can happen while RNA splicing occurs inside the nucleus. This results in variant spike proteins that are soluble.[2] Soluble S1 subunits were observed among recipients of the Moderna shots.[3

You can read more about it in this article: SARS-CoV-2 spike proteins detected in the plasma following Moderna shots.

Third, are exosomes released from cells containing MHCs with the spike proteins. [6] T-cells can interact with the spike proteins in the exosomes and cause inflammation [7]. Immunogenic spike proteins inside exosomes were demonstrated after Pfizer injection[8].

 Donor Blood Can Have Spike Protein Exosomes

The normal blood vessel

All organs in the body need an adequate blood supply, and blood vessels have to be in pristine working conditions for that to happen. They should be distensible to allow greater blood flow during exertion, smooth inside to prevent blood clot formation, and have working mechanisms to repair themselves and dissolve blood clots that may form.

All that work falls on the endothelial cells that line the inner wall of the blood vessels, andI talked about them at The Magical Endothelium. Any injury to the endothelium can elicit an inflammatory response and clot formation leading to organ dysfunctions like heart attacks, strokes, and deaths. 

Blood clots always start small, and once they develop, they initiate a chain reaction that promotes a more extensive clot. The good thing is that the body can do fibrinolysis, a built-in mechanism to dissolve clots.

13 ways Spike Proteins cause disease

The following are how the spike proteins and their S1 subunit can cause damage. They can work together and have four results, inflammationthrombosis or clot formation, auto-immunity, and amyloid formation.

Any foreign protein inside the body can elicit inflammation. That is why parts of the spike proteins in the form of their S1 subunits or shorter fragments are enough to cause damage.[9][10].

Inflammation and thrombosis

The S1 subunit activates Toll-like receptor 4 (TLR4) signaling to induce pro-inflammatory responses. It happens with the spike protein in COVID-19 [10] and the S1 subunits.[12]

The spike protein triggers cell signaling events that promote pulmonary vascular remodeling and pulmonary arterial hypertension (PAH), and other cardiovascular complications [13]Source: Suzuki and Gychka. Vaccines 2

The spike S1 initiates inflammatory responses from tumor-necrosis factor-alpha (TNF-α) and interleukin-6 (IL-6) to initiate a cytokine storm syndrome in the lungs. [14].

Spike proteins cause vascular leaks by degrading the barrier of the endothelium. [15] The leak may explain the proliferation of lymphocytes seen by German pathologists in the organs of deceased patients who died after the shots.

Spike proteins affect the cardiac pericytes, the cells that “supervise” the endothelial cells responsible for maintaining the smoothness of the blood vessels. [16]. Study shows spike proteins affect cardiac pericytes and explain why soccer players collapse

Spike proteins downregulate the ACE2 and impair endothelial function [17]

The S1 produces blood clots resistant to the body’s fibrinolysis and hospital clot-buster medications [18]. That’s why some have their limbs amputated after the shots. One woman had both legs and hands amputated. There’s a list here.

The spike protein can cause inflammation by activating the alternate complement pathway. [20]

Long COVID-Syndrome

  1. The S1 Proteins can persist in CD16+ Monocytes up to 15 Months Post-Infection and vaccination to induce chronic inflammation. This explains the symptoms of the Long COVID Syndrome. [19]

Amyloids formation and interaction

  1. Amyloids are fibrillar proteins. They are most commonly associated with neurodegenerative diseases like dementia. However, they can also form in the heart and lungs and make them rigid and form blood clots resistant to dissolution. [21The SARS-CoV-2 spike protein can form amyloids seen in lung, blood, and nervous system disorders
  2. The S1 protein contains heparin-binding sites that attract amyloids to initiate amyloid protein aggregation. Amyloid formation leads to neurodegeneration like Parkinson’s Disease, Alzheimer’s’ disease, and Frontal lobe dementia. [22]

Molecular mimicry

12. Molecular mimicry happens if protein sequences in the spike protein and peptides have similarities to human proteins. Antibodies made for those viral proteins may also attack the host proteins.[24][25][26].

This leads to several autoimmune diseases like immune thrombocytopenia (low platelet counts) [23], autoimmune liver diseasesGuillain-Barré syndromeIgA nephropathyrheumatoid arthritis, and systemic lupus erythematosus[27]

Cancer and Immune Deficiency

  1. Spike proteins impair DNA damage repair and result in ineffective antibodies and damaged tumor-suppressor genes like the BRCA1 and 53BP1 that lead to cancers. BRCA1 damage is associated with breast, ovarian, and prostate cancers.

53BP1 loss of function in tumor tissues is elated to tumor occurrence, progression, and poor prognosis in human malignancies.[30]

Parting thoughts

The disease-causing part of the SARS-CoV-2 virus is the spike protein, and it is present in COVID-19 and the COVID injections. Prevention and early treatment are possible for COVID-19. Once you have the shot, there is no way to control the spike protein.

It is unclear why not all have the adverse effects or die. What is sure is that there are over one million reported adverse effects on VAERS, and more than one hundred thousand have been killed. Vaccine-induced deaths in the U.S. and Europe are way higher than the VAERS reports!

This list is not all-inclusive, and I probably missed some. Indeed, more will be discovered in the future, and I don’t want my body to find out. Do you?

Don’t Get Sick!

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Related:

  1. Blood Vessel Damaging Proteins of the SARS-CoV-2
  2. Cerebral Thrombosis after the Pfizer Covid-19 Vaccine
  3. The High Risk of Deadly Brain Clots in the J & J COVID Vaccine
  4. This Study shows a Ten-Fold Risk of Developing Blood Clots after the COVID Vaccines.
  5. You got the COVID shot and found that others developed blood clots. Now what?
  6. Platelet Changes Causes Blood Clots in COVID-19
  7. Unidentified Foreign Bodies in the Vaccines Form Clots
  8. Retinal complications after COVID shots
  9. U.K. Study of COVID-19 shots and Excess Rates of Guillain-Barré Syndrome
  10. mRNA Vaccination Increases the Risk of Acute Coronary Syndrome
  11. German Analysis: The Higher the Vaccination Rate, the Higher the Excess Mortality
  12. Anti-Idiotype Antibodies against the Spike Proteins may Explain Myocarditis

References:

  1. Rijkers GT, Weterings N, Obregon-Henao A, et al. Antigen Presentation of mRNA-Based and Virus-Vectored SARS-CoV-2 VaccinesVaccines (Basel). 2021;9(8):848. Published 2021 Aug 3. doi:10.3390/vaccines9080848
  2. Kowarz E, Krutzke L, Reis J, et al. “Vaccine-Induced Covid-19 Mimicry” Syndrome: Splice reactions within the SARS-CoV-2 Spike open reading frame result in Spike protein variants that may cause thromboembolic events in patients immunized with vector-based vaccines. Research Square; 2021. DOI: 10.21203/rs.3.rs-558954/v1
  3. Ogata AF. et al. Circulating SARS-CoV-2 Vaccine Antigen Detected in the Plasma of mRNA-1273 Vaccine Recipients [published online ahead of print, 2021 May 20]. Clin Infect Dis. 2021;ciab465. doi:10.1093/cid/ciab465
  4. Hervier B, Ribon M, Tarantino N, Mussard J, Breckler M, Vieillard V, Amoura Z, Steinle A, Klein R, Kötter I, Decker P. Increased Concentrations of Circulating Soluble MHC Class I-Related Chain A (sMICA) and sMICB and Modulation of Plasma Membrane MICA Expression: Potential Mechanisms and Correlation With Natural Killer Cell Activity in Systemic Lupus Erythematosus. Front Immunol. 2021 May 3;12:633658. doi: 10.3389/fimmu.2021.633658. PMID: 34012432; PMCID: PMC8126610.
  5. Salih, Helmut & Goehlsdorf, Dennis & Steinle, Alexander. (2006). Salih HR, Goehlsdorf D, Steinle A. Release of MICB molecules by tumor cells: mechanism and soluble MICB in sera of cancer patients. Hum Immunol 67: 188-195. Human immunology. 67. 188-95. 10.1016/j.humimm.2006.02.008.
  6. Edgar JR. Q&A: What are exosomes, exactly?BMC Biol. 2016;14:46. Published 2016 Jun 13. doi:10.1186/s12915-016-0268-z
  7. Raposo G, Nijman HW, Stoorvogel W, Liejendekker R, Harding CV, Melief CJ, Geuze HJ. B lymphocytes secrete antigen-presenting vesicles. J Exp Med. 1996 Mar 1;183(3):1161-72. doi: 10.1084/jem.183.3.1161. PMID: 8642258; PMCID: PMC2192324.
  8. Bansal et al. Cutting Edge: Circulating Exosomes with COVID Spike Protein Are Induced by BNT162b2 (Pfizer–BioNTech) Vaccination prior to Development of Antibodies: A Novel Mechanism for Immune Activation by mRNA Vaccines. J Immunol November 15, 2021, 207 (10) 2405-2410
  9. Nuovo, G.J. et al. (2021) Endothelial cell damage is the central part of COVID-19 and a mouse model induced by injection of the S1 subunit of the spike protein. Ann. Diagn. Pathol. 51, 151682, https://doi.org/10.1016/j.anndiagpath.2020.151682
  10. Gu, T. et al. (2020) Cytokine signature induced by SARS-CoV-2 spike protein in a mouse model. Front. Immunol. 11, 621441,
  11. Aboudounya MM, Heads RJ. COVID-19 and Toll-Like Receptor 4 (TLR4): SARS-CoV-2 May Bind and Activate TLR4 to Increase ACE2 Expression, Facilitating Entry and Causing Hyperinflammation. Mediators Inflamm. 2021 Jan 14;2021:8874339. doi: 10.1155/2021/8874339. PMID: 33505220; PMCID: PMC7811571.
  12. Shirato K, Kizaki T. SARS-CoV-2 spike protein S1 subunit induces pro-inflammatory responses via toll-like receptor 4 signaling in murine and human macrophages. Heliyon. 2021 Feb 2;7(2):e06187. doi: 10.1016/j.heliyon.2021.e06187. PMID: 33644468; PMCID: PMC7887388. https://pubmed.ncbi.nlm.nih.gov/33644468/
  13. Suzuki YJ, et al. SARS-CoV-2 Spike Protein Elicits Cell Signaling in Human Host Cells: Implications for Possible Consequences of COVID-19 VaccinesVaccines (Basel). 2021;9(1):36. Published 2021 Jan 11. doi:10.3390/vaccines9010036
  14. Cao, X. et al. (2021) Spike protein of SARS-CoV-2 activates macrophages and contributes to induction of acute lung inflammation in male mice. FASEB J. 35, e21801
  15. Biering et al. SARS-CoV-2 Spike triggers barrier dysfunction and vascular leak via integrins and TGF-β signaling. bioRxiv 2021.12.10.472112
  16. Avolio E et al.  The SARS-CoV-2 Spike protein disrupts human cardiac pericytes function through CD147 receptor-mediated signaling: a potential non-infective mechanism of COVID-19 microvascular disease. Clin Sci (Lond). 2021 Dec 22;135(24):2667-2689. doi: 10.1042/CS20210735. PMID: 34807265; PMCID: PMC8674568.
  17. Lei, Y. et al. SARS-CoV-2 Spike Protein Impairs Endothelial Function via Downregulation of ACE 2. Circulation Research. 2021;128:1323–1326
  18. Grobbelaar, L.M. et al. (2021) SARS-CoV-2 spike protein S1 induces fibrin(ogen) resistant to fibrinolysis: implications for microclot formation in COVID-19. Biosci. Rep. 41 (8)
  19. Patterson B. et al. Persistence of SARS CoV-2 S1 Protein in CD16+ Monocytes in Post-Acute Sequelae of COVID-19 (PASC) Up to 15 Months Post-Infection.bioRxiv 2021.06.25.449905
  20. Yu J, Yuan X, Chen H, Chaturvedi S, Braunstein EM, Brodsky RA. Direct activation of the alternative complement pathway by SARS-CoV-2 spike proteins is blocked by factor D inhibition. Blood. 2020.
  21. Nyström et al. Amyloidogenesis of SARS-CoV-2 Spike Protein. bioRxiv 2021.12.16.472920. 
  22. Idrees D, Kumar V. SARS-CoV-2 spike protein interactions with amyloidogenic proteins: Potential clues to neurodegeneration. Biochem Biophys Res Commun. 2021 May 21;554:94-98. doi: 10.1016/j.bbrc.2021.03.100. Epub 2021 Mar 24. PMID: 33789211; PMCID: PMC7988450
  23. Nunez-Castilla, J et al. Spike mimicry of thrombopoietin may induce thrombocytopenia in COVID-19. bioRxiv 2021.08.10.455737
  24. Ehrenfeld M et al. Covid-19 and autoimmunity. Autoimmun Rev. 2020;19:102597.
  25. Kanduc D, Shoenfeld Y. On the molecular determinants of the SARS-CoV-2 attack. Clin Immunol. 2020;215. 
  26. Vojdani A, Kharrazian D. Potential antigenic cross-reactivity between SARS-CoV-2 and human tissue with a possible link to an increase in autoimmune diseases. Clin Immunol. 2020;217:108480
  27. Chen Y, Xu Z, Wang P, Li XM, Shuai ZW, Ye DQ, Pan HF. New-onset autoimmune phenomena post-COVID-19 vaccination. Immunology. 2021 Dec 27. doi: 10.1111/imm.13443. Epub ahead of print. PMID: 34957554.
  28. Jiang H, Mei YF. SARS-CoV-2 Spike Impairs DNA Damage Repair and Inhibits V(D)J Recombination In VitroViruses. 2021;13(10):2056. Published 2021 Oct 13. doi:10.3390/v13102056
  29. BRCA1 gene. Medline [website]
  30. Mirza-Aghazadeh-Attari M, et al.  53BP1: A key player of DNA damage response with critical functions in cancer. DNA Repair (Amst). 2019 Jan;73:110-119. doi: 10.1016/j.dnarep.2018.11.008. Epub 2018 Nov 20. PMID: 30497961.

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Abstract 10712: Mrna COVID Vaccines Dramatically Increase Endothelial Inflammatory Markers and ACS Risk as Measured by the PULS Cardiac Test: a Warning

Authors: Steven R Gundry Originally published 8 Nov 2021Circulation. 2021;144:A10712

Abstract

Our group has been using the PLUS Cardiac Test (GD Biosciences, Inc, Irvine, CA) a clinically validated measurement of multiple protein biomarkers which generates a score predicting the 5 yr risk (percentage chance) of a new Acute Coronary Syndrome (ACS). The score is based on changes from the norm of multiple protein biomarkers including IL-16, a proinflammatory cytokine, soluble Fas, an inducer of apoptosis, and Hepatocyte Growth Factor (HGF)which serves as a marker for chemotaxis of T-cells into epithelium and cardiac tissue, among other markers. Elevation above the norm increases the PULS score, while decreases below the norm lowers the PULS score.The score has been measured every 3-6 months in our patient population for 8 years. Recently, with the advent of the mRNA COVID 19 vaccines (vac) by Moderna and Pfizer, dramatic changes in the PULS score became apparent in most patients.This report summarizes those results. A total of 566 pts, aged 28 to 97, M:F ratio 1:1 seen in a preventive cardiology practice had a new PULS test drawn from 2 to 10 weeks following the 2nd COVID shot and was compared to the previous PULS score drawn 3 to 5 months previously pre- shot. Baseline IL-16 increased from 35=/-20 above the norm to 82 =/- 75 above the norm post-vac; sFas increased from 22+/- 15 above the norm to 46=/-24 above the norm post-vac; HGF increased from 42+/-12 above the norm to 86+/-31 above the norm post-vac. These changes resulted in an increase of the PULS score from 11% 5 yr ACS risk to 25% 5 yr ACS risk. At the time of this report, these changes persist for at least 2.5 months post second dose of vac.We conclude that the mRNA vacs dramatically increase inflammation on the endothelium and T cell infiltration of cardiac muscle and may account for the observations of increased thrombosis, cardiomyopathy, and other vascular events following vaccination.

Footnotes

Author Disclosures: For author disclosure information, please visit the AHA Scientific Sessions 2021 Online Program Planner and search for the abstract title.

Study Finds Teenage Boys Six Times More Likely To Suffer Heart Problems From Vaccine Than Be Hospitalized by COVID

Authors; Paul Joseph Watson via Summit News,

Research conducted by the University of California has found that teenage boys are six times more likely to suffer from heart problems caused by the COVID-19 vaccine than to be hospitalized as a result of COVID-19 itself.

“A team led by Dr Tracy Hoeg at the University of California investigated the rate of cardiac myocarditis – heart inflammation – and chest pain in children aged 12-17 following their second dose of the vaccine,” reports the Telegraph.

“They then compared this with the likelihood of children needing hospital treatment owing to Covid-19, at times of low, moderate and high rates of hospitalisation.”

Researchers found that the risk of heart complications for boys aged 12-15 following the vaccine was 162.2 per million, which was the highest out of all the groups they looked at.

This compares to the risk of a healthy boy being hospitalized as a result of a COVID infection, which is around 26.7 per million, meaning the risk they face from the vaccine is 6.1 times higher.

Even during high risk rates of COVID, such as in January this year, the threat posed by the vaccine is 4.3 times higher, while during low risk rates, the risk of teenage boys suffering a “cardiac adverse event” from the vaccine is a whopping 22.8 times higher.

The research data was based on a study of adverse reactions suffered by teens between January and June this year.

In a sane world, such data should represent the nail in the coffin for the argument that teenagers and children should be mandated to take the coronavirus vaccine, but it obviously won’t.

In the UK, the government is pushing to vaccinate 12-15-year-olds, even without parental consent, despite the Joint Committee on Vaccination and Immunisation (JCVI) advising against it.

Meanwhile, in America, Los Angeles County school officials voted unanimously to mandate COVID shots for all

Long covid—mechanisms, risk factors, and management

Authors: Harry Crook, research assistant1,  Sanara Raza, research assistant1,  Joseph Nowell, research assistant1,  Megan Young, clinical research officer1,  Paul Edison, clinical senior lecturer, honorary professor12

Abstract

Since its emergence in Wuhan, China, covid-19 has spread and had a profound effect on the lives and health of people around the globe. As of 4 July 2021, more than 183 million confirmed cases of covid-19 had been recorded worldwide, and 3.97 million deaths. Recent evidence has shown that a range of persistent symptoms can remain long after the acute SARS-CoV-2 infection, and this condition is now coined long covid by recognized research institutes. Studies have shown that long covid can affect the whole spectrum of people with covid-19, from those with very mild acute disease to the most severe forms. Like acute covid-19, long covid can involve multiple organs and can affect many systems including, but not limited to, the respiratory, cardiovascular, neurological, gastrointestinal, and musculoskeletal systems. The symptoms of long covid include fatigue, dyspnea, cardiac abnormalities, cognitive impairment, sleep disturbances, symptoms of post-traumatic stress disorder, muscle pain, concentration problems, and headache. This review summarizes studies of the long term effects of covid-19 in hospitalized and non-hospitalized patients and describes the persistent symptoms they endure. Risk factors for acute covid-19 and long covid and possible therapeutic options are also discussed.

Introduction

Coronavirus disease 2019 (covid-19) has spread across the world. As of 4 July 2021, more than 183 million confirmed cases of covid-19 have been recorded worldwide, and more than 3.97 million deaths have been reported by the World Health Organization .1 The clinical spectrum of covid-19 ranges from asymptomatic infection to fatal disease.23 The virus responsible for causing covid-19, severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), enters cells via the angiotensin-converting enzyme 2 (ACE2) receptor.4 Once internalized, the virus undergoes replication and maturation, provoking an inflammatory response that involves the activation and infiltration of immune cells by various cytokines in some patients.5 The ACE2 receptor is present in numerous cell types throughout the human body, including in the oral and nasal mucosa, lungs, heart, gastrointestinal tract, liver, kidneys, spleen, brain, and arterial and venous endothelial cells, highlighting how SARS-CoV-2 can cause damage to multiple organs.67

The impact of covid-19 thus far has been unparalleled, and long term symptoms could have a further devastating effect.8 Recent evidence shows that a range of symptoms can remain after the clearance of the acute infection in many people who have had covid-19, and this condition is known as long covid. The National Institute for Health and Care Excellence (NICE) defines long covid as the symptoms that continue or develop after acute covid-19 infection and which cannot be explained by an alternative diagnosis. This term includes ongoing symptomatic covid-19, from four to 12 weeks post-infection, and post-covid-19 syndrome, beyond 12 weeks post-infection.9 Conversely, The National Institutes of Health (NIH) uses the US Centers for Disease Control and Prevention (CDC) definition of long covid, which describes the condition as sequelae that extend beyond four weeks after initial infection.10 People with long covid exhibit involvement and impairment in the structure and function of multiple organs.11121314 Numerous symptoms of long covid have been reported and attributed to various organs, an overview of which can be seen in fig 1. Long term symptoms following covid-19 have been observed across the spectrum of disease severity. This review examines the long term impact of symptoms reported following covid-19 infection and discusses the current epidemiological understanding of long covid, the risk factors that may predispose a person to develop the condition, and the treatment and management guidelines aimed at treating it.

Multi-organ complications of covid-19 and long covid. The SARS-CoV-2 virus gains entry into the cells of multiple organs via the ACE2 receptor. Once these cells have been invaded, the virus can cause a multitude of damage ultimately leading to numerous persistent symptoms, some of which are outlined here.

For More Information: https://www.bmj.com/content/374/bmj.n1648