Understanding COVID-19 through genome-wide association studies

Authors: Tom H. Karlsen  Nature Genetics volume 54, pages368–369 (2022)Cite this article

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Defining the most appropriate phenotypes in genome-wide association studies of COVID-19 is challenging, and two new publications demonstrate how case-control definitions critically determine outcomes and downstream clinical utility of findings.

Exploring self-reported data from more than 700,000 participants in a direct-to-consumer ancestry genetics company, in this issue of Nature Genetics, Roberts et al. report how several commonly used phenotype definitions in COVID-19 genetics studies converge to represent either susceptibility to infection by the SARS-CoV-2 virus or risk of severe COVID-19 disease1. For pragmatic reasons, early genome-wide association studies (GWAS) in COVID-19 focused on hospitalized cases compared with unscreened and often previously genotyped controls2,3. While allowing for rapid assessments during the first and very challenging wave of the pandemic, such study designs are biased towards the biology of complications in COVID-19. The emphasis on patients with mild or no symptoms, including identification of household COVID-19 exposure as a high-risk measure, allowed the authors to conduct a deep investigation of susceptibility to SARS-CoV-2 infection through comparisons such as exposed individuals who tested positive for COVID-19 versus exposed individuals who tested negative. Not only did these assessments corroborate the controversial ABO locus as a bona fide susceptibility gene for SARS-CoV-2 infection2,4, they also suggested the presence of a hitherto unexplored pool of protective variants.

In a dedicated query of rare variants (minor allele frequency (MAF) < 0.005), also reported in this issue of Nature Genetics, Horowitz et al. identified an association signal between a non-coding X chromosome variant (rs190509934) upstream of angiotensin-converting enzyme 2 (ACE2) and protection against SARS-CoV-2 infection5. The authors go on to substantiate their finding using RNA sequencing – data from liver tissue, showing that the protective allele leads to an almost 40% reduction in ACE2 expression levels in carriers. The association inherently holds considerable plausibility, with the membrane-bound ACE2 serving as the binding site for the SARS-CoV-2 spike glycoprotein, initiating virus cell entry6. Furthermore, Horowitz et al.5 and Roberts et al.1 utilize rich phenotype data to dissect the chromosome 3p21.31 association into a susceptibility signal and a severity signal, which localize to SLC6A20 and LZTFL1, respectively, as also observed by others7SLC6A20 encodes the sodium–imino-acid (proline) transporter 1 (SIT1), which functionally interacts with ACE2 (ref. 8), and the risk allele has been shown to associate with increased expression of SLC6A20 (ref. 2). Along with data suggesting that the receptor-binding domain of the SARS-CoV-2 spike protein preferentially interacts with blood group A9, which is encoded by the risk variant at the ABO locus, genetics of the susceptibility to SARS-CoV-2 infection appear to converge on the cell entry apparatus for the virus.

Critical illness in COVID-19 develops in fewer than 10% of individuals infected with SARS-CoV-2 (ref. 10). Given the window from the first symptoms of COVID-19 to onset of severe disease with respiratory failure (typically about one week)10, prediction of a severe disease course following SARS-CoV-2 infection is of considerable clinical interest as well as from a therapeutic point of view. Reliable risk stratification may guide therapeutic interventions during this lead-in period, characterized by enhanced viral replication. These interventions potentially include antiviral therapies, convalescent plasma, neutralizing monoclonal antibodies or — possibly more important for hospitalized patients — immunomodulating drugs.

Horowitz et al. found that a high genetic risk score (top 10%) based on six established severity variants was associated with a 1.65-fold and 1.75-fold higher risk of severe disease, in individuals with or without the presence of clinical risk factors such as age and diabetes, respectively5. Others have found an odds ratio of 2.0 for the impact of the rs10490770 risk allele at the 3p21.31 locus on the combined end-point of death or severe respiratory failure in an overall COVID-19 patient population11, with almost double the effect size in individuals 60 years or younger (odds ratio of 3.5). These magnitudes are comparable with those associated with clinical risk factors. Findings of lower age in individuals homozygous for the chromosome 3p21.31 risk variant support enhanced utility of genetic risk stratification in the young patient population2.

The execution of GWAS in COVID-19 has been remarkably nimble, due in part to robust collaborative networks set up during past GWAS12, as well as the utilization of previously genotyped study populations such as the UK Biobank, AncestryDNA and 23andme1,3,4,5. The rapid phenotyping undertaken by several biobanks and direct-to-consumer genetics companies during the COVID-19 pandemic is unprecedented, and the resulting publications deserve acknowledgement as a form of ‘population-level testing’ for genetic clues in emerging diseases. The orchestration of projects by the COVID-19 Host Genetics Initiative has also been an important catalyzer of activities13. Figure 1 summarizes published and peer-reviewed GWAS articles on COVID-19. However, even at time of writing, the meta-analysis of the sixth data freeze of the COVID-19 Host Genetics Initiative has been released online, reporting on a total of 23 loci involving in COVID-19 susceptibility (7 loci) and severity (15 loci); adding 10 new loci to the consortium’s own publication only 3 months ago7. The 22-month period that has passed since the publication of the first COVID-19 GWAS2 appears even more impressive in comparison with the 7 years of Crohn’s disease genetics — spanning from the 2001 nucleotide-binding oligomerization domain 2 (NOD2) susceptibility gene discovery to a 2008 meta-analysis14,15 — that it took to achieve the same amount of insight. Further exemplified by the 20-year history of genetics of Crohn’s disease, translational studies of GWAS findings take time, but may reveal new and unexpected aspects of pathophysiology. It is in this context that the rapid unravelling of COVID-19 genetics becomes important. Some of the loci hold immediate biological plausibility (for example, ACE2 and some of the chemokines), whereas the underlying mechanisms of others remain obscure. Following this recent sprint of COVID-19 GWAS to which Horowitz et al.5 and Roberts et al.1 significantly contribute, the subsequent translational ultramarathon of biological studies can begin — and with this a deeper understanding of the pathophysiology of SARS-CoV-2 infection and its complications will emerge. Vaccination has proven the ultimate protection against SARS-CoV-2 infection. The hope is that the biological insights provided by COVID-19 GWAS will facilitate identification and development of novel treatment options of not only hospitalized and critically ill COVID-19 patients, but also treatment modalities that can prevent hospitalization.

figure 1
Fig. 1: Genetic loci from COVID-19 GWAS in peer-reviewed publications to date.

References

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SARS-CoV-2 Infection and the Liver

Authors: Katie Morgan,1,*Kay Samuel,2Martin Vandeputte,1Peter C. Hayes,1 and John N. Plevris1

Pathogens. 2020 Jun; 9(6): 430.Published online 2020 May 30. doi: 10.3390/pathogens9060430 PMCID: PMC7350360PMID: 32486188

Abstract

A novel strain of coronoviridae (SARS-CoV-2) was reported in Wuhan China in December 2019. Initially, infection presented with a broad spectrum of symptoms which typically included muscle aches, fever, dry cough, and shortness of breath. SARS-CoV-2 enters cells via ACE2 receptors which are abundant throughout the respiratory tract. However, there is evidence that these receptors are abundant throughout the body, and just as abundant in cholangiocytes as alveolar cells, posing the question of possible direct liver injury. While liver enzymes and function tests do seem to be deranged in some patients, it is questionable if the injury is due to direct viral damage, drug-induced liver injury, hypoxia, or microthromboses. Likely, the injury is multifactorial, and management of infected patients with pre-existing liver disease should be taken into consideration. Ultimately, a vaccine is needed to aid in reducing cases of SARS-CoV-2 and providing immunity to the general population. However, while considering the types of vaccines available, safety concerns, particularly of RNA- or DNA-based vaccines, need to be addressed.

1. Introduction

A novel strain of coronaviridae (SARS-CoV-2) was first reported in the Wuhan province of China in December 2019. As of 8 May 2020, it has spread to 215 countries with 265,961 deaths worldwide [1]. On 11 March 2020, the World Health Organisation categorised the outbreak as a pandemic [2,3].

The SARS-CoV-2 virus is a single stranded RNA, enveloped, beta coronavirus characterised by spikes protruding from the surface [4]. Normally found in mammals, birds, and reptiles, this strain has not previously been identified in humans [5]. Previous strains of coronavirus outbreaks in humans include Middle East Respiratory Syndrome (MERS) in 2012 and Severe Acute Respiratory Syndrome (SARS) in 2003 [5,6].

Similar to SARS, SARS-CoV-2 is primarily transmitted by respiratory droplets produced by infected persons when they sneeze, cough, or are deposited on surfaces, where they are transmitted through contact. However, as SARS-CoV-2 has been detected in the gastrointestinal tract, urine and saliva, other routes of transmission have been considered [7,8].

COVID-19 disease refers to infection with the SARS-CoV-2 virus. Incubation time is within 14 days following exposure, with a median of four days [7,9]. Although often asymptomatic (with frequency estimated between 17% and 88% of cases) [10,11,12,13,14], infection initially presents with a broad spectrum of symptoms that typically includes general malaise, fever (commonly over 37 °C), dry cough, shortness of breath, anosmia/dysgueusia, headaches, and muscle aches [7,11,15,16,17]. Some other viral related symptoms, albeit less common, can also be seen—sore throat, chest pain, nausea, vomiting, diarrhoea, skin rashes, and vasculitic manifestations. Severe infection seems to present a biphasic pattern [18,19,20,21]. A first phase (‘viremia’), corresponding to viral invasion of the body, causes symptoms as described above. This phase is followed by an ‘inflammatory’ phase, corresponding to excessive host inflammatory response (‘cytokine storm’), responsible for severe cardiopulmonary manifestations, sometimes leading to acute respiratory distress syndrome, shock, and death [18,19,20,21]. Respiratory symptoms, in particular hypoxia, have been the main indication for hospitalisation.

It has been reported that 14.8–53% of SARS-CoV-2 patients had liver injury indicated by abnormal liver function tests—mainly elevated alanine aminotransferase (ALT), hypoalbuminemia, and elevated gamma-glutamyl transferase (GGT) [22,23,24]. These abnormalities seem to occur during either the viremia or inflammatory phase. Reduced albumin can be due to inflammatory response while raised levels of GGT and bilirubin are associated with biliary damage. This is confirmed in recent reports that SARS-CoV-2 has a much greater affinity for biliary cells (cholangiocytes), which have higher expression of ACE2 receptors compared with hepatocytes [7,22,25]. Significant liver injury with raised levels of ALT, Bilirubin, variable levels of alkaline phosphatase and GGT has been reported in 58–78% of patients with severe clinical manifestations of COVID-19 disease, being a surrogate marker for adverse outcome [4,7,15,22,25] (Table 1).

Table 1

Liver enzyme abnormalities in COVID-19 disease vary and reflect the degree of inflammatory response, direct biliary injury by the virus, the presence or absence of ischemia/microthromboses, and possible drug-induced liver injury. Hypoalbuminaemia and high transaminases levels are associated with poor prognosis.

AlbuminTransaminasesGGTBilirubinAlkaline Phosphatase
COVID-19Severe liver injury from inflammatory response (cytokine storm)An external file that holds a picture, illustration, etc.
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Drug induced liver injuryVariableAn external file that holds a picture, illustration, etc.
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Direct biliary injuryVariableVariableAn external file that holds a picture, illustration, etc.
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This review summarises the up to date knowledge on liver injury in the context COVID-19 disease in patients with or without pre-existing liver disease. We also discuss possible mechanisms of liver injury and the current advice regarding management of liver disease patients including liver transplant recipients.Go to:

2. Viral Entry and Effect on Liver

SARS-CoV-2 enters the host via the Angiotensin-converting enzyme 2 (ACE2) receptor. It has been suggested that SARS-CoV-2 binds ACE2 receptors more efficiently than previous COVID viruses, allowing for its extensive transmission [26].

ACE2 is found in a variety of tissues (heart, liver, lung, bladder, kidney, and pancreas); however, it is known to be abundant in alveolar cells accounting for the viral injury to lungs of infected patients [22]. While there is conflicting evidence of ACE2 receptor density in the liver, current reports using single-cell RNA sequencing have confirmed that cholangiocytes have the highest levels of ACE2 receptors [7,22,25,27]. Hu et al. used in silico and in vitro techniques to sample hepatocytes, cholangiocytes, Kupffer cells and other components of fresh liver samples [25]. They found that 59.7% of cholangiocytes had ACE2 receptors in comparison to only 2.6% of hepatocytes. This data suggests that cholangiocytes have the same percentage of ACE2 receptors as aveolar type 2 cells [25]. Further, it has been suggested that infection of cholangiocytes may be the source of the virus found in faeces [28]. While the presence of a receptor is needed for the virus to gain entry into the host, it is still unclear if other conditions are also needed or could possibly aid the virus.Go to:

3. Possible Causes of Elevated Liver Enzymes

Emerging data for abnormal liver enzymes seen in SARS-CoV-2-infected patients raises several questions. Are these abnormalities due to direct viral damage, drug-induced liver injury (DILI), unknown pre-existing liver disease, or indirect consequence of viral damage to other systems (cardiopulmonary, haemostasis)? Liver samples from infected patients were examined, and moderate microvascular steatosis with mild lobular and portal activity were reported [29]. It does seem likely that damage that may affect liver function could principally be due to hypoxia and shock, although a direct effect of SARS-CoV-2 to the liver or DILI can also be contributing factors [29,30].

3.1. Direct Viral Damage

While mechanisms of direct damage to the liver remain unclear, concerns about viral damage have already been raised, e.g., with a case of SARS-CoV-2 infection concurrent with liver failure, without other apparent cause, recently described in Germany [31].

However, direct viral damage has been contested by some, and other explanations have been offered, which will be discussed below [30] (Figure 1).

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Figure 1

Liver injury in SARS-CoV-2. There are multiple reports of increased liver enzymes and liver dysfunction in SARS-CoV-2 patients presenting with elevated alanine transaminase (ALT), gamma-glutamyl transferase (GGT), bilirubin, and monocyte chemoattractant protein 1 (MCP1). Taken together with lower levels of albumin, this points to liver damage with possible injury to biliary cells. Liver injury is most likely multifactorial and seen mainly in patients at the severe end of the disease spectrum.

3.2. Drug-Induced Liver Injury

A study by Fan et al. [29] has raised the question of DILI as a possible cause of liver injury seen in COVID-19 patients. They show that patients given lopinavir or ritonavir after admission presented higher incidence of liver injury and required longer stay in hospital. It is also possible that these patients were given antivirals because they had a more severe presentation that might have affected their liver in the first place. Though recent evidence suggests lopinavir and ritonavir had no clinical effect on SARS-CoV-2, perhaps future application of antiviral drugs should also take into account their effects on the liver [32].

Many infected with SARS-CoV-2 regularly use paracetamol as it is the recommended antipyretic medication. Unintentional overdose with paracetamol contributing to raised ALT cannot be excluded in patients’ non-remitting pyrexia, as paracetamol is a well-recognised cause of fulminant hepatic failure [33]. This also needs to be taken into consideration when evaluating liver injury in these patients.

Several drugs have been trialled on SARS-CoV-2 patients such as hydroxychloroquine and azithromycin with ambiguous results on the virus but possibly exacerbating liver injury [34]. This ambiguity leads to many questions involving the management of SARS-CoV-2 and pre-existing liver disease.

3.3. Hypoxic Liver

Sepsis complicating severe COVID-19 illness and hypoxia can also be significant contributing factors [30]. Hypoxic liver injury can be characterised by an increase in transaminases due to an imbalance of oxygen supply [35]. This typically occurs in the elderly with right side congestive heart failure [35]. Though the median age of patients contracting SARS-CoV-2 is 47 years of age, the elderly have proven to be particularly vulnerable, with increasing age an indicator of mortality [7,27]. In the elderly population, it is likely that a rise in liver enzymes, particularly transaminases, is due to pre-existing conditions.

3.4. Microthromboses

SARS-CoV-2 has been shown to lead to a hypercoagulable state, therefore increasing thromboembolism risk [36,37,38]. It has recently been reported that in certain patient groups, often younger patients, micro vascular thromboses can cause end stage organ damage and may potentially affect the liver. It is also notable that high levels of alkaline phosphatase have been used as a prognostic value for ischemic stroke patients and in identifying high risk haemorrhagic transformation and are also shown to be high in COVID-19 patients suffered thrombotic events, although in other cases, alkaline phosphatase levels have been normal or very mildly raised [39,40,41].

Results of autopsies from Wuhan province, China, have also shown infiltration of lymphocytes and monocytes in the portal area with microthrombosis and congestion of hepatic sinuses [27]. The liver was described as having hepatocyte degeneration accompanied by lobular focal necrosis and neutrophil infiltration. Though histological features of liver failure and bile duct injuries were not observed in these cases [27].

3.5. SARS-CoV-2 in Patients with Pre-Existing Liver Disease

Patients with pre-existing conditions have shown increased susceptibility to SARS-CoV-2. At present, it is unclear to what extent pre-existing liver disease contributes to liver injury seen in SARS-CoV-2 patients. A very recent study conducted in the UK on more than 17 million people has identified pre-existing liver disease as an independent risk factor of death in SARS-CoV-2 infections [42].

For instance, it has been shown that patients with SARS-CoV-2 show an increase of monocyte chemoattractant protein 1 (MCAP1), which is a chemokine known to exacerbate steatohepatitis [34]. A recent short communication describes possible implications for patients with non-alcoholic fatty liver disease (NAFLD) [43]. NAFLD patients, alongside those with metabolic syndrome and type 2 diabetes, are often treated with ACE inhibitors, which have anti-inflammatory and anti-obesity effects. While there has not been a reported effect on mortality of ACE inhibitor drug use, it has been speculated that ACE inhibitors up-regulate the ACE2 receptor and therefore can increase viral load in patients taking these medications [43,44]. NAFLD patients often exhibit increased cytokine levels due to their chronic inflammatory stage. Prins and Olinga suggest that this predisposition, in patients infected with SARS-CoV-2, could expedite the progression of NAFLD to a more aggressive non-alcoholic steatohepatitis [43].

There is suggestion that derangement of liver function should be taken into consideration alongside other physiological values [11,30]. Patients with SARS-CoV-2 have exhibited increased levels of creatine kinase, lactate dehydrogenase, ferritin, C-reactive protein, and myoglobin alongside liver dysfunction, and it has been suggested that liver damage is collateral, caused by induced cytotoxic T cells and the induction of the innate immune response rather than direct injury from the virus itself, as observed with other respiratory viruses [11,16,30,45].

Regardless of the source of injury, it is clear that managing those with pre-existing liver disease needs to be thought out carefully during this pandemic and in future outbreaks of coronavirus infection. These patients are at higher risk of being infected and of more severe COVID-19 disease and should be practising strict social distancing or shielding if they take steroids or immunosuppressive therapies [46]. The British Liver Trust has recently called on the UK government to classify those with extreme liver disease as ‘extremely vulnerable’ [47]. Recent reports suggest that more than 1/3 of cirrhotic patients who developed SARS-CoV-2 died [48]. A new international registry developed between the University of Oxford and the University of North Carolina has shown that those with decompensated cirrhosis are at most risk and are calling on hospitals to routinely test patients with deranged liver function/enzyme results for SARS-CoV-2 so early observation and treatment may prevent further deterioration. The British Liver Trust also suggests that all patients with decompensated cirrhosis practice social shielding, a step up from social distancing, even though it is not yet part of the formal guidance [47].

Boettler et al. have published comprehensive recommendations for management and surveillance of patients with liver disease throughout the SARS-CoV-2 outbreak [28]. This paper now forms the official position of the European Association for the Study of Liver and the European Society of Clinical Microbiology and Infectious Disease [49]. They suggest prioritization of outpatient clinics, inpatient admission depending on presence of certain risk factors, reducing exposure through social distancing (remodelling waiting areas, reduction of waiting times, reduction of face to face contact through telemedicine), and carefully considering the benefits of patient care weighed against the risk of infection.Go to:

4. Disease Severity in the Immunocompromised and Transplant Patients

Under ordinary conditions, organ transplant recipients and those on immunosuppressants are at high risk of infection due in particular to the suppression of T cell response, making their susceptibility to SARS-CoV-2 and prognosis, if infected, unclear. On one hand, it has been postulated that reduction of systemic inflammation by immunosuppressants could improve outcome for COVID-19 patients as the severity of inflammatory response can be an indicator of prognosis [50]. However, it is also a case that immunosuppressed individuals tend to have a higher viral load, take longer to shed the virus, and may show more severe clinical symptoms with a poorer prognosis [51].

Zhu et al. reported on 10 SARS-CoV-2-positive renal transplant recipients in Wuhan, China [51]. All were admitted to hospital with significant progressive pneumonia. The severity of pneumonia in this group was recorded as greater than their infected family members and others in the local population. In accordance with Influenza A/HINI guidance, calcineurin inhibitors were stopped in seven patients for nine days and in one patient for 12 days [51,52]. Within this group, there was no acute renal graft rejection, and all patients eventually recovered from COVID-19, though it took longer for them to become SARS-CoV-2-negative than their infected family members [51]. They attributed the length of infection but eventual recovery to the hypothesis that long-term immunosuppression might delay viral clearance and prolong the course of disease but avoid fatal pneumonia caused by a hyperimmune response [51].

Another study of 90 SARS-CoV-2-positive transplant patients in New York City also described reducing antimetabolites, steroids, and/or calcineurin inhibitors in 55 patients [53]. Pereira et al. categorized patients as mild (outpatient care only), moderate (admission as general inpatient), or severe (mechanical ventilation, admission to intensive care unit, or death) [53]. Within this group, 24% presented with mild disease, 46% moderate, and 30% severe. As with other studies, advanced age and comorbidities were associated with disease severity [7,27,39,40,41]. Type of transplant and time of viral infection after transplant were not statistically significant factors [53]. Laboratory values were similar between moderate and severe cases, though albumin was lower in the severe group [53].

At present there is little data regarding the use of immunomodulatory agents such as tocilizumab or sarilumab when trying to suppress the ‘cytokine storm’ in these patients [53]. Pereira et al. noted that 14 patients receiving 1–3 doses each of tocilizumab and 16 patients receiving bolus steroids showed no adverse outcomes at the time of their publication [53]. They also noted that while all biomarkers of inflammation were elevated, procalcitonin was the only marker which differed between moderate and severe disease and suggested that the chronically immunosuppressed may undergo a uniquely dysfunctional inflammatory response to SARS-CoV-2. This was further supported by Lippi et al., who showed that high levels of procalcitonin can be a predictor of severe COVID-19 syndrome and potentially related to secondary bacterial infection [54]. From this study there were no confirmed cases of thromboembolic complications or organ rejection [53].

Many epidemiological reports regarding treatment and prognosis of COVID-19 syndrome are based on the general population who would have had healthy immunity before viral infection, thus overlooking important data for immunocompromised patients [53]. Many such patients present with atypical signs and symptoms leading to missed diagnosis, late presentation, and worse prognosis [50]. At the time of this publication, no significant conclusions have been drawn regarding the outcome of COVID-19 in patients in receipt of immunosuppressive therapy. More research into cytokine activation, T cell signalling and migration, and viral clearance are needed [53]. The postulated anti-inflammatory benefits of immunosuppression should be balanced against the possibility of inhibiting anti-viral immunity by delaying viral shedding and possible organ rejection for those patients having undergone transplant [50].Go to:

5. Vaccination for SARS-CoV-2

Ultimately, a vaccine against SARS-CoV-2 will be key in preventing spread of virus and loosening social restrictions, but many factors need to be considered in the development of a vaccine so as not to increase innate immune response, increase likelihood of autoimmune diseases, or further DILI.

Vaccinations are costly and usually take years to complete stringent animal and human trials before being made available to the public. However, in an epidemic or pandemic situation, the scientific community faces increasing pressure to rapidly respond with an effective vaccine. In previous epidemics such as Ebola, H1N1, SARS, and MERS, vaccine development was never completed due to the epidemic ending and funds being reallocated [55].

In the context of this review, it is important to highlight that one possible side effect of vaccinations could result in liver damage. Vaccines with the greatest potential, in pandemic situations, are RNA- or DNA-based vaccines [55]. These vaccines do not need to be cultured or require fermentation, they avoid risks of working with live pathogens, and can specifically encode key antigens without also coding for other toxins, but they are not without risks [55,56].

There are no approved RNA vaccines to date, as toxicity cannot always be predicted from animal studies due to species differences between human and animals [55]. Some effects seen in previous RNA-based vaccinations have been pancreatitis, lactic acidosis, liver steatosis, nerve damage, and death [55]. Liver toxicity was reported in preclinical studies using RNA therapy for Crigler–Nayjor syndrome, and in an RNA-based rabies vaccination trial, an increased and deleterious inflammatory response ended the trial [55]. This is possibly due to type 1 interferon induction by RNA, which is known to induce autoimmune diseases [55]. DNA-based vaccines have also been implicated in inducing an innate immune response through toll-like receptor (TLR) 9 and non-TLR activation [56].Go to:

6. Conclusions

SARS-CoV-2 is a novel coronavirus known to cause respiratory infections with severity ranging from mild cold- and flu-like symptoms to fatal pneumonia. While respiratory based, if severe, it can cause dysfunction of other organs such as the kidneys and liver. It is likely that the liver injury seen in SARS-CoV-2-positive patients is multifactorial and the result of a combination of inflammatory response, sepsis, hypoxia, microthrombotic events, DILI, and viral damage. Pre-existent liver disease is an independent risk factor of death in SARS-CoV-2 related infection, and severity of liver damage most likely correlates with COVID-19 disease severity. Nevertheless, abnormalities in the liver function tests of these patients, without pre-existing liver disease, may have prognostic significance and predict adverse outcomes. Patients with chronic liver disease and in particular those on immunosuppressive therapies including liver transplant recipients should be particularly careful and managed according to internationally accepted guidelines regarding strict social distancing or shielding.

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Intestinal Damage in COVID-19: SARS-CoV-2 Infection and Intestinal Thrombosis

Authors: Xiaoming Wu1Haijiao Jing1Chengyue Wang1Yufeng Wang1Nan Zuo1Tao Jiang2*Valerie A. Novakovic3 and Jialan Shi1,3,4* Front. Microbiol., 22 March 2022 | https://doi.org/10.3389/fmicb.2022.860931

The intestinal tract, with high expression of angiotensin-converting enzyme 2 (ACE2), is a major site of extrapulmonary infection in COVID-19. During pulmonary infection, the virus enters the bloodstream forming viremia, which infects and damages extrapulmonary organs. Uncontrolled viral infection induces cytokine storm and promotes a hypercoagulable state, leading to systemic microthrombi. Both viral infection and microthrombi can damage the gut–blood barrier, resulting in malabsorption, malnutrition, and intestinal flora entering the blood, ultimately increasing disease severity and mortality. Early prophylactic antithrombotic therapy can prevent these damages, thereby reducing mortality. In this review, we discuss the effects of SARS-CoV-2 infection and intestinal thrombosis on intestinal injury and disease severity, as well as corresponding treatment strategies.

Introduction

COVID-19 has become a worldwide pandemic causing widespread illness and mortality. SARS-CoV-2 mainly infects the respiratory tract through attachment to angiotensin-converting enzyme 2 (ACE2) receptors (Lan et al., 2020). ACE2 is also highly expressed on intestinal epithelial cells, allowing SARS-CoV-2 to infect the intestinal tract (Xiao et al., 2020a). Recent meta-analyses show that 48%–54% of fecal samples from COVID-19 patients have tested positive for viral RNA, and 15%–17% of patients have gastrointestinal (GI) symptoms (Cheung et al., 2020Mao et al., 2020Sultan et al., 2020). Additionally, live virus can be isolated from fecal samples of COVID-19 patients (Wang et al., 2020). Some studies have proposed fecal–oral transmission as the cause of intestinal infection (Guo et al., 2021). However, direct evidence for fecal–oral transmission is still lacking. Meanwhile, the virus has been detected in the blood of both symptomatic and asymptomatic patients (Chang et al., 2020), and disseminated virus could infect extrapulmonary organs (Jacobs and Mellors, 2020). Thus, the potential that intestinal infection occurs via blood transmission should be carefully considered.

Pulmonary infection triggers cytokine storm and induces a prothrombotic state (McFadyen et al., 2020Moore and June, 2020). Venous and arterial thrombosis are common in COVID-19 (Moore and June, 2020). Systematic reviews estimate that 14%–31% of in-hospital patients develop a clinically apparent thrombotic event (Suh et al., 2021Tan et al., 2021), while autopsy reports show a high prevalence of microthrombi in multiple organs, including lung, heart, liver, kidney, and gastrointestinal tract (Bradley et al., 2020Polak et al., 2020). A cohort study showed that COVID-19 patients with intestinal ischemia had markedly elevated D-dimer levels and poor outcomes (Norsa et al., 2020). Additionally, recent studies have shown that mesenteric thrombosis often results in intestinal resection and significantly increases mortality (Bhayana et al., 2020El Moheb et al., 2020). Therefore, it is essential to outline the mechanisms of intestinal thrombosis and its contribution to intestinal damage and disease progression.

In this review, we discuss blood transmission as a potential route for intestinal infection. We then summarize the characteristics and mechanism of intestinal thrombosis formation in COVID-19. Next, we focus on the effects of intestinal infection and thrombosis on intestinal damage and disease severity. Finally, we discuss therapeutic strategies to prevent intestinal damage.

Gastrointestinal Symptoms and SARS-CoV-2 Infection

Multiple studies have reported GI symptoms in COVID-19 patients, including diarrhea, nausea, vomiting, anorexia, and abdominal pain (Cheung et al., 2020Mao et al., 2020Sultan et al., 2020). According to a meta-analysis comprising 10,890 COVID-19 patients, the pooled prevalence estimates of GI symptoms were: diarrhea (7.7%), nausea or vomiting (7.8%), and abdominal pain (2.7%; Sultan et al., 2020) with 10% of these patients reporting GI symptoms as being their initial symptoms (Cheung et al., 2020). These data indicate potential gastrointestinal infection by SARS-CoV-2, which is reported to infect and replicate in epithelial cells of human small intestinal organoids (Zang et al., 2020). Both viral nucleocapsid proteins and viral particles have been detected in infected patient intestinal biopsies (Livanos et al., 2021). Additionally, SARS-CoV-2 RNA and live virus can be found in the stool of patients (Wang et al., 2020). More importantly, SARS-CoV-2 subgenomic mRNA is transcribed in actively replicating cells and has been detected in fecal samples (Wölfel et al., 2020). Further, rectal viral shedding persists for longer than that of the respiratory system (Zhao et al., 2020). All these data demonstrate that SARS-CoV-2 directly infects and replicates in intestinal epithelial cells of patients.

Intestinal Infection and Transmission Routes

With the deepening understanding of COVID-19, GI symptoms have been recognized as early signs of the disease. The high expression of ACE2 in the GI tract, isolation of live virus from fecal samples, and a subset of patients presenting with only GI symptoms seem to suggest fecal–oral transmission. However, problems with the feasibility of this mode of transmission remain. First, studies have shown that SARS-CoV-2 loses infectivity in simulated gastric acid within 10 min (Chan et al., 2020Zang et al., 2020Zhong et al., 2020). Secondly, SARS-CoV-2, as an enveloped virus, is largely unable to withstand the detergent effect of bile salts and the activity of digestive enzymes in the duodenum (Figure 1). Although some studies have suggested that highly viscous mucus in the gastrointestinal tract protects SARS-CoV-2, allowing the virus to retain its infectivity (Guo et al., 2021Zhang H. et al., 2021), there is still a lack of direct evidence. Bushman et al. (2019) had previously investigated the links between the structures of viruses and routes of transmission and found a strong association between fecal–oral transmission and the absence of a lipid envelope. Lastly, although some studies have isolated intact viruses from feces (Wang et al., 2020Zhang Y. et al., 2020Zhou et al., 2020Xiao et al., 2020b), most of them have not further confirmed the infectivity of these viruses (Wang et al., 2020Zhang Y. et al., 2020Xiao et al., 2020b). Zhou et al. (2020) confirmed viral propagation by RT-PCR, but only in a single fecal sample. Previous research has shown that SARS-CoV-2 is completely inactivated in simulated human colonic fluid over the course of 24 h, which may explain the sporadic detection of infection-active SARS-CoV-2 from feces samples.FIGURE 1

Figure 1. Intestinal infection and transmission routes. ① Direct evidence for fecal–oral transmission is still lacking. SARS-CoV-2 may be unable to enter the small intestine from the stomach due to gastric acid, bile and digestive enzymes. ② SARS-CoV-2 released from type II alveolar cells infects alveolar capillary endothelial cells (ECs). The virus replicates in ECs and is released into the blood to form viremia. ③ SARS-CoV-2 is released from infected ciliary cells of the nasal cavity and breaks through the basement membrane, infecting the vascular ECs and eventually entering circulation. ④ Blood transmission after alveolar or nasal infection is a potential route of intestinal infection. Eventually, SARS-CoV-2 is released into the gut and infects surrounding intestinal epithelial cells along the intestinal tract. ⑤ SARS-CoV-2 in the gut can also enter the capillaries and cause viremia, leading to recurrence of disease.

Several lines of evidence suggest that SARS-CoV-2 may infect the intestinal tract via the bloodstream. Deng et al. (2020) detected SARS-CoV-2 RNA in anal swabs from intratracheally but not intragastrically infected rhesus macaques, suggesting blood transmission. Indeed, SARS-CoV-2 RNA has been detected in blood and urine samples of patients (Wang et al., 2020). The virus can also be detected in multiple organs (including heart, brain, and kidney) and is associated with organ injury, indicating that the virus can reach and infect extrapulmonary organs (Puelles et al., 2020). Another study showed that SARS-CoV-2 viremia was associated with intestinal damage, independent of disease severity (Li Y. et al., 2021). Thus, blood transmission could be the cause of intestinal infection. Specifically, SARS-CoV-2 replicating in alveolar epithelial cells and capillary ECs is released into the bloodstream and infects new vascular ECs. The capillary network is then the main route by which the virus enters and infects extrapulmonary organs. The extensive surface area of intestinal capillaries makes intestinal epithelial cells more susceptible to infection than other extrapulmonary organs. Following infection of intestinal capillaries, SARS-CoV-2 is released into the gut and infects surrounding intestinal epithelial cells along the intestinal tract (Figure 1). Once established in the gut, SARS-CoV-2 can also reenter the capillaries, potentially leading to recurrence of disease. Consistent with this, in patients who experienced recurrence, the phylogenetic analysis of infection samples has shown that recurrent virus evolves from the original parent virus (Hu et al., 2020).

Additionally, SARS-CoV-2 RNA can also be detected in the blood and urine of asymptomatic patients, suggesting a second pathway to viremia through the nasal cavity (Chang et al., 2020Hasanoglu et al., 2021). The abundant blood vessels, thin mucous membrane, and higher levels of ACE2 (Huang et al., 2021) make it possible for the virus to initiate viremia from the nasal cavity. Specifically, SARS-CoV-2 is released from infected ciliary cells of the nasal cavity and breaks through the basement membrane, infecting the vascular ECs and eventually entering circulation (Figure 1). Blood transmission after nasal infection is therefore another potential route of intestinal infection.

Intestinal Damage, Malnutrition, and Poor Outcomes

A recent study has shown that a fecal sample positive for SARS-CoV-2 RNA at any time during hospitalization was associated with higher mortality [HR: 3.4 (1.2–9.9); Das Adhikari et al., 2021]. Similarly, another study showed that small-bowel thickening on CT was strongly associated with ICU admission (Wölfel et al., 2020). This relationship did not hold for colon or rectal thickening. These data indicates that small-bowel damage contributes to poor outcomes. As the main organ for nutrient absorption, damage to the small intestine will result in malabsorption and malnutrition, both of which commonly occur in COVID-19 patients (Di Filippo et al., 2021Lv et al., 2021) and are associated with disease severity (Luo et al., 2020Zhang P. et al., 2021). A fecal metabolome study showed that feces of COVID-19 patients were enriched with important nutrients that should be metabolized or absorbed, consistent with malabsorption (Lv et al., 2021). A prospective study showed that 29% of COVID-19 patients (31% of hospitalization patients and 21% of patients quarantined at home) had lost >5% of body weight [median weight loss, 6.5 (5.0–9.0) kg or 8.1 (6.1–10.9) %; Di Filippo et al., 2021]. Those patients with weight loss had greater systemic inflammation, impaired renal function and longer disease duration. A large, multicenter study (including 3,229 patients with GI symptoms) showed that 23% of patients had malnutrition, of whom 56.4% were unable to gain weight after 6 months follow-up (Rizvi et al., 2021). Studies also showed that malnutrition was associated with higher incidences of acute respiratory distress syndrome, acute myocardial injury, secondary infection, shock, and 28-day ICU mortality (Luo et al., 2020Zhang P. et al., 2021). Overall, malabsorption and malnutrition due to damaged small intestine increased disease severity and mortality.

Nutrient absorption in the small intestine is mainly through ATP-dependent active transport. Intestinal infection, hypoxemia, and intestinal ischemia contribute to malabsorption. SARS-CoV-2 adhesion depletes ACE2 levels on intestinal epithelial cells, which alters the expression of the neutral amino acid transporter B0AT1, reducing the intake of tryptophan and the production of nicotinamide (D’Amico et al., 2020). Meanwhile, uncontrolled viral replication consumes large amounts of ATP and nutrients, resulting in decreased nutrients entering the bloodstream. More importantly, anaerobic glycolysis caused by hypoxemia and intestinal ischemia significantly decreases ATP and active transport, leading to malabsorption. Additionally, hypoxemia and intestinal ischemia can also cause anorexia, nausea, vomiting, and enteral nutrition intolerance, reducing food intake. A prospective multicenter study showed that reduced food intake was associated with higher ICU admission and mortality (Caccialanza et al., 2021).

Intestinal Ischemia and Thrombosis

Intestinal ischemia is a common manifestation in COVID-19 patients. Autopsy results have shown that 31.6% of deceased patients had focal ischemic intestinal changes (Chiu et al., 2020). In a separate imaging study, bowel wall thickening and pneumatosis intestinalis, which indicate intestinal ischemia, were found on 38.1% (16 of 42) of abdominal CT images (Bhayana et al., 2020). Of these, 4 (9.5%) patients with pneumatosis intestinalis developed severe intestinal necrosis and needed resection. In another cohort study, 55.8% (58/104) of ICU patients developed an ileus (Kaafarani et al., 2020). Although mechanical factors cannot be ruled out, insufficient intestinal motility due to intestinal ischemia was more likely to be the cause of ileus in COVID-19 patients. In these patients with ileus, 4 (3.8%) developed severe intestinal ischemia and require emergency surgery. Both studies found microthrombi in these resected intestinal samples, which were the main cause of intestinal ischemia and increased mortality.

Additional intestinal ischemia and necrosis follows the formation of mesenteric thrombosis. However, there is currently relatively little data of mesenteric thrombus in COVID-19. Therefore, we have summarized the characteristics of 40 patients in 39 case reports published on PubMed (Supplementary Table 1). The median age of these patients was 50 (20–82) years, 26 (65%) were male, 38 (95%) developed bowel ischemia or necrosis, 30 (75%) needed bowel resection, 7 (17.5%) required no surgery, at least 3 (7.5%) developed sepsis, and 13 (32.5%) died. Other abdominal thrombotic events (such as celiac aortic thrombosis) leading to mesenteric ischemia can also result in severe intestinal necrosis and require intestinal resection (Zamboni et al., 2021).

Mild intestinal ischemia can lead to reduced diet and malabsorption. Severe intestinal ischemia or necrosis leads to the dissemination of gut bacteria, endotoxins, and microbial metabolites into the blood (Figure 2 bottom), aggravating hyperinflammation and the hypercoagulability state. Such patients need emergency excision of the necrotic bowel, which significantly increases mortality.FIGURE 2

Figure 2. Intestinal thrombosis leads to intestinal mucosal necrosis and dissemination of gut bacteria, endotoxins, and microbial metabolites in blood. (Top) Mesenteric vascular endotheliitis (initiated by viremia and accelerated by cytokines), hyperactivated platelets and high levels of phosphatidylserine (PS) promote a high rate of mesenteric thrombus in COVID-19 patients (mesenteric vein is shown in Supplementary Figure 1). (Bottom) Intestinal microthrombi and hypoxemia rapidly lead to intestinal mucosal ischemia and necrosis. The damaged gut–blood barrier leads to dissemination of gut bacteria, endotoxins, and microbial metabolites in blood.

Long-Term Gastrointestinal Sequelae

Long-term GI complications are common in recovering COVID-19 patients. In one systematic review of post-acute COVID-19 manifestations, diarrhea was among the top 10 most common complaints, with a prevalence of 6%. Other long-term GI symptoms include nausea, vomiting, abdominal pain, loss of appetite, and weight loss (Aiyegbusi et al., 2021Huang et al., 2021). The exact mechanisms of the GI sequelae remain unclear. Recently, persistent endotheliopathy, higher levels of thrombin (Fogarty et al., 2021), and residual SARS-CoV-2 viral antigens in the GI tract (Cheung et al., 2022) were described in convalescent COVID-19 patients. These data suggest that prolonged intestinal infection, persistent endothelial injury (abnormal intestinal–blood barrier), and microthrombi could be causes of the persistent GI symptoms.

The Mechanisms of Intestinal Thrombosis

Damaged Endothelial Cells

Resected bowel samples from COVID-19 patients routinely exhibit thrombi and endotheliitis, indicating the important role of EC injury in mesenteric thrombosis (Bhayana et al., 2020Chiu et al., 2020Kaafarani et al., 2020). SARS-CoV-2 infection (Varga et al., 2020) and elevated inflammatory cytokines (He et al., 2016) damage mesenteric vascular ECs. In response, EC cell margins retract, extending phosphatidylserine (PS) positive filopods and releasing endothelial microparticles (MPs; Figure 3BHe et al., 2016). The PS+ filopods and MPs can be co-stained by Xa and Va and support fibrin formation (Figures 3BD). The exposed PS then activates tissue factor on ECs, triggering the extrinsic coagulation pathway (Versteeg et al., 2013). Next, higher levels of FVIII and vWF released from damaged EC contribute to the hypercoagulable state and platelet aggregation, respectively (Goshua et al., 2020). Thrombomodulin is then released from ECs in its soluble form, which has an attenuated capacity to activate Protein C due to a lack of other cofactors on ECs, such as endothelial protein C receptor (Versteeg et al., 2013). Finally, upregulation of endothelial cell adhesion molecules recruits neutrophils and platelets and further contributes to thrombosis (Tong et al., 2020Li L. et al., 2021).FIGURE 3

Figure 3. Phosphatidylserine exposure on activated/apoptotic cells and microparticles (MPs) promotes fibrin formation. (A) Phosphatidylserine is usually confined to the inner leaflet of the cell membrane. This asymmetry is maintained through ATP-dependent inward transport of PS by flippases and outward transport of non-PS by floppases (left). Upon stimulation, calcium transients will inhibit ATP-dependent transport and stimulate the nonselective lipid transporter scramblase (ATP-independent), resulting in PS exposure (right). (B–D) Human umbilical vein ECs were treated with healthy human plasma and TNF-ɑ (our previous study; He et al., 2016). (B) ECs retracts the cell margins, extends PS positive filopods and releases endothelial-MPs. (C) The PS+ filopods and MPs can be co-stained by Xa and Va. (D) ECs (green) were incubated with MPs-depleted plasma (MDP) in the presence of calcium for 30 min and stained with Alexa Fluro 647-anti-fibrin for 30 min. Considerable fibrin stands among cultured ECs along with filopodia. (E) Confocal images showed PS expression on platelets of patients stained with Alexa 488 lactadherin (our previous study; Ma et al., 2017). MPs from the activated platelet (*) had formed at the margin area located between the distinct outlines. (F) MPs from plasma were co-stained by Xa and Va (or lactadherin and annexin V; our previous study; Gao et al., 2015). (G) MPs that were incubated with recalcified MDP for 30 min and stained with Alexa Fluro 647-anti-fibrin for 30 min. Converted fibrin networks were detected around MPs. The inset bars represent 5 μm in (B–D,G) and 2 μm in (E,F).

Hyperactivated Platelets and Phosphatidylserine Storm

Although COVID-19 patients exhibit mild thrombocytopenia, the remaining platelets are hyperactivated (Manne et al., 2020Taus et al., 2020Zaid et al., 2020). Studies have shown that platelets from COVID-19 patients have increased P-selectin and αIIbβ3 expression. P-selectin on activated platelets interacts with integrin αIIb3 on monocytes to form platelet-monocyte complexes, which induce monocyte tissue factor expression (Hottz et al., 2020). The activated platelets can also induce neutrophils to release neutrophil extracellular traps (NETs; Middleton et al., 2020). Furthermore, platelets from COVID-19 patients aggregate and adhere more efficiently to collagen-coated surfaces under flow conditions (Manne et al., 2020Zaid et al., 2020). Meanwhile, activated platelets release α- and dense-granule contents including FV, FXI, fibrinogen and vWF (Zaid et al., 2020). In addition, activated platelets also produce inflammatory cytokines, fueling cytokine storm (Taus et al., 2020Zaid et al., 2020). Most importantly, activated platelets expose higher levels of PS and release higher numbers of PS+ MPs (Figures 3EGZaid et al., 2020Althaus et al., 2021).

Phosphatidylserine is the most abundant negatively charged phospholipid in mammalian cells and is usually confined to the inner leaflet of the cell membrane (Versteeg et al., 2013). This asymmetry is maintained through ATP-dependent inward transport of PS by flippases and outward transport of other phospholipids by floppases (Figure 3A left). Upon stimulation, transiently increased calcium inhibits ATP-dependent transport and stimulates the nonselective lipid transporter scramblase (ATP-independent), resulting in PS exposure on the outer membrane (Figure 3A right). During this process, microvesicles derived from the budding of cellular membranes will be released. These MPs are typically <1 μm and express PS (Burnier et al., 2009). The exposure of PS on the surface of cells and MPs provides a catalytic surface for factor Xa and thrombin formation in vivo (Versteeg et al., 2013). We have previously demonstrated that PS mediates 90% of Xa and thrombin formation and significantly increases thrombosis in vivo (Shi and Gilbert, 2003).

Cytokines and virus infection can activate blood cells and ECs, resulting in higher levels of PS+ cells and MPs. As COVID-19 progresses, the developing cytokine storm activates more blood cells, leading to PS storm. Platelets are highly sensitive to circulating cytokines, releasing large amounts of cytokines and PS exposed MPs into the plasma (Taus et al., 2020Althaus et al., 2021) and thus are a major contributor to PS storm. Previous studies found an unusual elevation of FVa in severe COVID-19 patients (248 IU/dl, higher than any previous disease; Stefely et al., 2020von Meijenfeldt et al., 2021). The degree of FVa elevation in these patients may be the result of PS storm.

Collectively, SARS-CoV-2 infection is the initiating factor for injury of the intestinal vascular ECs, which is then aggravated by systemic cytokines, leading to endotheliitis. Subsequently, the hyperactivated platelets in circulation rapidly accumulate around the damaged ECs, inducing tissue factor expression, NET release, and activating the intrinsic/extrinsic coagulation pathways. Simultaneously, the high levels of PS expression in circulating cells and MPs further promote thrombin and fibrin formation (Figure 2 top).

Early Antithrombotic Treatment

Vaccines and antithrombotic therapy are effective measures to reduce intestinal damage and fight against the COVID-19 pandemic (Baden et al., 2021Chalmers et al., 2021). Vaccines induce adaptive immunity to clear the virus, reducing intestinal infection and intestinal damage. However, the usefulness of vaccines is limited by incomplete vaccine acceptance and viral mutations (Hacisuleyman et al., 2021Wang et al., 2021). Vaccines are also ineffective for already infected patients. Therefore, more attention should be paid to antithrombotic therapy. Studies had shown that thrombotic events mainly occurred within 7 days of COVID-19 diagnosis (both inpatients and outpatients; Mouhat et al., 2020Ho et al., 2021). Meanwhile, two large randomized controlled trials (RCTs) from the same platform showed that therapeutic anticoagulation reduced mortality in moderate cases but not in severe ones, suggesting that delayed anticoagulant therapy may lead to treatment failure (REMAP-CAP Investigators et al., 2021a,b). More importantly, a recent study reported three asymptomatic COVID-19 patients who developed abdominal (or intestinal) thrombosis leading to intestinal necrosis (Zamboni et al., 2021). All these data suggest that antithrombotic therapy should be initiated once COVID-19 is diagnosed (excluding patients with contraindications). Early prophylactic antithrombotic therapy can reduce the activation of vascular ECs and blood cells, preventing intestinal thrombosis, ensuring sufficient intestinal perfusion, maintaining the normal gut–blood barrier, avoiding malabsorption, malnutrition, and intestinal flora entering the bloodstream. Further, attenuated injury and decreased microthrombi in convalescent patients may lower the risk of long-term GI sequelae. Meanwhile, unobstructed systemic circulation can also accelerate the removal of SARS-CoV-2, inflammatory cytokines and damaged blood cells by the mononuclear phagocyte system.

Anticoagulation

Table 1 summarizes the RCTs of anticoagulant therapy in COVID-19 patients. For outpatients, early anticoagulant therapy reduced hospitalization and supplemental oxygen (Gonzalez-Ochoa). While, delayed treatment had no similar effect (ACTIV-4B and Ananworanich). Thus, oral anticoagulant therapy should be initiated in outpatients once COVID-19 is diagnosed. For non-critically ill patients, therapeutic doses of low molecular weight heparin (LMWH) reduced thrombotic events and mortality, and increased organ support-free days (REMAP-CAP, ACTIV-4a, ATTACC; RAPID; HEP-COVID). However, therapeutic doses of rivaroxaban did not improve clinical outcomes and increased bleeding (ACTION). This is potentially because novel oral anticoagulants do not share the anti-inflammatory and antiviral functions of heparin. Intestinal damage might also result in abnormal absorption of oral anticoagulants. Therefore, therapeutic LMWH should be the first choice for non-critically ill patients. For critically ill patients, RCTs showed that moderate and therapeutic doses were not superior to prophylactic ones. Results from several other studies suggest that the overwhelming thrombosis leads to failure of anticoagulant therapy at therapeutic doses (Leentjens et al., 2021Poor, 2021). Faced with this dilemma, an editorial in N Engl J Med argued that profibrinolytic strategies should be considered (Ten Cate, 2021). More studies are needed to explore optimal antithrombotic therapy in critically ill patients.TABLE 1

Table 1. Randomized clinical trials of anticoagulant therapy in COVID-19 patients.

Inhibition of Platelet Activation

As COVID-19 progresses, cytokine storm activates platelets, which not only participate in primary hemostasis, but also are the major components of PS storm. Autopsy results show a high prevalence of platelet-fibrin-rich microthrombi in lung and extrapulmonary organs, including the gastrointestinal tract (Bradley et al., 2020Polak et al., 2020). Early inhibition of platelet activation can reduce platelet activity and prevent PS storm, thus decreasing thrombosis and mortality. Several observational studies have shown that aspirin decreases mechanical ventilation, ICU admission, and mortality (Chow et al., 2020Santoro et al., 2022). The RCTs testing antiplatelet agents were still preliminary. A recent RCT suggested that aspirin was associated with an increase in survival and reduction in thrombotic events (RECOVERY Collaborative Group, 2022). In addition, anti-inflammatory therapy (e.g., dexamethasone, 6 mg once daily; RECOVERY Collaborative Group et al., 2020) inhibits cytokine storm, as well as platelet activation, reducing mortality. Overall, inhibition of platelet activation is also important to reduce mortality through the prevention of thrombosis and organs damage.

Factors Influencing Antithrombotic Treatment

Thrombotic Risk Factors or Co-morbidities

Studies have shown that obesity, hyperglycemia and diabetes are associated with increased thrombotic events (including intestinal thrombosis), COVID-19 severity, and mortality (Drucker, 2021Stefan et al., 2021). Other thrombotic risk factors include previous venous thromboembolism, active cancer, known thrombophilic condition, recent trauma or surgery, age ≥70 years, respiratory/cardiac/renal failure, and inflammatory bowel disease (Susen et al., 2020). These factors or co-morbidities heighten basal inflammatory levels and endothelial damage, leading to premature cytokine and PS storms, ultimately increasing thrombosis and mortality. Thus, more active antithrombotic therapy strategies should be adopted in these patients. For patients with mild COVID-19 with these factors, the French Working Group on Perioperative Hemostasis and the French Study Group on Thrombosis and Hemostasis recommend higher (intermediate) doses of anticoagulant therapy (Susen et al., 2020). For moderately ill patients, therapeutic doses of anticoagulant therapy should be initiated as soon as possible to prevent excessive microthrombus formation. The need for extended thromboprophylaxis in discharged patients remains controversial. However, a recent RCT showed that rivaroxaban (10 mg/day, 35 days) improved clinical outcomes in discharged COVID-19 patients with higher thrombotic risk factors (Ramacciotti et al., 2022), supporting extended thromboprophylaxis in patients with these risk factors or co-morbidities.

Vaccination

Although more than half the world population has received at least one dose of the vaccines, there are relatively little data of antithrombotic therapy in vaccinated patients. Studies of viral dynamics show that the viral loads of vaccinated patients are as high as that of unvaccinated patients, but drop significantly faster (Brown et al., 2021Klompas, 2021). Thus, vaccinated patients have shorter hospital stays, and are less likely to progress to critical illness and death (Tenforde et al., 2021Thompson et al., 2021). Nevertheless, antithrombotic therapy is still beneficial for the vaccinated patients. Firstly, heparin has anti-inflammatory and antiviral functions and can interfere with the binding of SARS-CoV-2 to ACE2 and shorten the duration of virus infection (Kwon et al., 2020Pereyra et al., 2021). Secondly, antithrombotic therapy protects cells from damage, PS exposure, and microthrombi formation, maintains unobstructed blood circulation, and facilitates virus clearance (by vaccine-induced adaptive immunity). Thirdly, thrombosis remains an important factor in disease progression. Antithrombotic therapy further reduces thrombosis and mortality, especially in vaccinated patients with high risk factors or co-morbidities. Lastly, although vaccines reduce the incidence, a subset of vaccinated patients will still develop long-term sequelae or Long Covid (Ledford, 2021Antonelli et al., 2022). Persistent viral infection and microthrombi are the primary causes (Ledford, 2021Xie et al., 2022), and early antithrombotic therapy is still needed to prevent them.

Conclusion and Future Research

During COVID-19 disease progression, SARS-CoV-2 infiltrates the blood stream from the initial respiratory tract infection, causing viremia, hyperactivated platelets and PS storm. The virus settles into the vascular beds of extrapulmonary organs, ultimately causing infection of intestinal epithelial cell. Damaged ECs, combined with hyperactivated platelets and PS storm, promote intestinal thrombosis, resulting in intestinal ischemia or necrosis. The damaged gut–blood barrier leads to malabsorption, malnutrition and intestinal flora entering the bloodstream, which significantly increase disease severity and mortality. Prolonged intestinal infection, persistent endothelial injury and microthrombi contribute to the long-term GI sequelae after discharge. Early prophylactic antithrombotic therapy can prevent microthrombi, ensuring sufficient intestinal perfusion, maintaining the normal intestinal function, and reducing the risk of long-term GI sequelae. More active antithrombotic therapy should be adopted in patients with other thrombotic risk factors or co-morbidities. Even in vaccinated COVID-19 patients, antithrombotic therapy is also important to decrease (intestinal) thrombosis, mortality and the risk of long-term GI sequelae.

With the Omicron pandemic, patients requiring hospitalization and ICU treatment decline rapidly. However, people are increasingly concerned about Long Covid. In terms of long-term GI sequelae, the detailed mechanisms of prolonged intestinal infection and persistent microthrombi remain unclear. And whether anticoagulant therapy can decrease GI symptoms in patients with long-term GI sequelae deserves further study. Finally, the impact of vaccines on long-term GI sequelae remains unclear in previously infected and breakthrough infected patients.

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The ongoing enigma of SARS‐CoV‐2 and platelet interaction

Authors: Younes Zaid, PhD 1 , 2 and Fadila Guessous, PhD 3 , 4 Res Pract Thromb Haemost. 2022 Jan; 6(1): e12642.Published online 2022 Jan 25. doi: 10.1002/rth2.12642 PMCID:  PMC8787413PMID :  35106430

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Abstract

Since the onset of the global pandemic of coronavirus disease 2019 (COVID‐19), there is an urgent need to understand the pathogenesis of the common inflammatory and thrombotic complications associated with this illness leading to multiorgan failure and mortality. It is well established that platelets are hyperactivated during COVID‐19. Data from independent studies reported an angiotensin‐converting enzyme (ACE2)‐dependent severe acute respiratory syndrome coronavirus 2 (SARS‐CoV‐2) platelet interaction, raising the concern whether ACE2 receptor is the “key receptor” in this process, while other platelet research groups demonstrated that thrombotic events occur via ACE2‐independent mechanisms, where the virus probably uses alternative pathways. In this study, we discuss the conflicting results and highlight the ongoing controversy related to SARS‐CoV‐2‐platelet interaction.

1. INTRODUCTION

Severe acute respiratory syndrome coronavirus 2 (SARS‐CoV‐2) is a highly transmissible and pathogenic coronavirus that caused an outbreak of uncommon viral pneumonia called coronavirus disease 2019 (COVID‐19), which originally emerged in Wuhan, China, in December 2019 (https://covid19.who.int/).

With the pandemic already in its second year, the disease’s impact on the respiratory tract is becoming increasingly obvious. Indeed, it is currently well established that, in addition to the staggering cytokine storm and lung inflammation associated with the infection, thrombotic complications, including microvascular, venous, or arterial thrombosis, and cardiovascular manifestations significantly contribute to the disease severity, leading to morbidity, multiorgan failure, and mortality. 1

Platelets, the small anucleated cellular fragments derived from their megakaryocytes precursors and traditionally linked to thrombosis and hemostasis, are also key players that mediate inflammation, infectious diseases, and immune response. 2

Upon exposure to invading pathogens, platelets contribute to the immunity either directly by cytokine production and anti‐microbial peptides release; or indirectly by amplifying the immune response through interaction with neutrophils, monocytes, and lymphocytes. 3 However, the immunothrombosis triggered during infections may adversely impact immunological and hemostatic processes; thus leading to adverse clinical outcomes. 5

Several recent reports highlighted the association of coagulopathy events and COVID‐19 severity; revealed by elevated levels of D‐dimers and fibrin‐degradation products and hyperactivated platelets in critically ill patients with COVID‐19. 6 However, how platelets interact with the SARS‐CoV‐2 remains controversial. Such controversy is undoubtedly triggering an exciting debate among researchers in the SARS‐CoV‐2–related coagulopathy field during this pandemic.

The recently published conflicting results of this interaction as well as its implication for better management of COVID‐19–related thrombotic complications will be debated here.Go to:

2. PLATELETS ARE HYPERACTIVATED IN COVID‐19

While the pandemic is progressing worldwide, numerous studies have reported that patients with COVID‐19 demonstrated platelet activation, aggregation, and platelet‐leukocyte aggregate formation, thus highlighting the essential role of platelets during SARS‐CoV‐2 infection and immunopathology. 8 10

Compared to donors not infected with SARS‐CoV‐2, platelets isolated from patients with COVID‐19 were hyperactivated when exposed to platelet agonists, such as thrombin, ADP, and collagen. 9 11 Such hypersensitivity could be partially due to increased mitogen‐activated protein kinase signaling pathway activation and thromboxane synthesis. 9 11

Additionally, platelet‐derived microvesicles can be involved in thrombosis, by providing anionic phospholipids, and support coagulation cascade. 12 Similarly, the release of neutrophil extracellular traps (NET), called NETosis, requires platelets and may participate to thrombosis during SARS‐CoV‐2 infection. 13 NETs are networks of extracellular fibers of decondensed chromatin carrying histones and antimicrobial peptides. 14 They also bind blood cells and generate a procoagulant and prothrombotic scaffold. 15 16 In fact, several studies have identified NETs as important components of micro‐ and macrovascular thrombi 17 18 19 and bronchoalveolar lavage fluid 20 in patients with COVID‐19, even after virus clearance from the lungs.

Moreover, upon SARS‐CoV‐2 infection, activated platelets release immune and inflammatory molecules including platelet‐derived growth factor, platelet factor 4 (PF4), RANTES, serotonin, soluble P‐selectin (sP‐selectin), and soluble glycoprotein VI (sGPVI) 8 11 21 22 ; and express a plethora of immune receptors, including CD40L, Toll‐like receptor, and the Fc receptor for the IgG (FcγRIIA). 23 24

A platelet transcriptome study conducted by Manne et al. 9 has revealed transcriptional changes in patients with COVID‐19 distinct from those reported in other viral infections.

Cytokine storm is an umbrella term encompassing several disorders of immune dysregulation characterized by constitutional symptoms, systemic inflammation, and multiorgan dysfunction that can lead to multiorgan failure if inadequately treated. 25 This phenomenon, also known as hypercytokinemia, is a hallmark of COVID‐19, leading to the accumulation of chemokines, cytokines, and several soluble factors, which may activate platelets and other inflammatory cross‐talk pathways. 11 26 Such activation triggers platelet adhesion to the subendothelium and results in thrombus formation, subsequently inducing ischemia and pulmonary embolism. 27

Other viruses can also trigger cytokine storm, including herpesviruses, such as herpes simplex virus, and other influenza viruses, such as H5N1. 25

Platelets are also known to be directly activated by certain viruses. Indeed, in response to direct infection by dengue and influenza viruses, megakaryocytes upregulate interferon‐α genes. 28 29 Similarly, simplex virus‐1 can activate platelet aggregate formation and thrombosis using the previously generated opsonizing antibodies and their interaction with the FcγRIIA. 30 Recently, a key role of platelet‐mediated immunothrombosis in COVID‐19 that signals through FcγRIIA and the C5a–C5a receptor pathway has been identified, revealing the role of platelet hyperactivation in complications associated with SARS‐CoV‐2 infection. 31

However, unlike for other viruses, mechanisms underlying the direct interaction of SARS‐CoV‐2 and platelets and/or megakaryocytes remain a true controversy. Indeed, two recent independent studies conducted in China resulted in conflicting findings. Zhang et al. 21 reported a possible activation of platelets and megakaryocytes directly by SARS‐CoV‐2 as evidenced by the presence of the virus mRNA in platelets of some patients with COVID‐19, while a most recent study revealed that platelet activation occurs through an angiotensin‐converting enzyme (ACE2)‐independent mechanism. 22Go to:

3. DOES SARS‐COV‐2 INTERACT DIRECTLY WITH PLATELETS?

SARS‐CoV‐2 is a positive‐sense single‐stranded RNA virus related to a number of naturally occurring betacoronaviruses. 32

ACE2, the negative regulator of the renin‐angiotensin system, has been recognized as the entry receptor for the SARS‐CoV‐2 infected host cells. 33

There is a little to no expression of ACE2 on most immune cells, including CD4+T cells, CD8+T cells, natural killer T cells, B cells, regulatory T cells, T helper 17 cells, monocytes, dendritic cells, and granulocytes. 34 On the contrary, this receptor is strongly expressed by alveolar epithelial cells, nasopharyngeal airway epithelial cells, and vascular endothelial cells, as well as lung macrophages. 35 Such virus tropism certainly explains the prevailing respiratory symptoms associated with the disease.

Another potential cellular entry process has been proposed for the viral invasion, using the transmembrane serine protease‐2 (TMPRSS2), which is essential for the cleavage of the SARS‐CoV‐2 S protein, thus allowing the fusion of viral and cell membrane and the virus internalization by the cell. 36

Following the onset of the COVID‐19 pandemic and considering its staggering proinflammatory feature, most studies have focused their interest on the expression of ACE2 on immune cells, but a little attention was given to platelets and their megakaryocytes precursors until a few research groups explored such expression on these cells.

Using RNA sequencing (RNA‐seq), reverse transcriptase polymerase chain reaction, and western blot analyses, Manne et al. 9 did not reveal any ACE2 or TMPRSS2 in CD45‐depleted platelets collected from either patients with COVID‐19 or healthy subjects. Using similar approaches, concomitant work by Zaid et al. 11 also reported that there is no detection of ACE2 on platelets derived from patients with COVID‐19 nor from healthy volunteers.

Consistent with these reports, a more recent retrospective survey of plasma samples from a cohort of 62 patients with severe and nonsevere COVID‐19 revealed an increased thrombosis and high levels of sP‐selectin and sGPVI as well as RANTES and PF4 release during platelet activation. However, the characteristics and mechanisms of the direct SARS‐CoV‐2–platelet interaction are yet to be elucidated. 22

In contrast, an independent study carried out by Zhang et al. 21 has shown a strong expression of ACE2 and TMPRSS2 mRNA and protein on platelets from healthy individuals and mice. Moreover, using in vitro assays and in vivo ACE2 transgenic mice, the same group ascertained their findings and reported that the SARS‐CoV‐2 virus and its spike protein induce direct platelet activation.

Another aspect of platelet–SARS‐CoV‐2 interaction was recently reported by Koupenova et al., which reported that SARS‐CoV‐2 initiates programmed cell death in platelets. Indeed, based on platelet RNA analysis by ARTIC v3 sequencing for SARS‐CoV‐2, transmission electron microscopy and immunofluorescence, this group showed that SARS‐CoV‐2 virions became internalized when they were attached to microparticles, bypassing the need for ACE2. Such internalization leads to rapid digestion, apoptosis, necroptosis, and extracellular vesicle release, thus contributing to dysregulated immunity and thrombosis. 37Go to:

4. TAKING TOGETHER THESE DISCORDANT FINDINGS ON WHETHER PLATELETS EXPRESS ACE2 AND/OR TMPRSS2, HOW COULD WE ARGUE IN FAVOR OF ONE OR THE OTHER STATEMENT?

Considering that investigations related to this topic were conducted in different parts of the world, the first possible explanation to such discrepancy would be ethnicity. Indeed, studies carried out by Zaid et al. 11 and Manne et al. 9 included individuals from North Africa and North America, while the cohort studied by Zhang et al. 21 was from Asia. However, in our opinion, this argument would not be valid given that Shen et al. 22 recently investigated patients from Asia as well. This group investigated in vitro SARS‐CoV‐2 infection in human platelets and their megakaryocyte cell line progenitor MEG‐01.

According to this study, the presence of SARS‐CoV‐2 RNA in both MEG‐01 cells and supernatant suggested that the virus may infect and reproduce in megakaryocytes despite insufficient efficiency; nevertheless, no viral particles were localized in MEG‐01 cells as revealed by electron microscopy and immunofluorescence assay (IFA). The authors speculated that platelets may not support SARS‐CoV‐2 duplication; a fact that was echoed by Zaid et al., 11 and Bury et al. 38 Additionally, the lack of ACE2 expression on platelets and megakaryocytes was also shown by western blot and IFA in the study of Shen et al. 22 Similarly, ACE2 and TMPRSS2 RNA were not detectable in a previous microarray‐based integrated plateletomics study that mainly included healthy Black subjects. 39

Besides the ethnicity argument, conflicting findings may be attributed to different technical approaches used to isolate RNA platelets in different investigations. Both studies conducted by Manne et al. 9 and Zaid et al. 11 used CD45+‐depleted washed platelets, a step that eliminated any remaining leukocytes from platelet preparation.

On the contrary, using gel‐purified platelets, Zhang et al. 21 confirmed the absence of white cells by using CD14 marker in their platelet preparation, a step that would probably leave a residual contamination by CD14‐nonexpressing white cells, such as lymphocytes and natural killer cells, thus explaining ACE2 detection in their preparation. Moreover, on the animal level, the same group demonstrated that the administration of SARS‐CoV‐2 spike protein in K18 hACE2 transgenic mice induces platelet hyperactivation and aggregation; however, the expression of this receptor either on platelets or megakaryocytes of these mice remains to be solved and needs to be further investigated, with still the main focus on how these findings in mice could be translated into humans. 39 It is also worth highlighting that a previous RNA‐seq analysis documented the lack of expression of ACE2 and TMPRSS2 by platelets and megakaryocytes in mice. 40

Despite the reported differences regarding the direct SARS‐CoV‐2‐platelet interaction, it is worth noticing that all studies converge toward the same finding that platelets are activated during COVID‐19; and some of them further ascertain that the virus RNA can be found within platelets. 38 All these data advocate in favor of a potential ACE2‐independent mechanism that SARS‐CoV‐2 might use for possible binding and/or entry into platelets.

Using RNA‐seq analysis, Shen et al. 22 showed unchanged levels of glucose‐regulated protein (GRP 78), ADAM1, cathepsin L, GRP1, and asialoglycoprotein 1 in platelets between intensive care unit (ICU) and non‐ICU patients with COVID‐19 and healthy individuals; and reported increased CD147 and kringle‐containing transmembrane protein 1 and reduced neuropilin 1 levels in patients as well as in MEG‐01 cells upon SARS‐CoV‐2 incubation. These data suggest a marked alteration of megakaryocyte and platelet transcriptomic profile, reflecting a similar finding to dengue virus infection. 28

Emerging evidence suggested CD147 as a potential receptor for SARS‐CoV‐2 and its overexpression is associated with certain diseases, such as chronic obstructive pulmonary disease, asthma, that represent risk factors associated with complications during the COVID‐19 pandemic. 28 Nonetheless, the binding of SARS‐CoV‐2 to CD147 is to be uncovered and the role of this receptor in SARS‐CoV‐2 infection remains disputable. 41

In contrast, in their report related to SARS‐COV‐2 directly interacting with platelets via ACE2, Zhang et al. 21 also supported a direct interaction of CD147, SARS‐CoV‐2, and the spike protein.

Based on recent structural studies, CD26 was suggested to be another SARS‐CoV‐2 receptor. 9 Such statement needs to be further investigated since the expression of this receptor on platelets is still debatable. Indeed, previous platelet RNA‐seq and proteomic analysis suggest that neither platelets nor megakaryocytes express CD26, under physiological or infectious conditions. 9 In contrast, the RNA abundance of 14 receptors and cofactors, including CD26, in human platelets and megakaryocytes was explored based on the RNA‐seq data reported in earlier studies and revealed the expression of CD26 on these cells, though at very low levels. 22 Together, these findings could be a hint of a possible CD26‐SARS‐CoV‐2 direct interaction but does not provide tangible data to support a binding solely through this receptor.

In addition to all discrepancy arguments cited above, it is worth mentioning that during this pandemic, there was a rapid and large volume of new COVID‐19 data published in a very short time in the quest to disseminate this new information and insights, thus helping containing the virus spread worldwide.

Therefore, considering that the technical barrier for COVID‐19–related studies was lowered and the well‐intended change of the publication process, the field is more likely to be inconsistent and needs to be revisited experimentally to clarify (Figure 1).FIGURE 1

Proposed model for the SARS‐CoV‐2 and platelet interaction. ACE2, angiotensin‐converting enzyme; CXCR, C‐X‐C chemokine receptor type 4; FcγRIIA, Fc receptor for the IgG; MHC‐1, major histocompatibility complex class 1; SARS‐CoV‐2, severe acute respiratory syndrome coronavirus 2; TLR, Toll‐like receptor; TMPRSS2, transmembrane serine protease‐2; TNFR, tumor necrosis factor receptorGo to:

5. CONCLUSIONS AND RECOMMENDATIONS

As the pandemic is still wreaking havoc across the globe, more studies are carried out and the understanding of the COVID‐19 pathophysiology is continuously evolving, shedding more light on still enigmatic mechanisms underlying platelet hyperactivation during SARS‐CoV2 infection. Following the cytokine storm triggered by the virus infection, platelets’ reactivity may be a critical step in the inflammatory and prothrombotic response, named immunothrombosis.

How the SARS‐CoV‐2–platelet interaction takes place is still obscure, and therefore more studies are warranted to uncover such mechanisms, taking into consideration ethnicity and gearing toward the expression of potential alternative SARS‐CoV‐2 entry receptors or pathways other than the previously established ACE2 receptor and the spike priming serine protease TMPRSS2.

Besides the uneven health care system efficiency in different countries, compiling clinical data worldwide demonstrated an unequal burden of this disease among certain populations, therefore urging the research community to explore a probable population‐based differential expression of SARS‐CoV‐2 key receptors on the surface of platelets and/or other immune cells.

Moreover, as the megakaryocytes are the platelet precursors and considered the cargo carrying all the molecules and factors necessary to platelets’ function before their release into circulation, all future studies should explore the SARS‐CoV‐2 and platelet interaction without losing sight of behavior differences between these two interdependent entities.

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Consequences of COVID-19 for the Pancreas

Authors: Urszula Abramczyk,1,*Maciej Nowaczyński,2Adam Słomczyński,2Piotr Wojnicz,2Piotr Zatyka,2 and Aleksandra Kuzan1 Int J Mol Sci. 2022 Jan; 23(2): 864.Published online 2022 Jan 13. doi: 10.3390/ijms23020864

Abstract

Although coronavirus disease 2019 (COVID-19)-related major health consequences involve the lungs, a growing body of evidence indicates that COVID-19 is not inert to the pancreas either. This review presents a summary of the molecular mechanisms involved in the development of pancreatic dysfunction during the course of COVID-19, the comparison of the effects of non-severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) on pancreatic function, and a summary of how drugs used in COVID-19 treatment may affect this organ. It appears that diabetes is not only a condition that predisposes a patient to suffer from more severe COVID-19, but it may also develop as a consequence of infection with this virus. Some SARS-CoV-2 inpatients experience acute pancreatitis due to direct infection of the tissue with the virus or due to systemic multiple organ dysfunction syndrome (MODS) accompanied by elevated levels of amylase and lipase. There are also reports that reveal a relationship between the development and treatment of pancreatic cancer and SARS-CoV-2 infection. It has been postulated that evaluation of pancreatic function should be increased in post-COVID-19 patients, both adults and children.

1. Effects of Severe Acute Respiratory Syndrome-Related Coronavirus (SARS-CoV) and Middle East Respiratory Syndrome-Related Coronavirus (MERS-CoV) on the Pancreas

Coronaviruses are enveloped, single- and positive-stranded RNA viruses that infect birds and mammals. In humans, coronaviruses cause respiratory tract infection, usually the common cold, but they can also cause severe respiratory illness including severe acute respiratory syndrome (SARS) and Middle East respiratory syndrome (MERS), caused by severe acute respiratory syndrome-related coronavirus (SARS-CoV) and Middle East respiratory syndrome-related coronavirus (MERS-CoV), respectively [1]. Coronaviruses tend to cause epidemics and even pandemics. The first coronavirus pandemic was the SARS outbreak in 2002–2003 [2]. With the experience gained during the SARS pandemic, it was possible to more quickly identify subsequent outbreaks of the MERS epidemic in 2012 [3]. The pathomechanism of both viruses is very similar—they even both use transmembrane protease serine 2 (TMPRSS2), except SARS-CoV uses angiotensin-converting enzyme 2 (ACE2) as its receptor, whereas MERS uses dipeptidyl peptidase-4 (DPP4) [4,5]. Moreover, there is a difference in terms of the severity and frequency of symptoms, which was observed in MERS patients as more frequent hospitalization in the intensive care unit (ICU) compared to SARS patients [2] (Table 1). Diabetes was one of the significant and independent predictors for developing severe SARS-CoV and MERS-CoV [6,7,8]. In MERS, no viral antigen was detected in any tissue other than pneumocytes [7], despite multiple organ dysfunction syndrome in critically ill patients. In SARS-CoV, the presence of the virus was detected not only in respiratory epithelial cells, but also in small intestinal and colonic epithelial cells, in which it also revealed replication features [9]. It is known that the ACE2 receptor is also present in tissues such as the heart, kidney, and pancreas [8,9]. According to some authors, the presence of the receptor is sufficient for tissue entry and pathogenic activity, although other researchers do not support this thesis [9,10]. Yang et al. were some of the first researchers who hypothesized that SARS coronavirus enters islets using ACE2 as its receptor and damages islets causing acute diabetes [8]. Yang’s study revealed that SARS-CoV had a much higher affinity for pancreatic islet cells than for pancreatic exocrine cells, which was consistent with the hyperglycemia observed in some patients and rarely reported acute pancreatitis (AP) [8]. Furthermore, insulin-dependent diabetes mellitus (IDDM) and high fasting blood glucose values were observed in some inpatients [8]. A 3-year follow-up revealed that both abnormalities were transient, which may be indicative of only temporary damage to the pancreatic islets [8]. However, another reason (different from that given by Young et al.) for high fasting blood glucose value in patients may result from increased stress hormones release. Cortisol, catecholamines, growth hormone, and glucagon, which are released during infection, fever, and trauma, can lead to hyperglycemia to the same degree as SARS-CoV can [11]. No information was found in the literature about a direct impact of the MERS virus on the pancreas or on glycemia during or after infection. This may be due to an insufficiently detailed analysis of the available data during previous studies that oscillated primarily, for laboratory tests, between complete blood count (CBC), lactate dehydrogenase (LDH), urea, and creatinine analysis. A summary of SARS-CoV, MERS, and SARS-CoV-2 is shown in Table 1.Table 1. The summary of characteristics of SARS and MERS coronaviruses. Dipeptidyl peptidase-4 (DPP4), transmembrane protease serine 2 (TMPRSS2), hospitalization in the intensive care unit (ICU), and cathepsin L (CTSL).

Table

In 2019, a new coronavirus named SARS-CoV-2 was identified, causing COVID-19. This virus has many characteristics that are analogous to SARS-CoV, for example, ACE2 is also used as its receptor [12]. Patients with diabetes are among those with the most severe forms of COVID-19 and related mortality; insights from recent experience can guide future management [17], particularly for the consequences on the pancreas. As the COVID-19 pandemic has been ongoing for nearly two years, this study aims to collect data concerning the impact of SARS-CoV-2 on the pancreas and analyze them to estimate the future health consequences of COVID-19 in populations.

2. Pancreatic Damage during Diabetes Mellitus and COVID-19

Pancreas tissue damage may cause to the lack of control over normal blood glucose levels in the body. Type 1 diabetes (T1D) is caused by insulin deficiency due to βcell dysfunction of immunologic or idiopathic cause. In contrast, β pancreatic cells in type 2 diabetes (T2D) become depleted over time due to compensatory insulin secretion caused by insulin resistance. There is also type 3 diabetes (T3D), which is described as diabetes associated with the development of Alzheimer’s disease [18]. It should not be confused with type 3c (pancreatogenic) diabetes, which relates to the exocrine and digestive functions of the pancreas. The issue concerning the impairing effect of hyperglycemia (glucotoxicity) on the secretory function of the islets of Langerhans has also been increasingly raised. In addition to endocrine dysfunction, some diabetic patients may also develop moderate exocrine pancreatic insufficiency (EPI), in which pancreatic enzyme secretion is impaired. EPI can be observed in almost all patients with type 3c (pancreatogenic) diabetes (secondary to pancreatic pathology), whereas the prevalence of this dysfunction in patients with T1D or T2D is 40% and 27%, respectively [19].With the ongoing SARS-CoV-2 pandemic, patients with reduced normal pancreatic function are at high risk for COVID-19 requiring hospitalization. In particular, elevated blood glucose levels in patient with and without diabetes makes them at high risk of mortality [20]. Hyperglycemia impairs the immune response (e.g., by reducing the activity of macrophages and polymorphonuclear leukocytes), which in addition influences the excessive cytokine response, and thus has a strong proinflammatory effect.The receptors for ACE2, which are also present in the pancreas, are a target of SARS-CoV-2 in the body, which may result in acute failure of both the islets of Langerhans and exocrine cells [15]. Infection-induced, transient β cell dysfunction may cause an uncontrolled hyperglycemic state, especially in patients whose pancreas is already affected by diabetes mellitus. Persistent hyperglycemia usually predisposes to severe COVID-19 and to viral infection complicated by secondary infections. The aforementioned risk can be found in T1D, T2D, and gestational diabetes mellitus (GDM). In T2D patients, the much more frequent coexistence of other risk factors such as atherosclerosis, hypertension, and obesity should be taken into consideration, which usually implies a worse prognosis for the course of COVID-19 [21,22]. In GDM, SARS-CoV-2 infection not only increases the risk of more severe course of the disease in a patient, but may also result in diabetic fetopathy or, in more advanced pregnancies, increase the risk of future pathologies involving glucose metabolism (such as T2D) in a child [23].

3. Pancreatic Damage in Patients without Pre-Existing Diabetes Infected with SARS-CoV-2

It has been postulated that, either by direct invasion of pancreatic cells by the virus or by indirect mechanisms described below, SARS-CoV-2 has a destructive effect on the pancreas and can lead to insulin deficiency and development of T1D [24].If the hypothesis that SARS-CoV-2 infection causes hyperglycemia is true, increased statistics of new T1D cases should be observed. Indeed, there are publications that describe such a phenomenon. For instance, Unsworth et al. and Kamrath et al. describe an increase in new-onset T1D in children during the COVID-19 pandemic [16,25]. Although pancreatic β cell damage induced transient hyperglycemia in SARS-CoV, it is still unclear whether β cell damage is transient or permanent in SARS-CoV-2 [22]. This information appears to be of great importance because COVID-19 in children is frequently considered “harmless”. Therefore, it is reasonable to sensitize parents to the fact that the consequences of COVID-19 may be potentially dangerous for their children.Below you will find the proposed molecular mechanisms that may participate in pancreatic damage that causes carbohydrate metabolism disorders.

4. Etiology Associated with ACE2, TMPRSS2, and Na+/H+ Exchanger

As previously mentioned, SARS-CoV infection of host cells is facilitated by ACE2, but also by the transmembrane protease serine 2 (TMPRSS2) and other host cell proteases such as cathepsin L (CTSL) [13].ACE2 is an enzyme that is expressed to varying degrees in most cells of the human body [14,26,27]. This enzyme catalyzes the conversion of angiotensin II to angiotensin 1–7, taking part in the maintenance of body homeostasis by influencing the regulation of blood pressure and water–electrolyte balance through the renin–angiotensin–aldosterone (RAA) system [28]. Moreover, ACE2/angiotensin (1–7) stimulates insulin secretion, reduces insulin resistance, and increases pancreatic βcell survival [27,28].In addition to the key role it plays in maintaining body homeostasis, ACE2 is now also the best-studied target for SARS-CoV-2 S glycoprotein, enabling infection of host cells [27,29]. ACE2 in the pancreas is expressed mainly within the pericytes of pancreatic microvessels and to a lesser extent on the surface of the islets of Langerhans, including pancreatic β cells [30]. SARS-CoV-2 shows 10–20 times more activity against ACE2 than SARS-CoV, which significantly increases the infectivity of SARS-CoV-2 [31,32]. Furthermore, studies indicate that SARS-CoV may also downregulate ACE2 expression in cells. This causes an imbalance between ACE and ACE2, consequently leading to blood pressure disorders and systemic inflammation [27,33,34]. Due to the 79% genetic similarity between SARS-CoV and SARS-CoV-2 [35], it is speculated that ACE2 expression may also be downregulated during SARS-CoV-2 infection, causing i.a. MODS observed in COVID-19 [27].During cell infection by SARS-CoV-2, in addition to the role played by ACE2, it is also appropriate to consider the significant pathogenic role of TMPRSS2 that is necessary for the preparation of S glycoprotein by its cleavage, thereby enabling fusion of the virus with the host cell [36,37]. The S1 and S2 domains can be distinguished in the SARS-CoV-2 S glycoprotein. The S1 domain is involved in binding to the ACE2 receptor and then TMPRSS2 intersects with the S protein, including at the boundary of the S1 and S2 domains and within the S2 domain, which enables the virus–cell fusion [38,39]. According to studies, TMPRSS2 expression is significantly increased in obese patients, which may contribute to the poorer prognosis that is observed during COVID-19 in this patient group [40]. Moreover, obese patients are frequently already burdened with problems such as insulin resistance at baseline, while the presence of ACE2 and TMPRSS2 within the pancreas as a binding site for SARS-CoV-2 may exacerbate insulin resistance causing problems in terms of diabetes management in COVID-19 patients.There are also other mechanisms by which COVID-19 may affect the development of hyperglycemia. It is reported that the virus may also affect the glucose regulation through the Na+/H+ exchanger and lactate pathways. The mechanism is that angiotensin II, which accumulates during infection, contributes to insulin resistance and—by activating the Na+/H+ exchanger in the pancreas—it leads to hypoxia and extracellular acidification, which, through the accumulation of calcium and sodium ions in the cells and the production of reactive oxygen species, damages pancreatic tissue [41]. Simultaneously, the concentration of lactate increases, which in COVID-19 infection is intensively released, among other things, from adipose tissue, and then monocarboxylate transporters transport lactate and H+ ion inward in the cell, which increases Na+/H+ exchanger activation, further disrupting pancreatic homeostasis [41].

5. The Etiology Associated with a Systemic Proinflammatory Environment, Immune System Aggression, and Production of Novel Autoantigens

A broad spectrum of proinflammatory cytokines, such as IL-2, IL-6, IL-7, IL-8, interferon-γ, and Tumor Necrosis Factor α (TNF-α), is released during, in particular severe, COVID-19 infection [42,43,44]. Based on current studies, it is reasonable to suspect that these cytokines are released in response to the binding of the virus to ACE2 receptors that are also located in the pancreas [9,42]. The cause of pancreatic damage during COVID-19 is the cytokine storm that plays a key role in this case, because in both acute pancreatitis (AP) and severe COVID-19, elevated levels of the aforementioned interleukins are associated with the severity of these both disease entities. Particular attention should be paid to IL-6, because it is suspected to play a key role in the pathogenesis of AP as well as acute respiratory distress syndrome (ARDS) that is the most common and most severe clinical manifestation of COVID-19. In COVID-19-induced ARDS, IL-6 levels are correlated with disease-related mortality [45,46,47]. At the same time, high IL-6 levels correlate with an increased risk of developing severe pancreatitis [48,49].The production of neutralizing antibodies is also an important response of the body in the course of COVID-19 [50,51,52]. It has been observed that early seroconversion and very high antibody titers occur in patients with severe SARS-CoV-2 infection [53,54]. The available literature details a mechanism called antibody-dependent enhancement (ADE), which is associated with a pathological response of the immune system [53]. ADE exploits the existence of FcRS receptors located on various cells of the immune system, for example, macrophages and B lymphocytes [53]. This relationship may lead to a likely bypass of the classical viral infection pathway by ACE2, and virus–antibody complexes may stimulate macrophages to overproduce cytokines including significant IL-6 [53,55].Molecular mimicry may be also one of potential causes of pancreatic cell damage [56]. There are similarities in the protein structure of the virus and β-pancreatic cells, which may induce cross-reactivity and lead to autoimmunity [56]. Furthermore, viral infection may also lead to increased cytokine secretion by surrounding dendritic cells and activation of naive T cells in genetically predisposed individuals [56].

6. Pancreatitis in COVID-19

Although the impact of the discussed coronavirus-induced disease on exocrine function is not fully understood, available literature is not able to unambiguously determine whether the tissue damage leading to AP occurs as a result of direct SARS-CoV-2 infection [57] or as a result of systemic MODS with increased levels of amylase and lipase [42]. Liu et al.’s study involving 121 COVID-19 patients with a mean age of 57 years and a variable course of infection proved above-normal levels of amylase and lipase in 1–2% of patients with moderate COVID-19 infection and in 17% of patients with severe COVID-19 infection. This may support the hypothesis that SARS-CoV-2-induced disease has a destructive effect not only on the endocrine portion of this gland, but also on the exocrine one [15].However, elevated levels of pancreatic enzymes in question do not have to mean the destruction of pancreatic cells—after all, such a situation may occur during kidney failure or diarrhea in the course of COVID-19. Furthermore, there remains the question of the effect of drugs administered during SARS-CoV-2 infection on changes in pancreatic function [42], discussed further in this article.According to the International Association of Pancreatology (IAP) and the American Pancreatic Association (APA), the diagnosis of AP is based on meeting two out of three of the following criteria: clinical (epigastric pain), laboratory (serum amylase or lipase > 3 × upper limit of normal), and/or imaging criteria (computed tomography, magnetic resonance imaging, ultrasound) [58]. Pancreatic lipase is considered as a potential marker of SARS-CoV-2 severity with concomitant AP. In Hemant Goyal et al.’s study, as many as 11.7% out of 756 COVID-19 patients had hyperlipidemia and they were three times more likely to have severe COVID-19 [59]. Those with higher lipase levels—17% out of 83 patients—required hospitalization [60]. However, it is difficult to distinguish whether these patients required hospitalization for severe systemic COVID-19 infection or for pancreatitis in the course of COVID-19 infection.AP in the course of COVID-19 was analyzed in different age groups; however, some studies only involve children [61]. Compared to pancreatic islet cells, cells of the exocrine pancreatic ducts are more abundant in ACE2 and TMPRSS2 that are necessary for the virus to penetrate the cell [62]. Infection of these cells may be one of the causes of AP [63]. Infections, both bacterial and viral, are one of the causes of AP. The definitive mechanism of how viral infections affect pancreatic cells is not known; however, a study by Maria K Smatti et al. found that there is infection of pancreatic islet cells and replication of the virus within them, ultimately resulting in autoimmune reactions that eventually affect both diabetes and AP in a negative way [64]. For non-SARS-CoV-2 patients, the etiology of AP is known and confirmed in most cases, although 69% of those undergoing infection do not have definite etiology of AP while meeting the AP-Atlanta criteria for diagnosis [65].Hegyi et al. show the mechanism of MODS formation during COVID-19 infection and AP [66]. This is lipotoxicity, involving an interstitial increase in pancreatic lipase levels, which leads to the breakdown of triacylglycerols contained in adipose tissue cells and the release of unsaturated fatty acids. These in turn exert a toxic effect on mitochondria causing the release of cytokines, which results in a cytokine storm.There is also a hypothesis, which claims that AP can develop because of blood circulatory centralization resulting from uncontrolled cytokine storm created by SARS-CoV-2 infection [67]. There exist reports that say that pancreatic ischemia may be the cause of different degrees of acute pancreatitis [68,69]. This statement can be supported by the reports that state that pancreatic blood reperfusion inhibits the development of AP and accelerate pancreas recovery [70].Another mechanism of developing AP during COVID-19 may be a coagulation cascade activation caused by active inflammatory process due to SARS-CoV-2 infection [71]. The ongoing inflammatory process causes not only hemostasis imbalance for blood clotting, but it also leads to intensification of coagulation by removing epithelial cell protein C receptor (EPCR) from epithelial by the means of inflammatory mediators and thrombin [71]. This means that both processes intensify each other. Simultaneously, it was proved that COVID-19 predisposes patients to venous thromboembolism resulting from excessive inflammation, platelet activation, and endothelial dysfunction [72]. It is also important to notice that AP is inherently connected with a coagulation cascade activation, increased fibrinolysis and, hence, higher level of D-dimers [73]. Acute pancreatitis severity may depend on hemostasis imbalance; local coagulation results in mild AP whereas, in more severe AP cases, the imbalance may lead to development of disseminated intravascular coagulation (DIC) [74]. These observations have been supported by the results of experimental studies showing that the inhibition of coagulation reduces the development of AP [75,76,77] and exhibits therapeutic effect in this disease [78,79]. Additionally it is worth noticing that infection-related hyperglycemia has powerful inflammation-promoting effects on the organism (especially when organism is under stress), thus increasing the number of inflammatory mediators [74]. Unfortunately, it is impossible to decide which process is dominant in causing AP in COVID-19 patients: local inflammation caused by SARS-CoV-2 or systemic hemostasis imbalance.Clinical reports on low molecular weight heparin (LMWH) treatment in AP seem to emphasize a more significant role of hemostasis imbalance in causing AP [74,80,81]. Heparin is extremely significant in the treatment of COVID19 patients due to its properties, mainly its similarity to heparan sulphate, which appears in a respiratory tract, its interactions with SARS-CoV-2 S protein, leading to viral adhesion inhibiting to the cell membrane [82], and its anti-inflammatory effects. Thanks to these properties, heparin may not only show its therapeutic effect as the anticoagulant, but also its protective role in acute pancreatitis or respiratory inflammations [83,84,85].

7. Drugs Used against SARS-CoV-2 Infection (Glucocorticoids, Lopinavir, Ritonavir, Remedesivir, Interferon-β1 (IFN-β1), and Azithromycin) Induce Pancreatic β Cell Damage

Statistical analyses revealed a significantly higher incidence of AP with the concomitant systemic use of glucocorticosteroids (GCS) [86]. In one study analyzing the development of drug-induced AP, dexamethasone, was classified as type IB—there was one case report in which administration of this drug-induced AP occurred; however, other causes of pancreatitis such as alcohol consumption could not be excluded [87]. Other GCS such as hydrocortisone, prednisone, and prednisolone were used in patients with mild to moderate AP; however, they cannot be classified into any group because they are frequently used together with other drugs that cause AP [86,87]. However, it has been determined that GCS independently increase the risk of AP, and patients with residual AP risk factors during GCS treatment should be more monitored for the development of AP [23]. Javier A. Cienfuegos et al. additionally observed that one of mechanisms of AP formation in COVID-19 patients may be GCS administered at the time of admission to the ICU with severe respiratory failure [88]. Because GCS were used in severe COVID-19 cases, it is difficult to say what true reason for AP was—either a severe course of COVID-19 or GCS application or both.GCS are used in the treatment of many diseases due to their immunosuppressive and anti-inflammatory nature. They induce diabetes in previously healthy patients as well as significantly exacerbate diabetes in diabetic patients [89,90]. Diabetes develops in these patients likely due to pancreatic β cell dysfunction, decreased insulin secretion, and increased insulin resistance in other tissues, which may depend on the timing and the dose of GCS used [89,91]. Long-acting or intermediate-acting insulin alone or combined with short-acting insulin should be used during the treatment [90]. At the same time, no advantage was found over the use of oral hypoglycemics [92]. Certainly, patients after long-term GCS therapy will need further observation for diabetes.Lopinavir/ritonavir was classified in the previously mentioned study as a type IV drug—medications reported with little information [87]. Both drugs are included in the group of antiretrovirals that act as protease inhibitors, and they are primarily used for HIV infection. Although Lopinavir is an active drug, it is not used alone. There have been reports about the occurrence of AP during the use of protease inhibitors in question, which is also described in the Summary of Product Characteristics (SmPC) of products approved by Committee for Medicinal Products for Human Use (CHMP). It has been proved that the use of lopinavir/ritonavir causes hyperglycemia [93,94].Remdesivir is an adenosine analogue with antiviral activity. There are single reports about the occurrence of pancreatitis as a result of the use of the aforementioned medication [95,96]. At the same time, it should be noted that other nucleoside-derivative drugs may cause pancreatitis [97].The current state of knowledge does not clearly indicate the therapeutic benefit of interferon-β in the treatment of COVID-19 patients [98,99]. To date, only single cases suggesting induction of pancreatitis by interferon-β have been reported. Based on this, Badalov et al. classified interferon into type III [87].There are few reports about the development of AP due to the use of azithromycin [100]. In the previously mentioned study by Badalov et al., two macrolide antibiotics were classified as type II and III. Unfortunately, there are no direct data concerning azithromycin. Interestingly, there were cases of patients with concomitant symptoms of AP and viral pneumonia caused by SARS-CoV-2 who were treated with azithromycin, which resulted in complete resolution of symptoms for both conditions [96,101]. Based on available data, the risk of azithromycin-induced AP is low.There is no clear evidence that azithromycin affects blood glucose levels in humans. However, it is known for its prokinetic effects, which may be helpful in patients who suffer from diabetic gastroparesis [102]). The incidence of hypo- and hyperglycemic episodes was not proved to be significant for azithromycin [103]; however, the risk of dysglycemia is emphasized [94]. In the SmPC, where azithromycin is the main ingredient, it is not possible to establish a causal relationship between the occurrence of pancreatitis and taking medications (Zithromax) based on the available data. In contrast, glycemic disturbances were not indicated as side effects (Zithromax) [104].Hydroxychloroquine has been extensively promoted for COVID-19 due to its anti-inflammatory and antiviral action; yet, the use of this agent in diabetes deserves particular attention for its documented hypoglycemic action, and its benefit on COVID-19 is controversial, although there is large usage [105].Table 2 shows a comparison of the side effects of medications in question.Table 2. Side effects of medications used in SARS-CoV-2 infection in the area of pancreatic effects and hyperglycemia.

Table

8. COVID-19, Pancreas, and Glycation

In T2D diabetics, oxidative stress leading to pancreatic damage may be stimulated by, among other things, the intense glycation that accompanies hyperglycemia [24]. Glycation is a non-enzymatic process involving reducing sugar and amino groups of proteins, which contributes to the formation of advanced glycation end products (AGEs). These products have significantly altered biochemical properties relative to the substrates, including proteins that have altered conformation, increased rigidity, resistance to proteolysis, etc. [106,107].Part of the pathomechanism involved in facilitating coronavirus infection in diabetics may be due to glycation of ACE2 and SARS-CoV-2 spike protein [108,109].An interesting hypothesis is that COVID-19 has a worse prognosis in patients with intense glycation, and thus high tissue AGE content. Glycated hemoglobin (HbA1c) is a commonly used diagnostic tool that estimates intensity of glycation. The parameter is not only a marker of long-term persistent hyperglycemia, but an active participant in immune processes, as HbA1c levels are associated with NK cell activity [110].Zhang et al.’s retrospective cohort study concerning COVID-19 patients revealed that glycated hemoglobin correlates negatively with saturation (SaO2) and positively with C-reactive protein (CRP), erythrocyte sedimentation rate (ESR), and fibrinogen (Fbg). It was concluded that determination of HbA1c levels may be helpful in assessing inflammation, hypercoagulability, and prognosis of COVID-19 patients [111].According to the meta-analysis by Chen et al. (2020), Hba1c levels were slightly higher in patients with severe COVID-19 compared to patients with mild COVID-19; however, this correlation was not statistically significant. However, it is of great importance to note that only two studies analyzing HbA1c in COVID-19 patients were included in this analysis because only these studies were available in May 2020 [112].Glycation plays its physiological effects not only directly by changing the properties of various proteins, but also indirectly through various receptors. RAGE is the most common receptor for AGEs. Binding of RAGE to its ligands activates a proinflammatory response primarily by mitogen-activated protein kinase (MAPK) and nuclear factor κβ (NFκβ) pathways. This interaction was proved to be significant in the pathogenesis of cancer, diabetes mellitus, and other inflammatory disorders [113]. RAGE was found to be expressed in the pancreas, and S100P-derived RAGE antagonistic peptide (RAP) reduces pancreatic tumor growth and metastasis [113]. The implications of this fact may also apply to the etiology and treatment of COVID-19. It has been postulated that targeting RAGE by various antagonists of this receptor may inhibit damage to various organs including the pancreas [114].

9. COVID-19 vs. Pancreatic Cancer

Immunosuppression as a treatment effect, elevated cytokine levels, altered expression of receptors for SARS-CoV-2, and a prothrombotic state in patients with various types of cancer may exacerbate the effects of COVID-19 [115].Focusing on pancreatic cancer, it can be observed that the pathomechanism of both diseases—COVID-19 and tumorigenesis in the pancreas—overlap in several molecular mechanisms. As mentioned above, SARS-CoV-2 infection of host cells is facilitated by ACE-2, TMPRSS2, and CTSL. Cathepsin L is upregulated in a wide variety of cancers, including pancreatic adenocarcinoma [13]. TMPRSS2 upregulation in pancreatic cancers is moderate, whereas ACE-2 is overexpressed in some cancers, including pancreatic carcinomas [115]. Interestingly, ACE2 upregulation seems to be associated with favorable survival in pancreatic cancer [116], and it is known that SARS-CoV-2 reduces ACE2 expression [22]. Furthermore, the above-mentioned RAGE may also participate in both pancreatic cancer development and SARS-CoV-2 infection. RAGE facilitates neutrophil extracellular trap (NET) formation in pancreatic cancer [117]. In conclusion, pancreatic cancer predisposes to an increased risk of COVID-19 and its more severe course, and coronavirus infection may contribute to pancreatic cancer.It also seems important how the COVID-19 epidemic has affected the treatment of patients with pancreatic cancer of SARS-CoV-2-independent etiology. According to the study by Pergolini et al., care of patients with pancreatic cancer can be disrupted or delayed, particularly in the context of treatment selection, postoperative course, and outpatient care [118].A separate issue is how patients after pancreatoduodenectomy respond to SARS-CoV-2 infection. A case series reported by Bacalbasa reveal that patients who develop SARS-CoV-2 infection postoperatively require re-admission in the ICU and a longer hospital stay; however, these infections are not fatal [119]. Although the analysis was performed on single cases, it is concluded that these results are an argument to perform elective oncological surgeries [119].There are also reports that chemotherapy in pancreatic cancer patients who become ill between treatment series can be successfully completed after a complete cure of the infection [120]. Guidelines for, e.g., prioritization and treatment regimens regarding pancreatic cancer treatment in the era of the pandemic, are developed and described, for example, by Catanese et al. or Jones et al. [121,122].

10. Conclusions

Evidence shows that SARS-CoV-2 infection contributes to damage within the pancreas. The mechanisms that are involved in this include but are not limited to direct cytopathic effect of SARS-CoV-2 replication and systemic and local inflammatory response [123]. At the current state of knowledge, it is certain that the virus attacks the endocrine portion of the pancreas as well as, to a much lesser extent, the exocrine portion. It has been shown that a bidirectional relationship between COVID-19 and diabetes exists; indeed, diabetes is associated with COVID-19 severity and mortality but, at the same time, patients with COVID-19 have shown new onset of diabetes [124]. SARS-CoV-2 virus infection not only directly affects glycemic levels, but also exacerbates already existing hyperglycemia through its negative impact on the functional competence of the islets of Langerhans. It cannot be excluded that the real cause of exocrine dysfunction of this gland is the negative effect of the drugs used for treatment of the infection. As the pandemic progresses, special attention should be given to the evaluation of chronic and acute pancreatic diseases, including pancreatic cancer, so that faster diagnosis enables faster implementation of treatment.

Author Contributions

Conceptualization, A.K.; investigation, U.A., M.N., A.S., P.W., P.Z. and A.K.; resources, U.A., M.N., A.S., P.W., P.Z. and A.K.; writing—original draft preparation, U.A., M.N., A.S., P.W., P.Z. and A.K.; visualization, U.A.; supervision, A.K. All authors have read and agreed to the published version of the manuscript.

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ACE2-independent infection of T lymphocytes by SARS-CoV-2

Authors: Xu-Rui ShenRong GengQian LiYing ChenShu-Fen LiQi WangJuan MinYong YangBei LiYong YangBei LiRen-Di JiangXi WangXiao-Shuang ZhengYan ZhuJing-Kun JiaXing-Lou YangMei-Qin LiuQian-Chun GongYu-Lan ZhangZhen-Qiong GuanHui-Ling LiZhen-Hua ZhengZheng-Li ShiHui-Lan ZhangKe Peng & Peng Zhou 

Signal Transduction and Targeted Therapy volume 7, Article number: 83 (2022) 

Abstract

SARS-CoV-2 induced marked lymphopenia in severe patients with COVID-19. However, whether lymphocytes are targets of viral infection is yet to be determined, although SARS-CoV-2 RNA or antigen has been identified in T cells from patients. Here, we confirmed that SARS-CoV-2 viral antigen could be detected in patient peripheral blood cells (PBCs) or postmortem lung T cells, and the infectious virus could also be detected from viral antigen-positive PBCs. We next prove that SARS-CoV-2 infects T lymphocytes, preferably activated CD4 + T cells in vitro. Upon infection, viral RNA, subgenomic RNA, viral protein or viral particle can be detected in the T cells. Furthermore, we show that the infection is spike-ACE2/TMPRSS2-independent through using ACE2 knockdown or receptor blocking experiments. Next, we demonstrate that viral antigen-positive T cells from patient undergone pronounced apoptosis. In vitro infection of T cells induced cell death that is likely in mitochondria ROS-HIF-1a-dependent pathways. Finally, we demonstrated that LFA-1, the protein exclusively expresses in multiple leukocytes, is more likely the entry molecule that mediated SARS-CoV-2 infection in T cells, compared to a list of other known receptors. Collectively, this work confirmed a SARS-CoV-2 infection of T cells, in a spike-ACE2-independent manner, which shed novel insights into the underlying mechanisms of SARS-CoV-2-induced lymphopenia in COVID-19 patients.

Introduction

Since its emergence in December 2019, SARS-CoV-2, the etiology of coronavirus disease 2019 (COVID-19), quickly spread to the majority of countries in the world and posed great threats to public health. The virus shares 79.5% genome identity with SARS-CoV-1 and also uses angiotensin-converting enzyme 2 (ACE2) as a cell entry receptor.1,2,3,4,5 Typical clinical symptoms of COVID-19 patients include fever, fatigue, dry cough, and pneumonia, whereas around 20% of the severe cases may die of multi-organ failure.6,7,8,9

Apart from the respiratory system, multiple organs including the immune system of COVID-19 patients were also targeted by SARS-CoV-2 infection. Notably, lymphopenia was observed in 83.2% of the patients on admission, and fatal infections were associated with more severe lymphopenia over time.6,7,8 Lymphocytes (particularly T cells) play a central role in the human immune system, a decrease of which would result in immune suppression and serious complications.10 It has been proposed that viral-induced lymphopenia might be due to direct infection, cytokine-mediated cell death, tissue sequestration of lymphocytes, or suppression of the bone marrow or thymus for T-cell generation.11 In the case of MERS-CoV, apoptosis induced by direct viral infection of T cells has been observed in vitro, which possibly explained lymphopenia in MERS patients.11 SARS-CoV-1 viral particles were also observed in multiple leukocytes from an autopsy study, suggesting that direct infection might account for the decrease in lymphocytes.12 Similarly, SARS-CoV-2 particles or proteins were also found in the spleen and lymph nodes from a study of 91 deceased COVID-19 cases, suggesting an infection of lymphocytes.13 Furthermore, in COVID-19 immune landscape depicted by single-cell RNA-seq studies, SARS-CoV-2 viral RNA has been found in multiple immune cells, including myeloid cells with phagocytic activity (neutrophil and macrophage) and lymphocytes without phagocytic activity (T, B, and NK cells).14,15 Notably, SARS-CoV-2 RNA-positive immune cells did not co-express the entry factors ACE2 and TMPRSS2, or other hypothesized entry co-factors.14,15 It is speculated that cell-associated SARS-CoV-2 viral positivity may represent a mixture of replicating virus, immune cell engulfment, and virions or virally infected cells attached to the cell surface.14,15

It has been shown that SARS-CoV-2-infected human monocytes, monocyte-derived macrophages, and dendritic cells in vitro, which potentially plays a major role in COVID-19 pathogenesis.16,17 However, whether SARS-CoV-2 infects lymphocytes, which do not express ACE2, to result in lymphopenia is still unknown. This knowledge gap also brings difficulty for our understanding of how lymphocytes lost the ability to control viral infection. Here, we provided evidence that activated T lymphocytes could be infected by SARS-CoV-2 in an ACE2-independent manner. The infection leads to pronounced T-cell apoptosis in vitro or in patients with COVID-19. Our findings shed light on the understanding of SARS-CoV-2 infection-induced lymphopenia.

Results

Presence of SARS-CoV-2 in lymphocytes from patients with COVID-19

Multiple immune cell types, including lymphocytes, have been shown enriched for SARS-CoV-2 viral RNA in multiple single-cell RNA-seq studies.14,15 To determine whether SARS-CoV-2 infects lymphocytes, we analyzed peripheral blood cells (PBCs) collected from COVID-19 patients. PBCs were prepared from 22 patients, who were all at severe condition during the study along with 15 healthy donors. We first analyzed major lymphocyte cell types including T (CD4 + helper T and CD8 + cytotoxic T), B, and natural killer (NK) cells for their population changes or the presence of viral antigen upon infection. For all patients tested, the ratios of blood T lymphocytes declined significantly compared to those in healthy donors, whereas B and NK cells appeared to be unaffected (Fig. 1a). Notably, CD4 + and CD8 + T lymphocytes almost declined to zero in some patients (Fig. 1b). The results suggested that lymphopenia in these patients is likely attributed to a decline of T lymphocytes.

figure 1
Fig. 1

We then analyzed the presence of SARS-CoV-2 viral antigens in PBCs using flow cytometry or by immunofluorescence assay (IFA). The results suggested that T lymphocytes were infected and in certain patient CD4 + T cells showed a high infection rate (Supplementary Fig. S1a). We also confirmed the presence of viral antigen in T lymphocytes from patient blood by immunofluorescence analysis (IFA) (Fig. 1c). Furthermore, we prepared postmortem lung sections from patients with a fatal infection and analyzed T lymphocytes infiltration and virus infection. We found T lymphocytes infiltration in the lung section, and many T lymphocytes were also positive for SARS-CoV-2 NP staining, indicating virus infection (Fig. 1d). A similar finding has also been reported.13 Taken together, we showed the presence of SARS-CoV-2 viral antigen in T lymphocytes either in the blood or in the lung section from the COVID-19 patients.

To further corroborate these findings, virus isolation was attempted from viral NP-positive PBCs. Patient PBCs were collected, determined for viral antigen using flow cytometry, and then co-cultured with Caco2 cells after three washes. Positive detection of viral RNA in the supernatant or viral protein in the Caco2 cells after co-culture indicated successful isolation and amplification of SARS-CoV-2 from PBCs of some COVID-19 patients (3 out of 5) but not from the healthy control (Supplementary Fig. S1b–e). Notably, in the three viral isolation positive samples, two also showed viral positive in the flow cytometry assay (P2 and P4), while the third one (P5) likely carried infectious virus at a level that was under the detection limit of flow cytometric analysis. Above all, we observed SAR-CoV-2 viral RNA and viral protein, and likely infectious virus in T lymphocytes from COVID-19 patients.

SARS-CoV-2 infection of T cells in vitro

Since T lymphocytes population decreased in COVID-19 patients and CD4 + T lymphocytes showed a high viral antigen-positive rate, we then investigated whether SARS-CoV-2 infects CD4 + T lymphocytes. For this purpose, we conducted a serial of experiments to test whether SARS-CoV-2 infects T cells. Upon infection, both viral RNA detection targeting at the receptor-binding domain (RBD) and viral subgenomic mRNA (sgRNA) targeting at M gene were tested. Viral sgRNA is transcribed only in infected cells during viral replication and is not packaged into virions, and therefore indicates the presence of actively infected cells in samples. Viral nucleocapsid protein (NP) and viral particles were also detected using western blot (WB), flow or electron microscopy (EM). Jurkat or MT4 cells, two commonly used CD4 + T cell lines, and primary T cells isolated from healthy donors were infected with SARS-CoV-2 (Fig. 2a). In some experiments, T cells were also activated by Phorbol myristate acetate (PMA) for 2 h for Jurkat cells or by a combination of IL2 + CD3 + CD28 for 3 days for primary T cells before infection, considering a large proportion of T cells is activated in human (Supplementary Fig. S2).

figure 2
Fig. 2

At 0, 24, 48, and 72 h post infection, it was observed that SARS-CoV-2-infected Jurkat T-cell line in a time-dependent manner, and the infection was more robust in activated T cells. Accumulation of viral RNA and sgRNA in cells or viral RNA in the culture supernatant was observed (Fig. 2b). Next, we sought to determine whether the qPCR detection assay represents only partial viral genome replication. We performed RNA-seq analysis of the SARS-CoV-2-infected activated Jurkat T cells at 0 or 24 h p.i. and analyzed the viral reads depth and coverage across the viral genome. Compared to 0 h infected, a much higher depth of viral genomes (as high as 5000 reads depth) can be observed in the 24 h-infected cells, demonstrating an effective replication (Fig. 2c). We then determined viral antigens by WB and flow assay. Our results showed a time-dependent increased level of viral NP in cells or in the supernatant, similar to the findings in viral RNA detection (Fig. 2d, e). We further employed electron microscopy to analyze SARS-CoV-2 infection of T-cell lines. Activated Jurkat or MT4 cells were infected with SARS-CoV-2 for 72 h and viral particles with typical coronavirus morphology were observed in the cytoplasm of the infected cells (Fig. 2f). Finally, to corroborate the findings from T-cell lines, we tested the infectivity of primary T cells isolated from healthy donors. In the three donors, SARS-CoV-2 showed time-dependent infection of T cells that is peaked at 8 h, probably because of extensive cell death induced by the virus at this time point (discussed below). Activation sensitized the cells to SARS-CoV-2 infection in two of the three donors. As comparison, primary colon organoid was also infected, which showed much higher infection efficiency compared to T cells (Fig. 2g, h). Taken together, our data clearly show that SARS-CoV-2 could infect T cells in vitro, although at a lower efficiency compared to tissue cells.

SARS-CoV-2 infection of T cells is ACE2 and TMPRSS2-independent

It is generally believed that ACE2 is the entry receptor for SARS-CoV-2. However, major cell populations in PBCs express extremely low levels of ACE2, raising the question whether ACE2 also mediates SARS-CoV-2 virus entry of T cells. We first tested whether an ACE2 knockdown could dampen SARS-CoV-2 infection of T cells. The data showed ACE2 was successfully knocked down by ACE2-shRNAs in Caco2 cells. Jurkat T cells do not express detectable ACE2 under either mock or knocked down conditions (Fig. 3a). Correspondingly, ACE2 knockdown resulted in dramatically decreased SARS-CoV-2 infection in Caco2 cells but not in Jurkat T cells (Fig. 3b). To further confirm this finding, we did ACE2 knocked out in Caco2 and Jurkat cells (Fig. 3c). Similar to ACE2-knockdown cells, viral load decreased in Caco2-ACE2-KO cells but not in Jurkat-ACE2-KO cells (Fig. 3d). These results suggested that SARS-CoV-2-infected T cells in an ACE2-independent manner.

figure 3
Fig. 3

It was reported that soluble human ACE2 protein could block SARS-CoV-2 infection through competing virus binding with the cellular receptor.3 Thus, ACE2 antibody pre-incubated cells or spike antibody pre-incubated SARS-CoV-2 should also block viral infection, if the infection depends on spike-ACE2 binding. To analyze whether these molecules affect SARS-CoV-2 infection of T cells, we incubated virus with soluble human ACE2 protein or a commercial mAb targeting at RBD-ACE2 binding, or incubated cells with ACE2 blocking antibody before the infection of Caco2 or activated Jurkat T cells. The intracellular viral RNA was analyzed after infection. In Caco2, the three blockers strongly blocked SARS-CoV-2 infection, and ACE2 protein appears to be more potent than the other two treatments. In contrast, none of the three treatments affected the SARS-CoV-2 infection of Jurkat T cells (Fig. 3e).

Lastly, it is known that SARS-CoV-2 uses the serine protease TMPRSS2 for S protein priming before binding to ACE2 receptor, and a TMPRSS2 inhibitor has been approved for clinical use (Camostat) to block SARS-CoV-2 entry.1 The RNA expression of TMPRSS2 in Caco2, Jurkat, and activated Jurkat cells was determined by qPCR. The result suggested that neither unactivated nor activated Jurkat cell-expressed TMPRSS2 (Fig. 3f). We observed that Camostat inhibited SARS-CoV-2 infection of Caco2 cells in a dose-dependent manner. At a dose of 20 μm, Camostat almost completely blocked viral infection of Caco2 cells. In contrast, Camostat showed no inhibitory effect on SARS-CoV-2 infection of Jurkat T cells even at a high dose (Fig. 3g). Collectively, these results suggested that SARS-CoV-2 infection of T cells does not rely on the spike-ACE2/TMPRSS2 interaction.

SARS-CoV-2 infection triggered T-cell death

It is known that severe patients with COVID-19 showed marked decreased lymphocyte populations. To determine whether SARS-CoV-2 infection contributes to T-cell death, we tested PBC T lymphocytes apoptosis collected from patients with COVID-19. T lymphocytes from patients or from healthy donors were dual-labeled with a CD3 antibody and a viral NP antibody, and apoptosis was analyzed with the TUNEL assay. T lymphocytes from COVID-19 patients underwent pronounced apoptosis compared to those from the healthy donors, showing a more than tenfold increase of apoptotic cells. In some patients, most of the apoptotic cells were also viral antigen-positive (e.g., 65% in patient 1), suggesting viral infection played a role in peripheral blood T lymphocytes death in these patients (Fig. 4a).

figure 4
Fig. 4

To confirm the role of viral infection in T-cell death, we experimentally infected primary T cells isolated from healthy donors. With or without activation, cells were experimentally infected with SARS-CoV-2 for 8 h and apoptosis was analyzed with TUNEL assay. It can be observed that SARS-CoV-2 infection induced pronounced apoptosis in infected T cells compared with the mock-treated cells. Activation sensitized T cell to viral infection, as shown by higher apoptotic cells in the activated group (Fig. 4b).

Finally, we determined the cellular responses in T cells upon SARS-CoV-2 infection by bulk RNA-seq analysis. Activated Jurkat T cells were infected with SARS-CoV-2 for 0, 24, 48, and 72 h before they were collected for TUNEL assay. It can be observed that virus induced significant apoptosis at 72 h post infection, compared to mock-infected or cells at other time points (Fig. 4c). We then determined the dynamic cellular responses in cells that have been infected for 24 or 48 h, as the cells in 72 h groups contained too many dead cells and were not suitable for RNA-seq analysis. Compared to the 24 h group, the hypoxia-related GO pathways are significantly upregulated in 48 h group, including “PID HIF1 TF pathway”, “response to hypoxia”, “positive regulation of cell death”, and “intrinsic apoptotic signaling pathway”. It has been shown that SARS-CoV-2 infection triggers mitochondrial ROS production, which induces stabilization of hypoxia-inducible factor-1a (HIF-1a) in monocytes.16 Similarly in T cells, multiple genes involved in this oxidative stress response were upregulated: BNIP3, PFKFB3, FOS, JUN, BHLHE40, GADD45B, PDK1, and DDIT4 (Fig. 4d). To corroborate the findings in T cell lines, we conducted RNA-seq analysis to primary peripheral blood mononuclear cells (PBMCs) collected from three healthy donors and three severe COVID-19 patients. Our data showed an upregulation of cell responses to stimuli, cell death, or response to hypoxia pathways, and a down-regulation of leukocytes activation and signaling pathways, similar to the findings in the T-cell line (Fig. 4e). In summary, SARS-CoV-2 infection induced pronounced T-cell death, which is probably dependent on mitochondria ROS-hypoxia pathways.

Exploration of potential receptors in T cells

Since our results suggested that the infection of SARS-CoV-2 to Jurkat T cell is ACE2-independent, we tried to identify the potential receptors. We first explored the expression of the known SARS-CoV-2 receptors or co-factors that have been identified in primary T cells from public single-cell NGS data14 and in Jurkat T cells in RNA-seq analysis with or without activation, including ACE2/TMPRSS2, AXL, NRP1, KIM-1/TIM-1, ASGR1, and KREMEN1.18,19,20 Moreover, ITGB2 (leukocyte-associated molecule-1, LFA-1), the leukocyte cell Adhesion molecule, has been suggested binding to SARS-CoV-1 ORF7a.21 As SARS-CoV-2 shares similar ORF7a as SARS-CoV-1, it would be interesting to evaluate whether LFA-1 also mediated SARS-CoV-2 infection of T cells.

Our data showed minimal expression of the following molecules in SARS-CoV-2-positive T cells from patients: ACE2, TMPRSS2, ASGR1, KREMEN1, and NRP1 (Fig. 5a and Supplementary Fig. S3a). In contrast, AXL and LFA-1 were expressed in these cells. In Jurkat cells, LFA-1 also showed very high expression, although it was not upregulated following a 2 h activation (Supplementary Fig. S3b). Taken together, AXL and LFA-1 appeared to be promising targets as entry molecules.

figure 5
Fig. 5

AXL was proposed to be a candidate receptor for SARS-CoV-2 in a previous study and the function in mediating SARS-CoV-2 infection is independent of ACE2.19 BEAS-2B that was used as a positive control for AXL-SARS-CoV-2 studies was pretreated with AXL proteins of different concentrations (25, 50, 100 μg/ml) for 30 min and then infected with SARS-CoV-2. The infection of SARS-CoV-2 could be significantly inhibited by AXL protein at a concentration of 25 μg/ml. In contrast, SARS-CoV-2 infection of Jurkat cells could not be inhibited even at 100 μg/ml (Fig. 5b). Next, we constructed AXL-knockdown or overexpression cell lines on Jurkat cells and then tested the effect on viral infection. Our data showed that AXL knockdown could not block SARS-CoV-2 infection, but an AXL overexpression could slightly enhance the infection (1.5-fold) (Fig. 5c). Taken together, AXL should not be a main receptor for SARS-CoV-2 in Jurkat cells but it may contribute to infection.

LFA-1 is widely expressed on the surface of many leukocytes, and T-cell activation changed the structure of LFA-1 to a high-affinity mode, but not expression level.22 We then overexpressed the high-affinity alpha subunit of LFA-1 protein in ACE2 knockdown Caco2 cells (Caco2-ACE2-shRNA) and Jurkat cells. Our qPCR data showed that the LFA-1 overexpression successfully restored the dampened infection in ACE2 knockdown Caco2 cells, and also enhanced viral infection in Jurkat cells (threefold increase), as shown in cellular viral RNA levels (Fig. 5d, e). To corroborate the finding, we also performed IFA to detect viral NP expression. After an 8 h infection, viral NP-positive cells were compared. Our data showed a dampened SARS-CoV-2 infection in ACE2-knockdown cells, and a much higher NP in LFA-1 overexpression ACE2-knowckdown cells (Fig. 5f, g).

Finally, the LFA-1-knockdown Jurkat cell line was constructed and infected by SARS-CoV-2 (MOI = 0.01). At a 24 h post infection, viral load in the knockdown cell line was significantly decreased compared to the control cell line (Fig. 5h). Lifitegrast, an inhibitor that blocked LFA-1 binding to its extracellular ligand, was also used to pretreat activated Jurkat cells before infection. The qPCR results showed that at a concentration of 200 nM, Lifitegrast could also reduce the viral load in Jurkat cells (Fig. 5i). Collectively, our results suggested that LFA-1 should be an attachment factor or potential entry molecular for SARS-CoV-2 during its infection in Jurkat cells.

Discussion

Here, we showed that SARS-CoV-2 infected T lymphocytes, mainly CD4 + T cells, in an ACE2-independent manner. SARS-CoV-2 infection triggered pronounced T-cell death, which potentially contributed to lymphopenia in patients with COVID-19. T-cell infection may also pose profound influences on patients. Infected T lymphocytes not only lost the ability to control viral infection but may also carry viruses to other parts of the body through blood circulation. In addition, this ACE2-independent infection mode may compromise the therapeutic effect of neutralizing antibodies targeting at spike-ACE2 binding. These may synergistically result in more severe infection outcomes in patients with COVID-19.

It has been debated whether SARS-CoV-2 impaired the functionality of immune cell populations through direct infection. Our results provided evidence to show that SARS-CoV-2-infected T cells, as viral RNA, viral sgRNA, viral protein, and the infectious virus could be detected from T cell upon infection or from patient PBCs, although the production of infectious virus particles may stay at a low level. Several recent studies also revealed that multiple immune cells carry viral antigen or viral RNA, including neutrophils, macrophages, inflammatory monocytes, plasma B cells, T cells, and NK cells through postmortem histology analysis and single-cell/single-nuclear RNA-seq to lung or BALF.13,14,15 This suggests that SARS-CoV-2 should have a broad tropism of target cells, including major immune cells populations.

Human ACE2 and TMPRSS2 proteins were recognized as the main proteins that mediated SARS-CoV-2 cell entry.14 The newly discovered binding molecules AXL and NRP1 are still dependent on ACE2 as the main receptor.18,19 Our discovery of ACE2-independent infection of T cells is surprising, but is also supported by previous discoveries that there are SARS-CoV-2 RNA+ cells which did not co-express ACE2 and TMPRSS2.15 In our data, SARS-CoV-2 showed significant infection of activated T cells, suggesting there should be a new entry mechanism in T cells. The identification of LFA-1, as an entry molecule that contributed to a SARS-CoV-2 infection of T cells would be important for developing clinical therapeutics, although future questions remain. For example, what is the LFA-1 binding protein in SARS-CoV-2 virion if it is not the spike protein. Since LFA-1 is expressed in a number of other leukocytes, it can be expected that other immune cells (including macrophages or monocytes) could also be infected by SARS-CoV-2 potentially through binding with LFA-1. These questions should be addressed in future studies.

The infection of CD4 + T lymphocytes by SARS-CoV-2 virus may be a major contributor of virus induced pathogenesis. Armed T cells play a pivotal role against pathogen infection.10 As shown in our data, these T cells are likely to be targets of SARS-CoV-2 infection and undergo apoptosis in the HIF-1a-dependent pathway. These events may lead to T-cell dysfunction, depletion, and eventually lymphopenia in patients. In addition, the dying CD4 + T lymphocytes could trigger excessive inflammation that leads to severe immunopathogenesis in patients. Notably, the population of CD8 + T lymphocytes is also significantly decreased in COVID-19 patients. Unlike CD4 + T lymphocytes, these cells were not determined to contain SARS-CoV-2 viral antigen in flow cytometry. The mechanism underlying SARS-CoV-2 infection-induced CD8 + T lymphocytes depletion is currently unknown. Besides viral infection, several mechanisms, including the presence of endogenous or exogenous glucocorticoids, over-activated neutrophil releasing inhibitors of T cell activation (Arginase 1 and CD274) and cytokine-regulated selective differentiation of bone marrow cells, might also contribute to lymphocytes depletion.11,23 Further in-depth investigation is needed to address the potentially multi-mode mechanisms that lead to lymphopenia in the COVID-19 patients. Considering the apparent correlation between lymphopenia and disease progression in COVID-19 patients, it is important to develop strategies to prevent virus-induced lymphopenia.

Materials and methods

Samples and ethics

Human blood and tissue samples from patients with COVID-19 or from healthy donors were collected by Tongji hospital with consent from all persons. Fresh lung biopsy sections were prepared from a deceased patient. The ethics committee of the designated hospitals for emerging infectious diseases approved all human samplings.

Cell lines and virus culture

Vero E6, Caco2, 293T-sg, GP2-293, and BEAS-2B in DMEM + 10% FBS, or MT4 and Jurkat T cells in RPMI1640 + 10% FBS (Gibco, C22400500BT), or A549 cells in DMEM/F12 + 10% FBS, or primary T cells in X-vivo (Lonza, 04-418Q) medium containing IL-2 (Peprotech, 200-02) were cultured at 37 °C in a humidified atmosphere of 5% CO2. All cell lines were tested free of mycoplasma contamination and applied to species identification and authenticated by microscopic morphologic evaluation. None of cell lines was on the list of commonly misidentified cell lines (by ICLAC). SARS-CoV-2 isolate WIV04 (GISAID accession number EPI_ISL_402124) was used in this study. WIV04 was isolated from Huh7 cells from the original sample and was passaged in Caco2 cells. Viral titer (TCID50/ml) was determined in Vero E6 cells.

Proteins and antibodies for SARS-CoV-2

SARS-CoV-2 strain WIV04 NP and predicted RBD were inserted into pCAGGS vector with an N-terminal S-tag. Constructed plasmids were transiently transfected into HEK293T-17. Supernatant collected for protein purification was purified using S-tag resin, the purity and yield were tested using anti-S-tag mAb (generated in-house). Rabbits were immunized with purified NP proteins three times at a dose of 700 ng/each, 2 weeks interval. Rabbit serum was collected at 10 days after the final immunization. Antibody titer was determined in an ELISA using purified NP protein as a detection antigen.

Peripheral blood cells (PBCs) preparation and SARS-CoV-2 infection

The blood samples from patients with COVID-19 or healthy donors were processed in BSL3 lab at WIV. In all, 1× RBC lysis buffer was made from eBioscience™ 10× RBC Lysis Buffer before the experiment (Multi-species, Invitrogen). Human blood samples were centrifuged at 500 × g for 10 min before being treated with 2 ml 1× RBC lysis buffer for no more than 15 min at room temperature. Cells were spun down at 500 × g for 10 min, followed by treatment using 2 ml 1× RBC lysis buffer for another 10 min at room temperature to remove the residue red blood cells. Cells were ready for use after centrifugation. Cells were spin washed (500 × g for 10 min each time) three times with PBS containing 2% BSA before staining of cell marker antibodies.

For infection, PBCs were seeded into 24-well plates in Roswell Park Memorial Institute 1640 culture medium (RPMI1640, ThermoFisher, 22400500BT) supplemented with 10% fetal bovine serum (FBS, Life Technologies, 10099141) at a density of 1 × 106 cells/ml. PBCs were infected with SARS-CoV-2 at 0.1 MOI. One hour after incubation, cells were spin washed for three times using RPMI1640. PBCs were then seeded with RPMI1640 supplemented with 10% FBS in new 24-well plates at 37 °C supplied with 5% CO2 for 12 h or 24 h before being collected for further analysis.

For IFA on patient PBCs, overnight fixed cells were evenly smeared over a glass coverslip. The presence of viral NP was detected with rabbit pAb against the SARS-CoV-2 NP protein (generated in-house, 1:1000) and a Cy3-conjugated goat anti-rabbit IgG (1:200, Abcam, ab6939). T lymphocytes were detected using a rabbit anti-human CD3 antibody (1:100, Abcam, ab5690). Nuclei were stained with DAPI (Beyotime, C1002). Staining patterns were examined using confocal microscopy on a FV1200 microscope (Olympus).

For immunohistochemistry analysis on patient lung, the biopsy tissues from a deceased patient were fixed with 4% paraformaldehyde for 24 h, paraffin-embedded and cut into 5-μm sections. Multiplex immunofluorescence staining was obtained using PANO 7-plex IHC kit (0004100100, Panovue, Beijing, China). Slides were deparaffinized and rehydrated, followed by 15-min heat-induced antigen retrieval with EDTA pH 9.0. The slides were washed with PBS/0.02% Triton X-100 then blocked with 10% BSA at RT for 30 min, rabbit pAb against the SARS-CoV-2 NP protein (generated in-house, 1:1000) and rabbit anti-human CD3 antibody (1:100, Abcam, ab5690) were then used in incubation at 37 °C 1 h, followed by horseradish peroxidase-conjugated secondary antibody incubation and tyramide signal amplification. The slides were microwave heat-treated after each TSA operation. Nuclei were stained with 4’-6’-diamidino-2-phenylindole (DAPI, Beyotime, C1002) at the final stage of staining. To obtain multispectral images, the stained slides were scanned using the Mantra System (PerkinElmer). The scans were combined to build a single stack image. Unstained images and single-stained sections were used to extract the spectrum of autofluorescence of tissues or each fluorescein, respectively. The extracted images were further used to establish a spectral library required for multispectral immixing by InForm image analysis software (PerkinElmer). Using this spectral library, we obtained reconstructed images of sections with the autofluorescence removed.

PBCs co-cultured with Caco2 cells

PBCs from five patients were washed three times with PBS before co-cultured with Caco2 cells for 4 days and tested for SARS-CoV-2 viral RNA in the supernatant or antigen in Caco2 cells. In the meantime, PBCs were analyzed for the presence of viral antigen by flow cytometry.

Human colon organoids culture and SARS-CoV-2 infection

Human colon organoids were generated and cultured as described in the previous study.24 Briefly, colon organoids in matrigel were digested and washed twice with medium before infection. SARS-CoV-2 was added to infect colon organoids at an MOI of 0.01. 24 h later, colon organoids were then spun down and washed twice with medium. Viral RNA in colon organoids was determined by qPCR.

Activation of Jurkat and primary T cells

To activate Jurkat cells, 2.5E + 06 of cells were seeded to a well of a six-well plate containing 2.5 ml of RPMI1640 medium containing 10% FBS. In total, 40 ng/ml of PMA (Invivogen, tlrl-pma) was added to cells and incubated at 37 °C for 2 h. Cells were centrifuged at 300 × g at room temperature for 10 min before discarding the supernatant and cultured with fresh RPMI1640 medium containing 10% FBS. Primary human CD3 T lymphocytes were isolated from blood of healthy donors using CD3 Microbeads of Human (Miltenyi, 130-050-101). To activate primary T cells, frozen T cells were thawed and cultured with X-vivo (Lonza, 04-418Q) containing 1 μg/ml of IL-2 (Peprotech, 200-02). Cells were cultured with a volume of 7.5 μl of T Cell TransAct (Miltenyi Biotec) in the medium for 3 days at 37 °C. Cells were then spun down and cultured with fresh IL-2/X-vivo medium before viral infection.

T-cells infection

Jurkat T cells or primary human CD3 T lymphocytes were infected with SARS-CoV-2 at a MOI of 0.01, 0.1, or 1 depending on the purpose of the experiment. Supernatant or cells were harvested at 0, 24, 48, or 72 hpi after three times PBS washing for Jurkat T cells, or 0, 4, 8, and 12 hpi for primary T cells. Cellular or supernatant viral RNA or protein expression was determined by qPCR, RNA-seq, WB, or flow cytometry. GAPDH was used in qPCR as internal control and beta-tubulin was used in WB (1:5000, 66240-1-Ig from Proteintech) as an internal control.

Flow cytometry analysis of human peripheral blood samples

For surface staining, PBCs were incubated with fluorochrome-labeled antibodies specific for humans before fixation: AF-700-anti-CD45 (2D1), percp-anti-CD19 (HIB19), APC/CY7-anti-CD3 (UCHT1), BV510-anti-CD4 (OKT4) and percp/Cy5.5-anti-CD8a (HTT8a). Antibody-stained PBCs were fixed overnight with 4% PFA at 4 °C and taken out of BSL3 lab for downstream analysis. Cells were stained further with in-house-made SARS-CoV-1 NP pAb (1:500) at 4 °C for 30 min after permeabilization. Then cells were stained with FITC-anti-Rabbit IgG (H + L) at room temperature for 30 min. AF-700-anti-CD45 (2D1), APC/CY7-anti-CD3 (UCHT1), BV510-anti-CD4, and percp/Cy5.5-anti-CD8a antibodies were purchased from Biolegend and all were used at 1:100. FITC-anti-Rabbit IgG (H + L) was from Proteintech (SA00003-2).

RNA extraction and qPCR

Whenever commercial kits were used, the manufacturer’s instructions were followed without modification. RNA was extracted from 140 μl of samples with the QIAamp® Viral RNA Mini Kit (QIAGEN). RNA was eluted in 50 μl of elution buffer and used as the template. The qPCR detection of SARS-CoV-2 was performed using HiScript® II One-Step qPCR SYBR® Green Kit plus One-Step qPCR Probe kit targeting at either M for sgRNA (designed in house) or RBD of spike gene (commercial) following the instructions of the manufacturer (Q222-CN, Vazyme Biotech Co., Ltd). QPCR was run in a Step-One Plus real-time PCR machine (ABI) machine using default settings.

SARS-CoV-2 genome depth and coverage analysis

RNA was extracted from SARS-CoV-2 24 h-infected activated Jurkat T cells with the RNAprep Pure Cell/Bacteria Kit (TIANGEN, DP430). RNA was eluted in 50 μl of elution buffer and used as the template for RNA-seq. Clean reads were mapping to SARS-CoV-2 genome (WIV04) using software HISAT2 v2.1.0. After sorted and indexed with samtools v1.10-24, the coverage was calculated using genomeCoverageBed function from bedtools v2.29.2.

Transcriptome analysis

The SARS-CoV-2 24 h- and 48 h-infected Jurkat T cells (3 replicates each), blood samples from three healthy donors, and 3 severe COVID-19 patients were subjected for RNA-seq analysis. After mapping clean reads to GRCh38.p13 with HISAT2 v2.1.0 and format conversion with samtools v1.10-24, we used stringtie v2.1.0 to assemble and quantitate transcripts. Reads counts table of transcriptome generated by prepDE.py, a tool in stringtie, was used for gene differential expression analysis in R v4.1.0 with package DESeq2 v1.32.0. The gene with log2 fold change >2 and P value <0.05 was selected to perform enrichment analysis using online tools Metascape.

Public single-cell NGS data analysis

Public single-cell NGS data were downloaded, COVID-19 patients’ data were downloaded from GSE15805514 and healthy donors’ data were from GSE134355 (human cell landscape). According to the original information of each article, we extracted data of primary T cells from lung, thymus, and peripheral blood of healthy donors and virus-positive T cells of COVID-19 patients. Following the standard Seurat v4.0.4 workflow, we normalized the data and scaled it with UMI information. The expression of candidate receptors or co-factors was visualized with Seurat function FeaturePlot.

Western blot (WB) analysis

Infected or transduced cells were harvested at the indicated time point and lysed with RIPA Lysis Buffer (Beyotime, P0013C) for WB. Proteins in cell lysates were then separated on 10–12% SDS-polyacrylamide gel electrophoresis (PAGE) and further transferred to polyvinylidene difluoride (PVDF) membranes (Millipore, SLHVR33RB). Blots were incubated with rabbit polyclonal anti-ACE2 (Servicebio, GB11267, 1:1000 dilution), rabbit polyclonal anti-2019-nCoV NP (1:1000 dilution), mouse monoclonal anti-beta-tubulin (Proteintech, 66240-1-Ig, 1:5000 dilution), and then appropriate rabbit or mouse peroxidase-conjugated secondary antibodies (Proteintech, 1:5000 dilution, SA00001-2, or SA00001-1). Immobilon western chemiluminescent HRP substrate (Millipore, WBKLS0500) was used for protein detection.

Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay

The TUNEL Assay kit purchased from Beyotime Biotechnology (C1088) was used to detect apoptosis in SARS-CoV-2-infected cells according to the manufacturer’s instructions. Briefly, cells fixed in 4% paraformaldehyde were permeabilized with 0.25% Triton X-100 for 20 min at 4 °C. Then the TdT reaction mixture containing TdT enzyme and fluorescent labeling solution was added to the cells to label the fragmented DNA. Cells were further stained with Rp3-CoV NP pAb (1:8000) or Rabbit anti-SARS-CoV-2 NP pAb (1:500) and CY3-anti-Rabbit IgG (H + L) (Proteintech, SA00009-2) after fixation. Labeled cells were analyzed with a flow cytometer (BD LSRFortessa).

ACE2 competition inhibition and antibody blocking experiments

Human recombinant full-length ACE2-Fc protein (GenScript, Z03484), Anti-ACE2 Ab (R&D, AF933) and RD#4-anti-Spike Ab (house-made monoclonal antibodies) were used. ACE2-Fc protein was diluted to 20 μg/μl in culture medium and then incubated with SARS-CoV-2 virus solution (MOI = 0.01) at a volume of 1:1 at 37 °C for 30 min. The RD#4-anti-Spike Ab was diluted to 320 ng/μl in culture medium and then incubated with SARS-CoV-2 virus solution (MOI = 0.01) at a volume of 1:1 at 37 °C for 30 min. The virus-ACE2 or virus–antibody mixtures were then added to Jurkat cells or Caco2 cells. Cells were collected for further analysis at 24 h post infection. For anti-ACE2 antibody blocking experiments, Jurkat cells or Caco2 cells were pretreated with 3.33 ng/μl anti-ACE2 antibody (R&D Systems, goat, AF933) at 37 °C for 30 min before infection.

Generation of KO, KD, overexpression cell lines

KO, KD, and overexpression plasmids were constructed on different vectors (pLenti-V2 for knockout, pLKO.1 vector for knockdown, and pQCXIH vector for overexpression). Knockout of ACE2 was accomplished by transduction of Caco2 and Jurkat cells with lentiviruses expressing specific sgRNAs targeting ACE2 (F: CACCG GCCTCCATCGATATTAGCAA; R: AAAC TTGCTAATATCGATGGAGGCC).

Knockdown of ACE2, AXL, LFA-1 was accomplished by transduction of Caco2 or Jurkat cells with lentiviruses expressing specific siRNAs (ACE2: 5′-GCCGAAGACCTGTTCTATCAA-3′; AXL: 5′- CCTGTGGTCATCTTACCTT-3′; LFA-1: 5′-GCCATCAATTATGTCGCGACA-3′ or scramble siRNA).

Then the transduced cells were cultured with puromycin (5 μg/ml for Caco2 or 1.5 μg/ml for Jurkat) for 7 days.

For overexpression, the full length of AXL or domain I of LFA-1 alpha subunit were amplified from Hep G2 cells or Jurkat cells respectively. Lentivirus transduced cells were cultured with hygromycin (35 μg/ml for Caco2 and Jurkat cells) for 7 days. For the infection, virus was added to the cells until the end of the experiment with 0.01 MOI. Infected cells were harvested at 24 hpi after twice washing with PBS. Intracellular viral protein expression was determined by western blotting assay with antibody against virus NP protein and viral RNA in the cytoplasm was determined by qPCR.

TMPRSS2 blocking assay

Camostat mesylate (MCE, HY-13512-10 mM) was diluted to a final concentration of 20 μM or 2 μM. In total, 100 μl (for a 48-well plate) or 200 μl (for a 24-well plate) of Camostat solutions were added to cells. One hour later, activated Jurkat and Caco2 cells were infected with SARS-CoV-2 at 0.01 MOI. The cell lysate was harvested at 24 hpi and viral RNA in the cytoplasm was determined by qPCR. Viral NP was analyzed by western blot.

Candidate receptor proteins competition inhibition experiments

Recombinant Human AXL Protein (MedChemExpress, HY-P7622) was diluted to different concentrations with culture medium and then incubated with SARS-CoV-2 virus (MOI = 0.01) at a volume of 1:1 at 37 °C for 30 min. Mixtures were then added to infect activated Jurkat cells and BEAS-2B cells. Samples were harvested at 24 hpi and cellular viral RNA was determined by qPCR.

LFA-1 inhibition experiment

Lifitegrast (MedChemExpress, HY-19344) was diluted to different concentrations and pretreated activated Jurkat cells at 37 °C before infection. Thirty minutes later, cells were infected with SARS-CoV-2 (MOI = 0.01) and samples were harvested at 24 hpi. Viral RNA in the cytoplasm was determined by qPCR.

Electron microscopy

Activated Jurkat and MT4 cells were infected with the SARS-CoV-2 (MOI = 1) for 72 h. Cells were collected and fixed with 2.5% (w/v) glutaraldehyde and 1% osmium tetroxide, dehydrated through a graded series of ethanol concentrations (from 30 to 100%), and embedded with epoxy resin. Ultrathin sections (80 nm) of embedded cells were prepared, deposited onto Formvar-coated copper grids (200 mesh), stained with uranyl acetate and lead citrate, and analyzed using a 200-kV Tecnai G2 electron microscope.

Statistical analysis

Data analyses were performed using GraphPad Prism 7.0 software. Data were shown as mean ± SD. Data were analyzed with Shapiro–Wilk normality test and confirmed to the Gaussian distribution. Statistical analysis was performed using Student’s t test with two-tailed, 95% confidence. P values less than 0.05 were considered statistically significant.

Data availability

Data presented in this study are available on request from the corresponding authors. The data are not publicly available due to limitations in the material transfer agreement.

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COVID-19 and the liver

Authors: Dinesh JothimaniRadhika Venugopal,Mohammed Forhad AbedinIlankumaran Kaliamoorthy, and Mohamed Rela

J Hepatol. 2020 Nov; 73(5): 1231–1240.Published online 2020 Jun 15. doi: 10.1016/j. jhep.2020.06.006PMCID: PMC7295524PMID: 32553666

Abstract

The current coronavirus disease 2019 (COVID-19) pandemic, caused by the severe acute respiratory syndrome coronavirus-2 (SARS-CoV-2), has become a major public health crisis over the past few months. Overall case fatality rates range between 2–6%; however, the rates are higher in the elderly and those with underlying comorbidities like diabetes, hypertension and heart disease. Recent reports showed that about 2–11% of patients with COVID-19 had underlying chronic liver disease. During the previous SARS epidemic, around 60% of patients were reported to develop various degrees of liver damage. In the current pandemic, hepatic dysfunction has been seen in 14–53% of patients with COVID-19, particularly in those with severe disease. Cases of acute liver injury have been reported and are associated with higher mortality. Hepatic involvement in COVID-19 could be related to the direct cytopathic effect of the virus, an uncontrolled immune reaction, sepsis or drug-induced liver injury. The postulated mechanism of viral entry is through the host angiotensin-converting enzyme 2 (ACE2) receptors that are abundantly present in type 2 alveolar cells. Interestingly, ACE2 receptors are expressed in the gastrointestinal tract, vascular endothelium and cholangiocytes of the liver. The effects of COVID-19 on underlying chronic liver disease require detailed evaluation and, with data currently lacking, further research is warranted in this area.

Introduction

Coronaviruses are enveloped single-stranded RNA viruses, belonging to the Coronaviridae family and Orthocoronavirinae subfamily. They are some of the largest viruses (with sizes ranging from 27–34 kilobases). Coronavirus infections are commonly seen in mammals and birds. They cause zoonotic, predominantly upper respiratory tract, infections in humans. Electron microscopic images shows a ‘halo’ or ‘crown’ around the virus which explains their name. Two coronaviruses, severe acute respiratory syndrome coronavirus (SARS-CoV) and the middle eastern respiratory syndrome coronavirus (MERS-CoV), caused relatively recent epidemics, in 2003 and 2012, respectively.

The current coronavirus, responsible for the coronavirus disease 2019 (COVID-19) pandemic, has been labelled severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) by the International Taxonomy group. Genome sequencing analysis showed SARS-CoV-2 is possibly a chimeric variant of a bat coronavirus identified in 2015 by Benvenuto and Colleagues.1 The resulting disease was termed COVID-19 by the World Health Organization (WHO) on the 11th February 2020. Viral detection studies by Zhou and colleagues2 showed an 80% homology between SARS-CoV (2003 pandemic) and the current novel coronavirus.

During the previous SARS epidemic, around 60% of patients developed various degrees of liver damage. Based on phylogenetic resemblance it is possible that SARS-CoV-2 also causes liver injury.

Epidemiology

Several cases of severe unexplained pneumonia were reported in Wuhan, China in December 2019. Bronchoalveolar lavage from an index case identified the presence of SARS-CoV-2 on the 3rd January 20203 and subsequently the WHO declared an ‘epidemic’. Following the rapid increase in COVID-19 infections across the world, the WHO declared a ‘pandemic’ on the 11th March 2020, an emergency public health situation. Wuhan was the initial epicentre for COVID-19, where the first 41 cases of severe pneumonia were reported following exposure to bats and pangolins at the Huanan Seafood Wholesale market.4 Subsequent cases were reported from the same locality by Chen and colleagues.5 However, several patients in the outbreak did not have exposure to animals, likely indicating person to person transmission.

Key point

COVID-19 is a pandemic caused by SARS-CoV-2, a virus that has 80% homology with SARS-CoV.

A WHO report from the 19th May 2020, confirmed 4,731,458 COVID-19 positive cases from 213 countries worldwide, of which 1,477,516 cases were reported in the United States of America, 231,606 cases in Spain, 225,886 cases in Italy, 246,410 in the United Kingdom, 84,500 cases in China (the origin of the pandemic) and 101,139 cases in India.6 These data indicate the rapid spread of the disease around the world, with a doubling rate of 7.2 days.

ACE2 receptors

As with SARS-CoV, angiotensin-converting enzyme 2 (ACE2) appears to be the susceptible receptor for SARS-CoV-2 and is expressed in more than 80% of alveolar cells in the lungs. In vitro studies from the SARS epidemic identified ACE2 as the host receptor for viral entry.7 Immunohistochemical studies from human tissues during the SARS pandemic showed high expression of the ACE2 receptor protein in the vascular endothelium of small and large arteries and veins. In the lungs, ACE2 is highly expressed in type 2 alveolar cells. Interestingly, fibrotic lungs had much higher staining for ACE2, whereas bronchial epithelial cells showed weaker expression. A recent study showed that SARS-CoV-2 possessed 10-20-fold higher receptor binding affinity.8 Immunohistochemical studies identified higher expression of ACE2 receptors in the gastrointestinal tract. ACE2 expression is high in the basal layer of the squamous epithelium. of the nasal, oral and nasopharyngeal mucosa. Smooth muscles of the gastric and intestinal colonic mucosa also express ACE2. In addition, ACE2 is abundantly expressed in enterocytes in the duodenum, jejunum and ileum.9

Key point

ACE2 is the host cell receptor for SARS-CoV-2; it is present in type 2 alveolar cells, the gastrointestinal tract and the liver.

Hepatic distribution of ACE2 is peculiar. It is highly expressed in the endothelial layer of small blood vessels, but not in the sinusoidal endothelium. Chai and colleagues10 found that the ACE2 cell surface receptor was more highly expressed in cholangiocytes (59.7%) than hepatocytes (2.6%). The level of ACE2 expression in cholangiocytes was similar to that in type 2 alveolar cells of the lungs, indicating that the liver could be a potential target for SARS-CoV-2. Immunohistochemistry stains for ACE2 were negative on Kupffer cells, as well as T and B lymphocytes.

A recent study from Wuhan showed that Asian men had higher expression of ACE2, indicating the possibility of a higher susceptibility to COVID-19 in this population.11 , 12

Transmission

SARS-CoV-2 started as a zoonotic infection; however, the disease spreads rapidly from person to person through coughing and sneezing, particularly amongst close contacts. SARS-CoV-2 is resilient and can remain viable for 2 hours to 14 days depending on the fomite and the weather condition.13

The transmission potential of an infection in the community is based on its basic reproduction rate which is usually denoted as disease transmission ratio (R0). This represents the number of secondary cases resulting from an index case in a susceptible population. The (R0 – R naught) of COVID-19 is 2.2.14

Previous studies showed that 19.6% to 73% of patients with SARS presented with gastrointestinal symptoms.[15][16][17][18] Active replication of SARS-CoV was detected in the enterocytes of the small intestine.15 Moreover, SARS-CoV RNA was detected in patient stool samples during the SARS pandemic, which highlighted the possibility of faeco-oral transmission. A similar pattern has been observed with SARS-CoV-2; between 3% and 79% of patients with COVID-19 develop gastrointestinal symptoms, predominantly nausea, vomiting and diarrhoea. Zhang et al. found that 53.3% and 26.7% of oral and anal swabs remained positive for SARS-CoV-2 RNA, respectively, for several days after treatment. The same study group performed paired samples on a different cohort of patients with COVID-19 and found that on day 0, 80% of patients were positive on oral swabs whereas on day 5, 75% of patients were positive on anal swabs, indicating the dynamic changes in viral tests during the course of the illness.19 Xiao and colleagues20 showed that patients with SARS-CoV-2-related respiratory illness can continue to shed the virus in stool even after a negative respiratory sample. In a series of 73 patients with COVID-19, about 53.42% had detectable RNA in their stool, of whom about 23.29% continued to have positive RT-PCR for SARS-CoV-2 RNA in faecal samples even after a negative respiratory sample.20 Yeo and colleagues21 showed that faecal shedding can continue to occur for a longer period after clinical recovery and these patients can potentially infect others. These findings illustrate the multiple routes of viral entry into a single host, viral persistence in various organ systems and possible faecal-oral transmission of SARS-CoV-2 even during the convalescence period.

Key point

In addition to droplets, SARS-CoV-2 also transmits through the faeco-oral route.

With limited therapeutic options, prevention by social distancing appears to be the cornerstone of COVID-19 management. Virus transmission can be reduced by various methods described in the WHO protocol.6 These include, maintaining safe social distance, regular hand washing for 20 seconds, using 60% alcohol hand rub, and avoiding crowded places and public events. Countries have taken different measures to reduce viral transmission and most countries have gone into ‘Lockdown’ in order to stop viral transmission. Being a large virus particle, a surgical face mask should provide adequate protection against viral inhalation. N-95 masks should be reserved for treating teams. Personal protective equipment should be worn according to institutional policy. All patients with a history of travel to affected regions should be screened for SARS-CoV-2 even if they are asymptomatic. People with high temperature, dry cough, profound tiredness, diarrhoea or other unusual symptoms with recent travel history should be tested for COVID-19. Nations will need to continually monitor their prevention, testing and treatment strategies based on guidelines issued by the WHO.

Clinical features

Initial reports from China showed that the incubation period of SARS-CoV-2 was between 3 to 7 days and occasionally 2 weeks. The longest incubation period identified was 12.5 days.14

Large studies from a Chinese population reported fever (≥38°C), dry cough, fatigue, myalgia, leukopenia and raised liver enzymes as the most common clinical features of COVID-19 on presentation, as shown in Table 1 and ​and2 .2 . Nausea, vomiting and diarrhoea were seen in 2–10% of patients with COVID-19.

Table 1

Spectrum of clinical manifestations and their frequency from recent studies on COVID-19 in China.

Clinical featuresWang et al.22
n = 138
Zhou et al.51
n = 191
Guan et al.23
n = 1,099
Fever98.6%94%88.7%
Cough59.4%79%67.8%
Sputumn.a.23%33.7%
Myalgian.a.15%14.9%
Fatigue69.6%23%38.1%
Diarrhoean.a.5%3.8%
Nausea/vomitingn.a.4%5.0%
Sore throatn.a.n.a.13.9%
Lymphopenia (<0.8 × 109/L)70.3%40%n.a.
Prolonged PT (>13.5 seconds)58%n.a.n.a.
Raised LDH (>261 U/L)39.9%n.a.n.a.

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COVID-19, coronavirus disease 2019; LDH, lactate dehydrogenase; PT, prothrombin time; n.a., data not available.

Table 2

Classification of COVID-19 into 3 groups based on severity of clinical manifestations by Chinese Center for Disease Control.23

Mild disease (reported in 81% cases)Fever, dry cough, mild dyspnoea (respiratory rate <30/min).
Severe disease (reported in 14% cases)Dyspnoea, respiratory rate >30 and/or lung infiltrates >50% within 24 to 48 hours.
Critical disease (reported in 5% cases)Respiratory failure, septic shock and/or multiple organ dysfunction or failure.

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COVID-19, coronavirus disease 2019.

In the latest case series from Wuhan by Wang and colleagues,22 138 hospitalised patients (including 40 healthcare workers and 17 already hospitalised for other conditions) with COVID-19; median age was 56 years (IQR 22–92 years) and 54.3% were males. Clinical features were fever (98.6%), fatigue (69.6%), dry cough (59.4%), lymphopenia <0.8 × 109/L (70.3%), prolonged prothrombin time (58%), and raised lactate dehydrogenase (LDH) 261 U/L (39.9%). Thirty-six patients (26.1%) received intensive care unit (ICU) care for acute respiratory distress syndrome (ARDS) (61.1%), cardiac arrhythmias (44.4%) and shock (30.6%). Onset and the progression of symptoms were dramatic, with a median time from symptoms to ARDS of only 8 days. Patients requiring intensive care were older (66 vs. 51, years) and more often had comorbidities (72% vs. 32%). Patients admitted to the ICU had higher LDH (435 U/L vs. 212 U/L, p <0.001), aspartate aminotransferase (AST) (52 U/L vs. 29 U/L, p <0.001) and hypersensitive cardiac troponin (11 ng/ml vs. 5.1 ng/ml, p = 0.004). All 138 patients showed bilateral pneumonia in the thoracic scan. Analysis between the survivors and non-survivors showed higher white blood cell count with severe progressive lymphopenia in the non-survivors. With disease progression, these patients required organ support with progressive deterioration in renal function before death.

In the largest database analysis of 1,099 patients with confirmed COVID-19 from China, by Guan and colleagues,23 the median age of presentation was 47 years (IQR 35–58 years) and 58% were male. The most common presenting symptoms were fever (88.7%), cough (67.8%), nausea or vomiting (5%), and diarrhoea (3.8%). CT chest radiography revealed ground glass opacity (56.4%) and bilateral patchy shadows (51.8%). Of 1,099 patients, 5% were admitted to the ICU, 2.3% underwent invasive ventilation and 1.4% died. COVID-19 disease was classified according to the clinical severity into 3 groups by the Chinese CDC by Guan and colleagues23 as shown in Table 2.

Key point

2–11% of patients with COVID-19 have been reported to have underlying chronic liver disease.Go to:

COVID-19 and hepatic dysfunction

It is intriguing to know the pattern of liver injury in COVID-19. Hepatic involvement in COVID-19 could be related to the direct cytopathic effect of the virus, an uncontrolled immune reaction, sepsis or drug-induced liver injury. Given the higher expression of ACE2 receptors in cholangiocytes, the liver is a potential target for SARS-CoV-2. Moreover, COVID-19 may cause worsening of underlying chronic liver disease, leading to hepatic decompensation and acute-on-chronic liver failure, with higher mortality.

A summary of recently published studies is provided in Table 3 . Overall, 2–11% of patients with COVID-19 were reported to have underlying chronic liver disease and 14-53% with COVID-19 developed hepatic dysfunction,24 particularly those with severe COVID-19. Hepatic dysfunction was significantly higher in critically ill patients and was associated with poor outcome.

Table 3

Studies of COVID-19 and hepatic manifestations.

AuthorCountryComments
Chen et al.26ChinaHigher ALT and AST in deceased patients.
High mortality in patients with acute liver injury (76.9%).
Li et al.74China7% of patients with COVID-19 had underlying chronic liver disease.
Wang et al.22China3.9% of patients with COVID-19 had underlying chronic liver disease.
Mortality 4.3%.
Guan et al.23China2.1% of patients with COVID-19 had chronic hepatitis B infection.
Mortality 1.4%.
Huang et al.4ChinaMortality 15%.
1 (4%) patient with COVID-19 had underlying chronic liver disease.
Fan et al.28ChinaPatients with abnormal LFT had longer hospital stay (16.4 vs. 12.6 days).
Cai et al.75ChinaHigher AST, ALT and GGT in patients with severe disease.
Patients with NAFLD had severe disease.
Cao W.76ChinaHigher ALT and AST in patients with severe COVID-19.
Shi et al.77China7 (3%) patients with COVID-19 had underlying chronic liver disease.
Wu et al.78China3% (7) had underlying CLD.
Bilirubin was significantly higher in patients with ARDS-related death.
Graselli et al.79Italy15-30% mortality in patients between 50–70 years of age.
Arentz et al.80USA3 (14.7%) patients developed acute liver injury.
Zhang et al.24ChinaMortality 1.7%.

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ALD, alcohol-related liver disease; ALT, alanine aminotransferase; ARDS, acute respiratory distress syndrome; AST, aspartate aminotransferase; CLD, chronic liver disease; COVID-19, coronavirus disease 2019; GGT, gamma glutamyltransferase; ICU, intensive care unit; LFT, liver function test; NAFLD, non-alcoholic fatty liver disease.

In the recent series from Wuhan, by Wang and colleagues,22 4 patients (2.9%) with COVID-19 had underlying chronic liver disease. Another study from China23 showed that 23 (2.1%) patients were positive for HBsAg, of whom only one had severe COVID-19. Interestingly, a study from outside Wuhan by Xu and colleagues25 identified 26 patients with COVID-19 in whom 11% had underlying chronic liver disease. In another study, comparing 113 non-survivors and 161 survivors showed that 4% had underlying chronic hepatitis B.26 Cases of acute liver injury were reported in 13 (5%) out of 274 patients of whom 10 (76.9%) died.26

With the current evidence, it is clear that elevated liver enzymes are observed predominantly in severe and critical cases of COVID-19. Raised AST was noted in 8/13 (62%) patients in ICU compared to 7/28 (25%) in the non-ICU setting.24 The peak alanine aminotransferase (ALT) and AST levels noted were 7,590 U/L and 1,445 U/L, respectively, in severe COVID-19.27 Interestingly, a higher proportion of enzyme elevation was noted in patients receiving lopinavir/ritonavir therapy (56.1% vs. 25%).28 It was unclear whether the elevated liver enzymes were due to the disease per se or drug-induced liver injury in this population. There is a possible effect of liver damage due to inflammatory cytokine storm in severe COVID-19.29

Key point

14–53% of patients with COVID-19 have been reported to develop some form of hepatic dysfunction.

Interestingly, despite the presence of ACE2 in cholangiocytes, more patients developed raised transaminases. An, unpublished data from Wuhan, China, by Xu et al. showed increased gamma glutamyltransferase (GGT) levels in severe cases of COVID-19.30 Whether COVID-19 aggravates cholestasis in patients with primary biliary cholangitis and primary sclerosing cholangitis requires further analysis.31 It is possible that hepatic dysfunction may result from cytokine storm rather than the direct cytopathic effects of the virus. More data is required to ascertain the pattern and the degree of liver injury in patients with COVID-19.

COVID-19 liver histology

Xu et al. reported the first post-mortem findings of a patient who succumbed to severe COVID-19. In his study, the liver histology revealed moderate microvesicular steatosis and mild inflammatory infiltrates in the hepatic lobule and portal tract. However, at this stage, it is unclear whether these changes are related to the viral infection or to the drugs. In addition, peripheral blood examination showed significantly reduced but hyper-reactive CD4 and CD8 cells in a proinflammatory state, with increased CCR6+ Th17 CD4 T cells and cytotoxicity granulations in CD8 cells, which may also contribute to hepatocellular dysfunction.32

In another report by Tian S et al., post-mortem liver biopsies in 4 patients with COVID-19 showed mild sinusoidal dilatation and focal macrovesicular steatosis. There was mild lobular lymphocytic infiltration, which was not significant in portal areas. SARS-CoV-2 RNA was isolated from liver tissue through RT-PCR in one of the patients. Though the bile duct epithelium expresses higher levels of ACE2 receptors, there was not much evidence to point towards bile duct damage.33

During the SARS-CoV outbreak in 2002, 23% to 60% of patients had hepatic dysfunction and few patients underwent liver biopsy. This revealed mild to moderate lobular lymphocytic inflammation, ballooning of hepatocytes and apoptosis. The most prominent feature was high mitotic figures indicative of a rapidly proliferative state (positive Ki-67). The Ki proliferative index of hepatocytes in chronic hepatitis C infection is around 0.45 to 1% suggestive of high replicative phase of hepatocytes in chronic hepatitis C infection. Immunohistochemistry studies showed that the Ki proliferative index of hepatocytes during SARS-CoV infection was much higher than during chronic hepatitis C infection and liver regeneration. The mitotic index was probably due to cell cycle arrest following SARS-CoV infection. It is possible that COVID-19 has a similar pathogenesis.34

Liver abnormality in SARS

SARS was a major pandemic in 2003. Hepatic dysfunction was described in patients with SARS. Up to 10% of patients had underlying chronic liver disease, particularly, chronic hepatitis B, probably owing to the geographic location of the SARS outbreak. Over 50% of patients developed abnormal liver function tests (mostly mild) and the majority recovered. However, in some studies, elevated liver function tests were associated with severe disease and, in particular, high ALT predicted ICU admission and death. This raised the possibility that SARS caused liver dysfunction rather than simply being associated with it.[35][36][37][38][39][40][41]

Liver abnormality in MERS

The first case of MERS-CoV infection was reported in 2012 in Saudi Arabia.42 Unlike SARS-CoV and SARS-CoV-2, MERS-CoV utilises dipeptidyl peptidase-4 (DPP-4), which is abundant in the liver, as the cell entry receptor.43 Low albumin was found to be an independent predictor of severe MERS-CoV infection.44 The liver biopsy in patients with MERS showed lobular lymphocytic infiltration and mild hydropic degeneration of hepatocytes.45 , 46 In patients with MERS, non-survivors had a higher incidence of liver injury than survivors (91.3% vs. 77.9%, respectively).47 , 48 Mortality was higher in patients with comorbidities.49 , 50

Clinical outcome of COVID-19

According to Wang and colleagues,22 disease progression manifested as increasing respiratory distress leading to pneumonia. In these patients, CT showed bilateral ground glass appearance and patchy pneumonia in almost 100% of patients. Most patients recovered with no sequalae. Overall, in patients with severe COVID-19, 19.6% developed ARDS, 16.7% had myocarditis which manifested as arrhythmias and 8.7% developed septic shock. However, this number was higher in patients admitted to the ICU; ARDS (61%), arrhythmias (44.4%), and shock (30.6%). These patients required mechanical ventilation and extracorporeal membrane oxygenation (ECMO).

Case fatality rates of 3.6–15% have been reported in 4,292 Chinese patients. Mortality was higher in men (3.25:1), those aged >75 years and those with comorbidities (diabetes mellitus, hypertension and cardiovascular disease). These comorbidities were noted in 48% of patients in a study by Zhou and colleagues51 reporting on 191 patients with COVID-19: 54 died (28.2% mortality) of whom 36 (66.6%) had underlying chronic disease. Fig. 1 illustrates the distribution of comorbidities in deceased patients. In the largest case series by Wu and colleagues,52 the overall mortality was 2.3%; however, the mortality rate was 49% in patients with critical disease. In a recent report from Italy by Remuzzi and colleagues,53 mortality related to COVID-19 was 6% (827 patients), with a male:female ratio of 4:1 and a mean age of 81 years among those who died. More than 60% of these patients had comorbidities. The median time from presentation to death was 14 days.4 , 22 Age-adjusted mortality in these 2 large series is shown in Fig. 2 .

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Fig. 1

Distribution of comorbidities in deceased patients with COVID-19.

COVID-19, coronavirus disease 2019.

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Fig. 2

Comparison of the case fatality rates of COVID-19 based on respective age groups in 2 large cohorts from China52 and Italy.53

COVID-19, coronavirus disease 2019. n.a., no data were available for age groups 50-59, 60-69, >90 years in the Chinese cohort.

According to a meta-analysis of 8 studies, including 46,248 patients, which analysed the prevalence of comorbidities in COVID-19, the most common comorbidities were hypertension (14–22%), followed by diabetes mellitus (6–11%), cardiovascular diseases (4–7%) and respiratory disease (1–3%).54 The mortality rate was higher in patients with hypertension (48%), followed by 21% in diabetics, 14% in patients with cardiovascular illness, 10% in those with chronic lung disease, and 4% each for malignancy, chronic kidney disease and cerebrovascular diseases.26 However, the mortality rate in patients with underlying chronic liver disease was 0–2%.55 In this analysis, hypertension (48% vs. 24%), diabetes (21% vs. 14%), and cardiovascular disease (14% vs. 4%) were more common in non-survivors. Fatty liver disease is likely seen as part of the metabolic syndrome in this group of patients, which can complicate the issue.

Another study from Wuhan reported on the characteristic features of deceased patients (n = 113). AST, ALT, alkaline phosphatase, GGT and bilirubin levels were significantly higher in non-survivors than survivors. Elevated AST (>40 U/L) was observed in 59 (52%) deceased and 25 (16%) recovered patients and likewise elevated ALT (>41 U/L) was found in 30 (27%) deceased and 30 (19%) recovered patients. Similarly, hypoalbuminemia (<32 g/L) was found in 74 (65%) deceased patients compared to 22 (14%) recovered patients. Serum bilirubin was 12.6 μmol and 8.4 μmol in the deceased and recovered patients, respectively. In a recent report by Chen et al., 13 (5%) patients with COVID-19 developed acute liver injury during the course of the illness of whom 10 (76.9%) died.26 Although the numbers are small, this conveys an important message on patients with COVID-19 and hepatic dysfunction.

Key point

Hepatic dysfunction was significantly more frequent in critically ill patients and was associated with poor outcome.

Diagnosis

Diagnosis of COVID-19 was based on Real time reverse transcription polymerase chain reaction (RT-PCR). In the case series described by Wang and colleagues22 centrifuged throat swab samples were used for testing. The total viral RNA was extracted within 2 hours using an RNA isolation kit. RT-PCR of the suspension was performed and amplification of Open reading frame (ORIF) and nucleocapsid protein were carried out using respective forward, reverse primers and the probe. Diagnosis were also obtained using nasal swabs, oral and rectal swabs. Interestingly, Xiao and colleagues20 showed patients with SARS CoV-2 related respiratory illness can continue to shed virus in stool even after a negative respiratory sample.

Management

Although the evidence is less clear, the current treatment recommendations include antiviral drugs, antibiotics, intravenous fluids and corticosteroids. Oseltamivir was utilised in 89.9% of patients in the Wuhan series. Remdisivir was though initially promising, a recent randomized control study did not show clinical benefit in COVID-19 except non significant faster clinical recovery. Moreover, liver injury was observed in 10-13 % of remdisivir treated group.56 Being an RNA virus, one would expect broad spectrum ribavirin to work; unfortunately, during the SARS outbreak, ribavirin was associated with significant toxicity including severe haemolysis. Interestingly, Omrani and colleagues57 found interferon alpha 2 A in combination with ribavirin to improve survival at day 14 (70% vs. 17%, p = 0.004) but not day 28 (30% vs. 17%, p = 0.054) during the MERS-CoV outbreak.

Lopinavir/ritonavir, approved for HIV infection showed in vitro activity against SARS-CoV and was beneficial in MERS-CoV.58 These drugs are being tried in COVID-19. Lopinavir, a protease inhibitor, has been shown to be effective in controlling SARS-CoV. Ritonavir was added to increase the trough level of lopinavir through CYP450 enzyme inhibition in liver. A recently published open labelled, randomised controlled trial on 199 patients with severe COVID-19 showed no benefit of lopinavir and ritonavir (99 patients). It was debated

Key point

Current treatment recommendations for COVID-19 include corticosteroids, antiviral drugs, antibiotics and intravenous fluids.

whether the trial should have been conducted in less sick patients and treatment should have been initiated in an earlier phase of COVID-19. In this study, 20.5% and 41% of patients had elevated AST and ALT prior to randomisation, respectively; however, the presence of cirrhosis, ALT or AST >5 times the upper limit normal were exclusion criteria in this trial. Increased bilirubin and elevated AST were noted in 3.2% and 2.1% of patients in the treatment group, respectively.59 Importantly, using ritonavir to inhibit CYP450 will increase the trough levels of calcineurin inhibitors, the most commonly used immunosuppression in solid organ transplant recipients, leading to potential drug toxicity.

Antibiotics such as fluoroquinolones and third-generation cephalosporins were used to reduce secondary infection. Corticosteroids (methylprednisolone) have been used in patients with COVID-19 to curtail inflammation22 and, recently, dexamethasone has been found to reduce mortality. Their use can lead to the reactivation of chronic hepatitis B. Thus, HBsAg-positive patients should be given antiviral therapy and we recommend checking hepatitis B core antibody status and, if positive, treating patients with antivirals for the duration of steroid therapy.

Recently, Chen et al. constructed a 3-dimensional crystal structure model of SARS-CoV-2 proteases. Virtual screening of the active viral site demonstrated that hepatitis C NS5A inhibitors could be effective in controlling SARS-CoV-2. Ledipasvir and velpatasvir readily inhibited SARS-CoV proteases in their model. However, more evidence is required.

COVID-19 and HCC

Patients with underlying cancer are often immunosuppressed, as result of the natural history of the disease and chemotherapy. In a nationwide study of 1,590 cancer patients with COVID-19 across 575 hospitals in China, it was observed that patients with cancer were at higher risk of contracting SARS-CoV-2 infection and developing severe illness. They also had worse outcomes than those without cancer.61 Most patients with hepatocellular carcinoma (HCC) have underlying chronic liver disease and therefore, they fall under this high-risk category and are likely to have worse outcomes. AASLD currently recommends delaying HCC surveillance by 2 months; however, HCC-related treatments should be carried out without much delay.31 EASL recommends avoiding HCC surveillance in COVID-19-positive patients, postponing locoregional therapy and temporarily withholding immune checkpoint inhibitor therapy.62

COVID-19 and deceased donor transplantation

There has been a significant decline in cadaveric organ donation during the COVID-19 pandemic.63 This can affect patients awaiting liver or other solid organ transplantation, leading to increased waiting list mortality. There has been a recent debate on harvesting organs from SARS-CoV-2-positive donors, like the discussion around HCV-positive donors.64 However, the risk of disease transmission to the transplant team remains a major concern.65 This may be an interesting option in the future, when effective vaccination comes available.

Post-liver transplant COVID-19

COVID-19 leaves no stone unturned, including liver transplant recipients. A recent case report from Wuhan described a 37-year-old man with hepatitis B and HCC, who developed fever on the third day post transarterial chemoembolisation. He was initially treated with antibiotics and subsequently liver transplantation on day 7. His fever continued on day 9, and a CT scan of his chest showed hypostatic changes in both lung fields. A repeat CT on the third week showed bilateral ground glass appearance. His nasopharyngeal swab confirmed COVID-19. His tacrolimus dose was reduced and maintained under 10 ng/ml. His liver enzymes increased by the fourth week but settled gradually. His PCR remained positive for nearly 2 months and subsequently cleared.66

Another case of post-transplant COVID-19 was described recently. The patient underwent cadaveric liver transplantation in July 2017. He presented recently with high fever and developed severe COVID-19. His tacrolimus was discontinued for a month, but he received corticosteroid therapy. His allograft function remained normal.67

Some immunosuppressive drugs possess antiviral activity by virtue of their mechanism of action. Studies from SARS identified an interaction between SARS-CoV non-structural proteins and cyclophilins, resulting in modulation of T cell immune responses. In vitro studies showed that cyclosporine inhibited SARS-CoV at higher doses. However, its clinical utility was limited by its profound immunosuppressive effects.68 Similarly, mycophenolic acid exhibited potent antiviral properties against MERS-CoV in vitro.69 Interestingly, mTOR inhibitors (everolimus) showed effectiveness against SARS-CoV and MERS-CoV infections by blocking early viral entry and post-entry consequences.70 , 71 Although in vitro studies, the antiviral properties of these drugs may offer some protection against COVID-19 in transplant recipients, particularly to ameliorate disease severity.

Literature from SARS-CoV and MERS-CoV showed that post-liver transplant patients on immunosuppression were not at higher risk of mortality. Similar data on SARS-CoV-2 are very limited.72

Rapid clinical deterioration in COVID-19 is often due to a cytokine storm associated with elevated interleukin (IL)-6, IL-8 and tumour necrosis factor-alpha levels. The combined effects of SARS-CoV-2 infection and immunosuppression are not well established. However, stopping immunosuppressive medications in transplant patients may lead to rejection. In patients with COVID-19 on high dose steroids, the dose needs to be tapered and maintained at 10 mg/day. When there is lymphopenia, fever and worsening lung condition, azathioprine, mycophenolate and calcineurin inhibitor doses need to be reduced but not stopped. Caution needs to be exercised when considering initiation of steroids or other immunosuppressive therapy in patients with severe alcoholic hepatitis, autoimmune hepatitis etc.31 Patients on immunosuppression may be more infectious as they have higher viral titres.73

The American Society of Transplantation has provided a few recommendations specifically for those awaiting liver transplantation and transplant recipients during the current pandemic. The recommendations include patient education, hand hygiene and social distancing, provision for patients to contact the transplant centre via telephone if they develop fever, cough or flu-like symptoms. Each hospital should provide layout protocols for managing these high-risk patients. Allograft function and drug interactions should be carefully monitored in transplant recipients with COVID-19, because ritonavir can potentially inhibit the CYP34A enzyme, leading to increasing trough levels of mTOR and calcineurin inhibitors, and possibly drug toxicity. In addition, they have recommended postponing elective surgeries including living donor transplantation and non-urgent deceased donor transplantations in areas with a high prevalence of COVID-19. In addition, potential deceased donors should be adequately tested for SARS-CoV-2 with nucleic acid assays.73

Conclusion

COVID-19 is currently a pandemic, with an overall mortality rate of 2–6% in infected patients, which increases with age and comorbidities. COVID-19 causes pneumonia, but hepatic dysfunction can occur in severe cases and is associated with fatal outcome. Cases of severe acute liver injury have been reported with higher mortality. Larger studies with long-term follow-up are required to characterise the extent and cause of liver damage in COVID-19. The effects of COVID-19 on underlying chronic liver disease require detailed evaluation, with further research warranted in this area.

Abbreviations

ACE2, angiotensin-converting enzyme 2; ALT, alanine aminotransferase; ARDS, acute respiratory distress syndrome; AST, aspartate aminotransferase; COVID-19, coronavirus disease 2019; GGT, gamma glutamyltransferase; HCC, hepatocellular carcinoma; ICU, intensive care unit; MERS, Middle East respiratory syndrome; MERS-CoV, MERS coronavirus; NAFLD, non-alcoholic fatty liver disease; RT-PCR, reverse transcription PCR; SARS, severe acute respiratory syndrome; SARS-CoV, SARS coronavirus; SARS-CoV-2, SARS coronavirus 2; SIRS, systemic inflammatory response syndrome; WHO, World Health Organization.

Financial support

The authors received no financial support to produce this manuscript.

Authors’ contributions

Dinesh Jothimani: Conceptualization; Project Administration; Supervision; Writing -original draft; Writing – review & editing. Radhika Venugopal: Data curation; Resources; Software; Writing – review & editing. Mohammed Forhad Abedin: Resources; Writing – review & editing; Ilankumaran Kaliamoorthy: Supervision; Validation; Writing – review & editing; Mohamed Rela: Conceptualization; Supervision; Writing – review & editing.

Conflict of interest

The authors declare no conflicts of interest that pertain to this work.

Please refer to the accompanying ICMJE disclosure forms for further details.

Footnotes

Author names in bold designate shared co-first authorship

Supplementary data to this article can be found online at https://doi.org/10.1016/j.jhep.2020.06.006.

Supplementary data

disclosures.pdf:Click here to view.(185K, pdf)Go to:

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Does COVID-19 Cause Hypertension?

Authors: Mahmut Akpek, MD  December 10, 2021 Research Article https://doi.org/10.1177/00033197211053903

Abstract

The coronavirus disease 2019 (COVID-19) outbreak remains a major public health challenge worldwide. The present study investigated the effect of COVID-19 on blood pressure (BP) during short term follow-up. A total of 211 consecutive COVID-19 patients who were admitted to Parkhayat Kutahya hospital were retrospectively screened. Information was obtained from the electronic medical records and National health data registry. The study outcome was new onset of hypertension according to the Eight Joint National Committee and European Society of Cardiology Guidelines. Finally, 153 confirmed COVID-19 patients (mean age 46.5 ± 12.7 years) were enrolled. Both systolic (120.9 ± 7.2 vs 126.5 ± 15.0 mmHg, P <.001) and diastolic BP (78.5 ± 4.4 vs 81.8 ± 7.4 mmHg, P <.001) were significantly higher in the post COVID-19 period than on admission. New onset hypertension was observed in 18 patients at the end of 31.6 ± 5.0 days on average (P <.001). These findings suggest that COVID-19 increases systolic and diastolic BP and may cause new onset hypertension.

Introduction

The coronavirus disease 2019 (COVID-19) outbreak remains a major public health challenge worldwide. COVID-19 disease is caused by a novel coronavirus: severe acute respiratory syndrome coronavirus-2 (SARS-CoV-2).1 The number of new cases continues to increase and >4 million people died due to the COVID-19 pandemic.2 With the growing understanding of the pathophysiology of this disease and its effect on multiple systems, various comorbidities were investigated.3 The post COVID-19 infection period should also be investigated in order to identify the short-, mid-, and long-term adverse outcomes and new onset comorbidities that may develop.

Hypertension is associated with an increased risk of severe COVID-19 and higher mortality rate in these patients.4 However, the effect of COVID-19 on blood pressure (BP) has not yet been elucidated. Therefore, the present study aimed to investigate the effect of COVID-19 on BP and change in the prevalence of hypertension in COVID-19 patients.

Methods

In the present retrospective cohort study, a total of 211 consecutive COVID-19 patients who admitted to Parkhayat Kutahya hospital from December 15, 2020 to April 01, 2021 were screened. The following patients were excluded: those under the age of 18, who received steroid therapy, with systemic inflammatory disease history, previous kidney or liver failure history, with hypertension, who left the follow-up and with missing data. Finally, 153 eligible patients (75%) were analyzed (Figure 1).

Figure 1. Study participants.

The diagnosis of COVID-19 was confirmed by the detection of the presence of SARS-CoV-2 ribonucleic acid on an oropharyngeal and nasopharyngeal swab using reverse transcriptase polymerase chain reaction in the Public Health Microbiology Laboratory of the Ministry of Health according to World Health Organization guidance.5 Oropharyngeal and nasopharyngeal swabs were collected at the time of admission to the outpatient unit.

The study outcome was new onset hypertension. New onset hypertension was defined as values ≥140 mmHg systolic BP and/or ≥90 mmHg diastolic BP in office measurements and ≥135 mmHg systolic BP and/or ≥85 mmHg diastolic BP in home BP monitoring according to Eighth Joint National Committee (JNC 8) and European Society of Cardiology guidelines.6,7 In the COVID-19 unit, BP was measured 3 times on the right upper-arm in the seated position by a trained nurse using a sphygmomanometer after 15 min resting. The average of the 3 measurements was used. Proper cuff size was determined based on arm circumference. The measurement was performed under controlled condition in a quiet room.

The patient characteristics, laboratory results, treatment protocol and outcome data of patients were obtained from the electronic medical records of Parkhayat Kutahya hospital and National health data registry (e-Nabız®). For all patients, blood samples for routine laboratory analysis were drawn upon admission and follow-up in the COVID-19 outpatient unit. Laboratory analyses were performed in the laboratories of Parkhayat Kutahya hospital.

Complete blood count was measured with ELite 580 advanced hematology analyzer (Erba, Czech Republic). C-reactive protein, lactate dehydrogenase (LDH), alanine aminotransferase (ALT), aspartate aminotransferase (AST), and creatinine were measured by Beckman Coulter AU 640 (Japan) analyzer. Ferritin and high sensitive troponin-I (HsTrop-I) were measured by Beckman Coulter DXI 800 (Japan) analyzer. D-dimer was determined with the Getein 1600 immunofluorescence quantitative analyzes (China). Bio-Speedy® SARS-CoV-2 (2019-nCoV) qPCR Detection Kits (Bioeksen, Istanbul, Turkey) were used to detect COVID-19. Repeatability of the kit is 100% and the reproducibility combined with the robotic extraction is 100% at concentrations over the LOD (Limit of detection). LOD for all the sample types is 20 genomes/mL. Sensitivity and specificity of the Bio-Speedy® kit were 99.4–99.0%, respectively.8

The management of the treatment protocol for COVID-19 was left to the discretion of the pandemic team consisting of infection disease, radiology, chest disease, anesthesiology, cardiology, and internal medicine specialized medical doctors and pharmacists as recommended by updated guideline by the Turkish Ministry of Health. The present study was approved by the institutional review board (Protocol no: E-41997688-050.99-8877) and the Republic of Turkey Ministry of Health.

Statistical Analysis

Continuous variables were tested for normal distribution by the Kolmogorov–Smirnov test. The variables are expressed as means ± standard deviation or median (interquartile range). Dependent continuous variables were compared with paired sample t-tests or Wilcoxon signed rank tests, as appropriate. Dependent categorical variables were compared with McNemar’s test. We performed a power analyses according to changes in systolic and diastolic BP by the follow-up period and found a power of >.98 (P = 1-β error probability) for both. The power analyses for new onset hypertension as a dependent categorical variable was .88 (P = 1-β error probability). A two-tailed P <.05 was considered significant. All statistical analyses were performed using the SPSS statistical package for Windows version 15.0 (SPSS Inc., Chicago, IL, USA).

Results

A total of 153 confirmed COVID-19 patients (mean age 46.5 ± 12.7 years) were enrolled; 101 patients (66%) were female. Table 1 shows the baseline characteristics of the study population. Body mass index was 25.8 ± 4.4. The common symptoms were fatigue, cough, and fever (74%, 65% and 49%, respectively). Sore throat was seen in 42% of patients while dyspnea was seen in 39% and myalgia was seen in 39% of the study population. Hyposmia, dysosmia, anosmia, headache and diarrhea were rare symptoms on admission in patients with COVID-19. Favipiravir and chloroquine/hydroxychloroquine were the most given drugs (78% and 77%, respectively). Anti-coagulants were administered for 38% of patients. Only 8 patients (5%) were hospitalized. Mean hospitalization time was 6.1 ± 1.0 days. Mean follow-up time was 31.6 ± 5.0 days.

Table 1. Baseline characteristics.

Table 1. Baseline characteristics.View larger version

Clinical characteristics and laboratory findings are shown in Table 2. There was no significant difference in hemoglobin, white blood cell, and lymphocyte count on admission and after COVID-19 (P = .728, P = .224, P = .272, respectively). The serum CRP level (5.0 (2.0–10.4) vs 3.0 (2.0–5.0) mg/L, P <.001) and D-dimer level (149.0 (100.0–300.0) vs 119.9 (100.0–187.7) ng/mL, P <.001) were significantly higher on admission than in post COVID-19 period. High sensitive troponin-I significantly decreased in the post COVID-19 period (9.6 ± 6.4 vs 3.8 ± 3.4 pg/mL, P <.001). Ferritin, lactate dehydrogenase, creatinine, and transaminases levels were not significantly different between on admission and the post COVID-19 period. New onset hypertension was observed in 18 patients (12%) during post COVID-19 period (P <.001), while diabetes mellitus, coronary artery disease, and chronic obstructive pulmonary disease were not significantly different between admission and post COVID-19 period (P = .375, P = .500 and P = .125, respectively). Both systolic (120.9 ± 7.2 vs 126.5 ± 15.0 mmHg, P <.001) and diastolic BP (78.5 ± 4.4 vs 81.8 ± 7.4 mmHg, P <.001) were significantly higher in the post COVID-19 period when compared with on admission (Figure 2).

Table 2. Clinical characteristics and laboratory findings.

Table 2. Clinical characteristics and laboratory findings.View larger version

Figure 2. Systolic and diastolic blood pressure on admission and post COVID-19 period.

Discussion

Since the outbreak of COVID-19 was recognized, there have been 188,650,179 confirmed cases and >4,000,000 deaths, reported to the WHO.2 Since the pandemic started, published research focused on evaluating the optimal treatment to reduce COVID-19 mortality. Recent studies also focused on the determination of independent predictors of mortality in patients with COVID-19.9 However, data about outcomes in post COVID-19 short- and long-term follow-up period is limited. Therefore, the present study was designed to evaluate the effect of COVID-19 on hypertension in the short term post COVID-19 period. In the present study, 153 eligible COVID-19 patients enrolled and followed up 31.6 days on average. At the end of this period, systolic and diastolic BP was significantly increased. The incidence of new hypertension was also increased.

Various biomarkers and comorbidities have been identified as independent predictors of severe disease and adverse outcomes in COVID-19.1012 With respect to hypertension, its relation with COVID-19 has been discussed since the early stages of the pandemic. In a review by Tadic et al, a search of 14 studies was performed to determine the relationship between hypertension and COVID-19 and the role of hypertension on outcome in these patients. Tadic et al concluded that arterial hypertension represented one of the most common comorbidities in patients with COVID-19.13 Due to the role of angiotensin converting enzyme (ACE) 2 in SARS-CoV-2 infection, it was suggested that hypertension may be involved in the pathogenesis of COVID-19.13 In a recent study, Lippi et al found that hypertension is associated with a 2.5-fold increased risk of both increased disease severity and mortality in COVID-19 patients. They also showed that this effect was mainly observed in older patients (age >60 years).4 On the other hand, ACE 2, as a receptor for SARS-CoV-2, is increased in the use of ACE inhibitors or angiotensin (ANG) receptor blockers. Concerns have been raised over the risk of SARS-CoV-2 infection and poor prognosis of COVID-19 in patients who are on these drugs. Various studies focused on this issue.14,15 A review of 16 studies showed that the evidence does not suggest higher risks for SARS-CoV-2 infection or poor prognosis for COVID-19 patients treated with renin angiotensin aldosterone system (RAAS) inhibitors.16 The American Heart Association and European Society for Cardiology confirmed this issue.17,18

The RAAS plays a key role in the cardiovascular system.19 It is well known that the hyperactivation of RAAS and increases in ANG 2 levels are related with adverse outcomes (via the ANG 1 receptors) in cardiovascular diseases including heart failure, hypertension, myocardial infarction, and diabetic cardiovascular complications.20 On the other hand, ACE 2 is an enzyme has a negative regulator role in RAAS activation mainly by converting ANG 1 and ANG 2 into ANG 1–9 and ANG 1–7, respectively. There is a balance between the protective arm ACE 2/ANG 1–7/Mas receptor axis and pathogenic arm ACE/ANG 2/ANG 2 receptor type 1 receptor axis.21 ACE 2 is also the cellular receptor for the SARS-CoV-2 that is responsible the infectivity of COVID-19. ACE 2 is widely expressed in the cardiovascular system and in the lung, as well. Considering that ACE 2 plays a negative role in RAAS, a decrease in the ACE 2 and an increase in the ANG 2 level may lead to increase in BP. In a cohort study circulating, ANG 2 levels were significantly elevated in COVID-19 patients when compared with healthy individuals and increase of ANG 2 was linearly correlated with virus load.22 Therefore, a direct link between ACE 2 down regulation and systemic RAAS imbalance may lead to increase ANG 2 levels and BP. Accordingly, the present study showed that both systolic and diastolic BP were significantly increased in COVID-19 patients in short term follow-up period. The new onset hypertension was observed in 18 patients at the end of the follow-up period.

The effect of COVID-19 on BP has not been elucidated yet. However, few cases of hypertension after mRNA-based vaccination for COVID-19 have been reported. Athyros et al23 reported a hypertensive crisis with intracranial hemorrhage 3 days after anti–COVID-19 vaccination. In a case series, Meylan et al24 shared their 1-month experience in their vaccination center. They identified 9 patients with stage 3 hypertension after vaccination. In both reports,23,24 the authors stated that the underlying mechanism was uncertain. An analogy may be coagulopathy which can occur both during COVID-19 infection and after vaccination. Hematological and thromboembolic events were observed after first doses of mRNA vaccines. The risks of such events were higher and more prolonged after SARS-CoV-2 infection than after vaccination.25 The rise in BP after COVID-19 is also indirectly supported by the hypertension occurring after vaccination. More research is needed in order to confirm the occurrence of hypertension after mRNA-based vaccination and SARS-CoV-2 infection.

Stress and anxiety are the main reasons for white coat hypertension (WCH). In the present study, WCH was unlikely to affect the results because of several reasons. First, in the study population, stress or anxiety due to possible diagnosis of COVID-19 would be likely to be higher on admission. In the control visit, however, the patients knew that they had recovered from COVID-19 disease and therefore were likely to be in a better psychological condition. Despite better psychological status, both systolic and diastolic BP were significantly higher in the post COVID-19 period. Second, the first measurements and second measurements of BP were compared in the same patient. This, to some extent compensates for an anxious personality. In other words, if present, anxiety could be seen in the first as well as in the second measurement. Third, BP measurement was performed in a quiet room before the nasopharyngeal and blood sample collections. Therefore, no uncomfortable/painful procedure was applied before BP measurement. The present study has some limitations. First, the follow-up period was short. Further studies should be planned in order to investigate whether the causative effect of COVID-19 on hypertension is not evident after long term follow-up. Second, the results of the present study should be supported by the detection of biomarkers, including ANG 2 and ACE 2 levels. Third, the present study was a single-center experience and represents a small number of patients. However, our study population of unselected COVID-19 patients mirrors the real world scenario.

In conclusion, the present study showed that COVID-19 leads to increase both systolic and diastolic BP and causes new onset hypertension. The clinical implication of the present study is that, physicians should be aware of the potentially risk for new onset hypertension during the post COVID-19 period and take early action.

Acknowledgments

I thank all collaborators of the coronavirus disease 2019 unit in Parkhayat Kutahya Hospital. Special thanks to Zeynep Selcen Akpek, MD, for insightful comments.

Declaration of Conflicting Interests
The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding
The author(s) received no financial support for the research, authorship, and/or publication of this article.

ORCID iD

Mahmut Akpek https://orcid.org/0000-0002-2867-4993

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Close relatives of MERS-CoV in bats use ACE2 as their functional receptors

Authors: Qing Xiong,  View ORCID , , ei Cao,  Chengbao Ma,  Chen Liu, Junyu Si,  Peng Liu,  Mengxue Gu,  Chunli Wang, Lulu Shi, Fei Tong, Meiling Huang, Jing Li, Chufeng Zhao,  Chao Shen,   Yu Chen,   Huabin Zhao,  Ke Lan,  Xiangxi Wang,  Huan Yan

Summary

Middle East Respiratory Syndrome coronavirus (MERS-CoV) and several bat coronaviruses employ Dipeptidyl peptidase-4 (DPP4) as their functional receptors14. However, the receptor for NeoCoV, the closest MERS-CoV relative yet discovered in bats, remains enigmatic5. In this study, we unexpectedly found that NeoCoV and its close relative, PDF-2180-CoV, can efficiently use some types of bat Angiotensin-converting enzyme 2 (ACE2) and, less favorably, human ACE2 for entry. The two viruses use their spikes’ S1 subunit carboxyl-terminal domains (S1-CTD) for high-affinity and species-specific ACE2 binding. Cryo-electron microscopy analysis revealed a novel coronavirus-ACE2 binding interface and a protein-glycan interaction, distinct from other known ACE2-using viruses. We identified a molecular determinant close to the viral binding interface that restricts human ACE2 from supporting NeoCoV infection, especially around residue Asp338. Conversely, NeoCoV efficiently infects human ACE2 expressing cells after a T510F mutation on the receptor-binding motif (RBM). Notably, the infection could not be cross-neutralized by antibodies targeting SARS-CoV-2 or MERS-CoV. Our study demonstrates the first case of ACE2 usage in MERS-related viruses, shedding light on a potential bio-safety threat of the human emergence of an ACE2 using “MERS-CoV-2” with both high fatality and transmission rate.

Introduction

Coronaviruses (CoVs) are a large family of enveloped positive-strand RNA viruses classified into four genera: Alpha-, Beta-, Gamma- and Delta-CoV. Generally, Alpha and Beta-CoV can infect mammals such as bats and humans, while Gamma- and Delta-CoV mainly infect birds, occasionally mammals68. It is thought that the origins of most coronaviruses infecting humans can be traced back to their close relatives in bats, the most important animal reservoir of mammalian coronaviruses 910. Coronaviruses are well recognized for their recombination and host-jumping ability, which has led to the three major outbreaks in the past two decades caused by SARS-CoV, MERS-CoV, and the most recent SARS-CoV-2, respectively1114.

MERS-CoV belongs to the linage C of Beta-CoV (Merbecoviruses), which poses a great threat considering its high case-fatality rate of approximately 35%15. Merbecoviruses have also been found in several animal species, including camels, hedgehogs, and bats. Although camels are confirmed intermediate hosts of the MERS-CoV, bats, especially species in the family of Vespertilionidae, are widely considered to be the evolutionary source of MERS-CoV or its immediate ancestor16.

Specific receptor recognition of coronaviruses is usually determined by the receptor-binding domains (RBDs) on the carboxyl-terminus of the S1 subunit (S1-CTD) of the spike proteins17. Among the four well-characterized coronavirus receptors, three are S1-CTD binding ectopeptidases, including ACE2, DPP4, and aminopeptidase N (APN)11819. By contrast, the fourth receptor, antigen-related cell adhesion molecule 1(CEACAM1a), interacts with the amino-terminal domain (NTD) of the spike S1 subunit of the murine hepatitis virus2021. Interestingly, the same receptor can be shared by distantly related coronaviruses with structurally distinct RBDs. For example, the NL63-CoV (an alpha-CoV) uses ACE2 as an entry receptor widely used by many sarbecoviruses (beta-CoV linage B)22. A similar phenotype of cross-genera receptor usage has also been found in APN, which is shared by many alpha-CoVs and a delta-CoV (PDCoV)7. In comparison, DPP4 usage has only been found in merbecoviruses (beta-CoV linage C) such as HKU4, HKU25, and related strains24.

Intriguingly, many other merbecoviruses do not use DPP4 for entry and their receptor usage remains elusive, such as bat coronaviruses NeoCoV, PDF-2180-CoV, HKU5-CoV, and hedgehog coronaviruses EriCoV-HKU3152325. Among them, the NeoCoV, infecting Neoromicia capensis in South Africa, represents a bat merbecovirus that happens to be the closest relative of MERS-CoV (85% identity at the whole genome level)2627. PDF-2180-CoV, another coronavirus most closely related to NeoCoV, infects Pipisrellus hesperidus native to Southwest Uganda2328. Indeed, NeoCoV and PDF-2180-CoV share sufficient similarity with MERS-CoV across most of the genome, rendering them taxonomically the same viral species2729. However, their S1 subunits are highly divergent compared with MERS-CoV (around 43-45% amino acid similarity), in agreement with their different receptor preference23.

In this study, we unexpectedly found that NeoCoV and PDF-2180-CoV use bat ACE2 as their functional receptor. The cryo-EM structure of NeoCoV RBD bound with the ACE2 protein from Pipistrellus pipistrellus revealed a novel ACE2 interaction mode that is distinct from how human ACE2 (hACE2) interacts with the RBDs from SARS-CoV-2 or NL63. Although NeoCoV and PDF-2180-CoV cannot efficiently use hACE2 based on their current sequences, the spillover events of this group of viruses should be closely monitored, considering their human emergence potential after gaining fitness through antigenic drift.

Results

Evidence of ACE2 usage

To shed light on the relationship between merbecoviruses, especially NeoCoV and PDF-2180-CoV, we conducted a phylogenetic analysis of the sequences of a list of human and animal coronaviruses. Maximum likelihood phylogenetic reconstructions based on complete genome sequences showed that NeoCoV and PDF-2180-CoV formed sister clade with MERS-CoV (Fig. 1a). In comparison, the phylogenetic tree based on amino acid sequences of the S1 subunit demonstrated that NeoCoV and PDF-2180-CoV showed a divergent relationship with MERS-CoV but are closely related to the hedgehog coronaviruses (EriCoVs) (Fig. 1b). A sequence similarity plot analysis (Simplot) queried by MERS-CoV highlighted a more divergent region encoding S1 for NeoCoV and PDF-2180-CoV compared with HKU4-CoV (Fig. 1c). We first tested whether human DPP4 (hDPP4) could support the infection of several merbecoviruses through a pseudovirus entry assay30. The result revealed that only MERS-CoV and HKU4-CoV showed significantly enhanced infection of 293T-hDPP4. Unexpectedly, we detected a significant increase of entry of NeoCoV and PDF-2180-CoV in 293T-hACE2 but not 293T-hAPN, both of which are initially set up as negative controls (Fig. 1dExtended Data Fig.1).

Extended Data Figure 1

Extended Data Figure 1

Expression level of coronaviruses spike proteins used for pseudotyping.

Fig. 1

Fig. 1A clade of bat merbecoviruses can use ACE2 but not DPP4 for efficient entry.

a-b, Phylogenetical analysis of merbecoviruses (gray) based on whole genomic sequences (a) and S1 amino acid sequences (b). NL63 and 229E were set as outgroups. Hosts and receptor usage were indicated. c, Simplot analysis showing the whole genome similarity of three merbecoviruses compared with MERS-CoV. The regions that encode MERS-CoV proteins were indicated on the top. Dashed box: S1 divergent region. d, Entry efficiency of six merbecoviruses in 293T cells stably expressing hACE2, hDPP4, or hAPN. e-f, Entry efficiency of NeoCoV in cells expressing ACE2 from different bats. EGFP intensity (e); firefly luciferase activity (f). g-h, Cell-cell fusion assay based on dual-split proteins showing the NeoCoV spike protein mediated fusion in BHK-21 cells expressing indicated receptors. EGFP intensity (g), live-cell Renilla luciferase activity (h). i, Entry efficiency of six merbecoviruses in 293T cells stably expressing the indicated bat ACE2 or DPP4. Mean±SEM for di; Mean±SD for f, and h.(n=3). RLU: relative light unit.

To further validate the possibility of more efficient usage of bat ACE2, we screened a bat ACE2 cell library individually expressing ACE2 orthologs from 46 species across the bat phylogeny, as described in our previous study31(Extended Data Figs.2-3, Supplementary Table 1). Interestingly, NeoCoV and PDF-2180-CoV, but not HKU4-CoV or HKU5-CoV, showed efficient entry in cells expressing ACE2 from most bat species belonging to Vespertilionidae (vesper bats). In contrast, no entry or very limited entry in cells expressing ACE2 of humans or bats from the Yinpterochiroptera group (Fig. 1e-fExtended Data Fig.4). Consistent with the previous reports, the infection of NeoCoV and PDF-2180-CoV could be remarkably enhanced by an exogenous trypsin treatment28(Extended Data Fig.5). As indicated by the dual split protein (DSP)-based fusion assay 32, Bat37ACE2 triggers more efficient cell-cell membrane fusion than hACE2 in the presence of NeoCoV spike protein expression (Fig. 1g-h). Notably, the failure of the human or hedgehog ACE2 to support entry of EriCoV-HKU31 indicates that these viruses have a different receptor usage (Extended Data Fig.6). In agreement with a previous study2328, our results against the possibility that bat DPP4 act as a receptor for NeoCoV and PDF-2180-CoV, as none of the tested DPP4 orthologs, from the vesper bats whose ACE2 are highly efficient in supporting vial entry, could support a detectable entry of NeoCoV and PDF-2180-CoV (Fig. 1iExtended Data Fig.7). Infection assays were also conducted using several other cell types from different species, including a bat cell line Tb 1 Lu, ectopically expressing ACE2 or DPP4 from Bat40 (Antrozous pallidus), and each test yielded similar results (Extended Data Fig.8).

Extended Data Figure 2

Extended Data Figure 2

Receptor function of ACE2 from 46 bat species in supporting NeoCoV and PDF-2180-CoV entry.

Extended Data Figure 3

Extended Data Figure 3

The expression level of 46 bat ACE2 orthologs in 293T cells as indicated by immunofluorescence assay detecting the C-terminal 3×FLAG Tag.

Extended Data Figure 4

Extended Data Figure 4

Entry efficiency of PDF-2180-CoV (a-b), HKU4-CoV (c), and HKU5-CoV (d) pseudoviruses in 293T cells expressing different bat ACE2 orthologs

Extended Data Figure 5

Extended Data Figure 5

TPCK-trypsin treatment significantly boosted the entry efficiency of NeoCoV and PDF-2180-CoV on 293T cells expressing different ACE2 orthologs.

Extended Data Figure 6

Extended Data Figure 6

Hedgehog ACE2 (hgACE2) cannot support the entry of Ea-HedCoV-HKU31. (a) The expression level of ACE2 was evaluated by immunofluorescence detecting the C-terminal fused Flag tag. (b) Viral entry of SARS-CoV-2 and HKU31 into cells expressing hACE2 or hgACE2.

Extended Data Figure 7

Extended Data Figure 7

ACE2 and DPP4 receptor usage of different merbecoviruses. a, Western blot detected the expression levels of ACE2 and DPP4 orthologs in 293T cells.b, The intracellular bat ACE2 expression level by immunofluorescence assay detecting the C-terminal 3×FLAG-tag. c-d, Viral entry (c) and RBD binding (d) of different coronaviruses on 293T cells expressing different ACE2 and DPP4 orthologs.

Extended data Figure 8

Extended data Figure 8

NeoCoV and PDF2180-CoV infection of different cell types expressing either Bat40ACE2 or Bat40DPP4. The BHK-21, 293T, Vero E6, A549, Huh-7, and Tb 1 Lu were transfected with either Bat40ACE2 or Bat40DPP4. The expression and viral entry (GFP) (a) and luciferase activity (c) were detected at 16 hpi.

S1-CTD mediated species-specific binding

The inability of NeoCoV and PDF-2180-CoV to use DPP4 is consistent with their highly divergent S1-CTD sequence compared with the MERS-CoV and HKU4-CoV. We produced S1-CTD-hFc proteins (putative RBD fused to human IgG Fc domain) to verify whether their S1-CTDs are responsible for ACE2 receptor binding. The live-cell binding assay based on cells expressing various bat ACE2 showed a species-specific utilization pattern in agreement with the results of the pseudovirus entry assays (Fig. 2a). The specific binding of several representative bat ACE2 was also verified by flow-cytometry (Fig. 2b). We further determined the binding affinity by Bio-Layer Interferometry (BLI) analysis. The results indicated that both viruses bind to the ACE2 from Pipistrellus pipistrellus (Bat37) with the highest affinity (KD=1.98nM for NeoCoV and 1.29 nM for PDF-2180-CoV). In contrast, their affinities for hACE2 were below the detection limit of our BLI analysis (Fig. 2cExtended Data Fig.9). An enzyme-linked immunosorbent assay (ELISA) also demonstrated the strong binding between NeoCoV/PDF-2180-CoV S1-CTDs and Bat37ACE2, but not hACE2 (Fig. 2d). Notably, as the ACE2 sequences of the hosts of NeoCoV and PDF-2180-CoV are unknown, Bat37 represents the closest relative of the host of PDF-2180-CoV (Pipisrellus hesperidus) in our study. The binding affinity was further verified by competitive neutralization assays using soluble ACE2-ectodomain proteins or viral S1-CTD-hFc proteins. Again, the soluble Bat37ACE2 showed the highest activity to neutralize viral infection caused by both viruses (Fig. 2e-f). Moreover, NeoCoV-S1-CTD-hFc could also potently neutralize NeoCoV and PDF-2180-CoV infections of cells expressing Bat37ACE2 (Fig. 2g). We further demonstrated the pivotal role of S1-CTD in receptor usage by constructing chimeric viruses and testing them for altered receptor usage. As expected, batACE2 usage was changed to hDPP4 usage for a chimeric NeoCoV with CTD, but not NTD, sequences replaced by its MERS-CoV counterpart (Fig. 2h). These results confirmed that S1-CTD of NeoCoV and PDF-2180-CoV are RBDs for their species-specific interaction with ACE2.

Extended data Figure 9

Extended data Figure 9

BLI analysis of the binding kinetics of PDF-2180-CoV S1-CTD interacting with different ACE2 orthologs.

Fig. 2

Fig. 2S1-CTD of NeoCoV and PDF-2180-CoV was required for species-specific ACE2 binding.

a, Binding of NeoCoV-S1-CTD-hFc with 293T bat ACE2 cells via immunofluorescence detecting the hFc. b, Flow cytometry analysis of NeoCoV-S1-CTD-hFc binding with 293T cells expressing the indicated ACE2. The positive ratio was indicated based on the threshold (dash line). c, BLI assays analyzing the binding kinetics between NeoCoV-S1-CTD-hFc with selected ACE2-ecto proteins. d, ELISA assay showing the binding efficiency of NeoCoV and PDF-2180-CoV S1-CTD to human and Bat37ACE2-ecto proteins. e, The inhibitory activity of soluble ACE2-ecto proteins against NeoCoV infection in 293T-Bat37ACE2. f, Dose-dependent competition of NeoCoV infection by Bat37ACE2-ecto proteins in 293T-Bat37ACE2 cells. g, The inhibitory effect of NeoCoV, PDF-2180-CoV S1-CTD-hFc and MERS-CoV RBD-hFC proteins on NeoCoV infection in 293T-Bat37ACE2 cells. h, Receptor preference of chimeric viruses with S1-CTD or S1-NTD swap mutations in cells expressing the indicated receptors. Mean±SD for deg, and h, (n=3).

Structural basis of ACE2 binding

To unveil the molecular details of the virus-ACE2 binding, we then carried out structural investigations of the Bat37ACE2 in complex with the NeoCoV and PDF-2180-CoV RBD. 3D classification revealed that the NeoCoV-Bat37ACE2 complex primarily adopts a dimeric configuration with two copies of ACE2 bound to two RBDs, whereas only a monomeric conformation was observed in the PDF-2180-CoV-Bat37ACE2 complex (Figs. 3a-bExtended Data Fig. 1011). We determined the structures of these two complexes at a resolution of 3.5 Å and 3.8 Å, respectively, and performed local refinement to further improve the densities around the binding interface, enabling reliable analysis of the interaction details (Figs. 3a-bExtended Data Fig. 1213 and Table 1-2). Despite existing in different oligomeric states, the structures revealed that both NeoCoV and PDF-2180-CoV recognized the Bat37ACE2 in a very similar way. We used the NeoCoV-Bat37ACE2 structure for detailed analysis (Figs. 3a-b and Extended Data Fig. 14). Like other structures of homologs, the NeoCoV RBD structure comprises a core subdomain located far away from the engaging ACE2 and an external subdomain recognizing the receptor (Fig. 3c and Extended Data Fig. 15). The external subdomain is a strand-enriched structure with four anti-parallel β strands (β6–β9) and exposes a flat four-stranded sheet-tip for ACE2 engagement (Fig. 3c). By contrast, the MERS-CoV RBD recognizes the side surface of the DPP4 β-propeller via its four-stranded sheet-blade (Fig. 3c). The structural basis for the differences in receptor usage can be inferred from two features: i) the local configuration of the four-stranded sheet in the external domain of NeoCoV shows a conformational shift of η3 and β8 disrupting the flat sheet-face for DPP4 binding and ii) relatively longer 6-7 and 8-9 loops observed in MERS-CoV impair their binding in the shallow cavity of bat ACE2 (Fig. 3c and Extended Data Fig. 15).

Extended data Figure 10

Extended data Figure 10

Flowcharts for cryo-EM data processing of Neo-CoV RBD-Bat37ACE2 complex.

Extended data Figure 11

Extended data Figure 11

Flowcharts for cryo-EM data processing of PDF-2180-CoV RBD-Bat37ACE2 complex.

Extended Data Figure 12

Extended Data Figure 12

Resolution Estimation of the EM maps, density maps, and atomics models of NeoCoV RBD-Bat37ACE2 complex.

Extended Data Figure 13

Extended Data Figure 13

Resolution Estimation of the EM maps, density maps, and atomics models of PDF-2180-CoV RBD-Bat37ACE2 complex.

Extended Data Figure 14

Extended Data Figure 14

Superimposition of overall structures of NeoCoV RBD-Bat37ACE2 complex (red) and PDF-2018-COV RBD-Bat37ACE2 complex (bule).

Extended Data Figure 15

Extended Data Figure 15

Structures and sequence comparison of RBDs from different merbecoviruses.

Fig. 3

Fig. 3Structure of the NeoCoV RBD-Bat37ACE2 and PDF-2018-CoV RBD-Bat37ACE2 complex.

ab, Cryo-EM density map and cartoon representation of NeoCoV RBD-Bat37ACE2 complex (a) and PDF-2018CoV RBD-Bat37ACE2 complex (b). The NeoCoV RBD, PDF-2180-CoV RBD, and Bat37ACE2 were colored by red, yellow, and cyan, respectively. c, Structure comparison between NeoCoV RBD-Bat37ACE2 complex (left) and MERS-CoV RBD-hDPP4 complex (right). The NeoCoV RBD, MERS-CoV RBD, NeoCoV RBM, MERS-CoV RBM, Bat37ACE2, and hDPP4 were colored in red, light green, light yellow, gray, cyan, and blue, respectively. d, Details of the NeoCOV RBD-Bat37ACE2 complex interface. All structures are shown as ribbon with the key residues shown with sticks. The salt bridges and hydrogen bonds are presented as red and yellow dashed lines, respectively. ef, Verification of the critical residues on NeoCoV RBD affecting viral binding (e), and entry efficiency (f) in 293T-Bat37ACE2 cells. gh, Verification of the critical residues on Bat37ACE2 affecting NeoCoV RBD binding (g), and viral entry efficiency(h). Mean±SD for f (n=3) and h (n=4).

In the NeoCoV-Bat37ACE2 complex structure, relatively smaller surface areas (498 Å2 in NeoCoV RBD and 439 Å2 in Bat37ACE2) are buried by the two binding entities compared to their counterparts in the MERS-CoV-DPP4 complex (880 Å2 in MERS-CoV RBD and 812 Å2 in DPP4; 956 Å2 in SARS-CoV-2 RBD and 893 Å2 in hACE2). The NeoCoV RBD inserts into an apical depression constructed by α11, α12 helices and a loop connecting α12 and β4 of Bat37ACE2 through its four-stranded sheet tip (Fig. 3d and Extended Data Table. 2). Further examination of the binding interface revealed a group of hydrophilic residues at the site, forming a network of polar-contacts (H-bond and salt-bridge) network and hydrophobic interactions. These polar interactions are predominantly mediated by the residues N504, N506, N511, K512, and R550 from the NeoCoV RBM and residues T53, E305, T334, D338, R340 from Bat37ACE2 (Fig. 3d, Extended Data Table. 2). Notably, the methyl group from residues A509 and T510 of the NeoCoV RBM are partially involved in forming a hydrophobic pocket with residues F308, W328, L333, and I358 from Bat37ACE2 at the interface. A substitution of T510 with F in the PDF-2180-CoV RBM further improves hydrophobic interactions, which is consistent with an increased binding affinity observed for this point mutation (Figs. 3d, Extended Data Table. 2). Apart from protein-protein contacts, the glycans of bat ACE2 at positions N54 and N329 sandwich the strands (β8–β9), forming π-π interactions with W540 and hydrogen bonds with N532, G545, and R550 from the NeoCoV RBD, underpinning virus-receptor associations (Fig. 3d and Extended Data Table. 2).

The critical residues were verified by introducing mutations and testing their effect on receptor binding and viral entry. As expected, mutations N504F/N506F, N511Y, and R550N in the NeoCoV RBD, abolishing the polar-contacts or introducing steric clashes, resulted in a significant reduction of RBD binding and viral entry (Fig.3e-f). Similarly, E305K mutation in Bat37ACE2 eliminating the salt-bridge also significantly impaired the receptor function. Moreover, the loss of function effect of mutation N54A on Bat37ACE2 abolishing the N-glycosylation at residue 54 confirmed the importance of the particular protein-glycan interaction in viral-receptor recognition. In comparison, N329A abolishing the N-glycosylation at site N329, located far away from the binding interface, had no significant effect on receptor function (Fig.3g-h).

Evaluation of zoonotic potential

A major concern is whether NeoCoV and PDF-2180-CoV could jump the species barrier and infect humans. As mentioned above, NeoCoV and PDF-2180-CoV cannot efficiently interact with human ACE2. Here we first examined the molecular determinants restricting hACE2 from supporting the entry of these viruses. By comparing the binding interface of the other three hACE2-using coronaviruses, we found that the SARS-CoV, SARS-COV-2, and NL63 share similar interaction regions that barely overlapped with the region engaged by NeoCoV (Fig. 4a). Analysis of the overlapped binding interfaces reveals a commonly used hot spot around residues 329-330 (Fig.4b). Through sequences alignment and structural analysis of hACE2 and Bat37ACE2, we predicted that the inefficient use of the hACE2 for entry by the viruses could be attributed to incompatible residues located around the binding interfaces, especially the difference in sequences between residues 337-342 (Fig.4c). We replaced these residues of hACE2 with those from the Bat37ACE2 counterparts to test this hypothesis (Fig.4c-d). The substitution led to an approximately 15-fold and 30-fold increase in entry efficiency of NeoCoV and PDF-2180-CoV, respectively, confirming that this region is critical for the determination of the host range. Further fine-grained dissection revealed that N338 is the most crucial residue in restricting human receptor usage (Fig. 4e-g).

Fig. 4

Fig. 4Molecular determinants affecting hACE2 recognition by the viruses.

a, Binding modes of ACE2-adapted coronaviruses. The SARS-CoV RBD, SARS-CoV-2 RBD, NL63-CoV RBD, and NeoCoV RBD were colored in purple, light purple, green, and red, respectively. b, A common virus-binding hot spot on ACE2 for the four viruses. Per residue frequency recognized by the coronavirus RBDs were calculated and shown. c, Schematic illustration of the hACE2 swap mutants with Bat37ACE2 counterparts. de, The expression level of the hACE2 mutants by Western blot (d) and immunofluorescence (e). fg, Receptor function of hACE2 mutants evaluated by virus RBD binding assay (f) and pseudovirus entry assay (g). h, Molecular dynamics (MD) analysis of the effect of critical residue variations on the interaction between NeoCoV and Bat37ACE2 by mCSM-PPI2. i, Structure of NeoCoV RBD-hACE2 complex modeling by superposition in COOT. The NeoCoV RBD and hACE2 were colored in red and sky blue, respectively. Details of the NeoCoV RBD key mutation T510F was shown. All structures are presented as ribbon with the key residues shown with sticks. j-k, The effect of NeoCoV and PDF-2180-CoV RBM mutations on hACE2 fitness as demonstrated by binding (j) and entry efficiency (k) on 293T-hACE2 and 293T-Bat37ACE2 cells. l, hACE2 dependent entry of NeoCoV-T510F in Caco2 cells in the presence of 50μg/ml of Anti-ACE2 (H11B11) or Anti-VSVG (I1). m, Neutralizing activity of SARS-CoV-2 vaccinated sera against the infection by SARS-CoV-2, NeoCoV, and PDF-2180-CoV. n, Neutralizing activity of MERS-RBD targeting nanobodies against the infections by MERS-CoV, NeoCoV, and PDF-2180-CoV. Mean± SD for g,k-ng(n=4),kl (n=3), gmn (n=10).

We further assessed the zoonotic potential of NeoCoV and PDF-2180-CoV by identifying the molecular determinants of viral RBM, which might allow cross-species transmission through engaging hACE2. After meticulously examining the critical residues based on the complex structures and computational prediction tool mCSM-PPI233(Fig. 4h, Extended Data Table. 4), we predicted increasing hydrophobicity around the residue T510 of NeoCoV might enhance the virus-receptor interaction on hACE2 (Fig. 4 i). Interestingly, the PDF-2180-CoV already has an F511 (corresponding to site 510 of NeoCoV), which is consistent with its slightly higher affinity to human ACE2 (Extended Data Fig.16). As expected, T510F substitutions in NeoCoV remarkably increased its binding affinity with hACE2 (KD=16.9 nM) and a significant gain of infectivity in 293T-hACE2 cells (Fig. 4 j-k, Extended Data Fig.17-18). However, PDF-2180-CoV showed much lower efficiency in using hACE2 than NeoCoV-T510F, indicating other unfavorable residues are restricting its efficient interaction with hACE2. Indeed, a G to A (corresponding to A509 in NeoCoV) mutation in site 510, increasing the local hydrophobicity, partially restored its affinity to hACE2 (Fig.4 j-k). In addition, the NeoCoV-T510F can enter the human colon cell line Caco-2 with much higher efficiency than wild-type NeoCoV. It enters the Caco-2 exclusively through ACE2 as the infection can be neutralized by an ACE2-targeting neutralizing antibody H11B1134 (Fig. 4l). Humoral immunity triggered by prior infection or vaccination of other coronaviruses might be inadequate to protect humans from NeoCoV and PDF-2180-CoV infections because neither SARS-CoV-2 anti-sera nor ten tested anti-MERS-CoV nanobodies can cross-inhibit the infection caused by these two viruses35. (Fig. 4m-n).

Extended Data Figure 16

Extended Data Figure 16

Comparison of the binding affinity of NeoCoV and PDF-2180-CoV RBD with hACE2 using SARS-CoV-2 RBD as a positive control.

Extended Data Figure 17

Extended Data Figure 17

Expression level of the NeoCoV and PDF-2180-CoV spike proteins and their mutants.

Extended Data Figure 18

Extended Data Figure 18

BLI analysis of the binding kinetics of NeoCoV S1-CTD WT and T510F interacting with human ACE2.

Discussion

The lack of knowledge of the receptors of bat coronaviruses has greatly limited our understanding of these high-risk pathogens. Our study provided evidence that the relatives of potential MERS-CoV ancestors like NeoCoV and PDF-2180-CoV engage bat ACE2 for efficient cellular entry. However, HKU5-CoV and EriCoV seem not to use bat DPP4 or hedgehog ACE2 for entry, highlighting the complexity of coronaviruses receptor utilization. It was unexpected that NeoCoV and PDF-2180-CoV use ACE2 rather than DPP4 as their entry receptors since their RBD core structures resemble MERS-CoV more than other ACE2-using viruses (Fig. 4aExtended Data Fig. 15).

Different receptor usage can affect the transmission rate of the viruses. Although it remains unclear whether ACE2 usage out-weight DPP4 usage for more efficient transmission, MERS-CoV appears to have lower transmissibility with an estimated R0 around 0.69. Comparatively, the ACE2 usage has been approved able to achieve high transmissibility. The SARS-CoV-2 estimated R0 is around 2.5 for the original stain, 5.08 for the delta variant, and even higher for the omicron variant3638. This unexpected ACE2 usage of these MERS-CoV close relatives highlights a latent biosafety risk, considering a combination of two potentially damaging features of high fatality observed for MERS-CoV and the high transmission rate noted for SARS-CoV-2. Furthermore, our studies show that the current COVID-19 vaccinations are inadequate to protect humans from any eventuality of the infections caused by these viruses.

Many sarbecoviruses, alpha-CoV NL63, and a group of merbecoviruses reported in this study share ACE2 for cellular entry. Our structural analysis indicates NeoCoV and PDF-2180-CoV bind to an apical side surface of ACE2, which is different from the surface engaged by other ACE2-using coronaviruses (Fig.4a). The interaction is featured by inter-molecular protein-glycan bonds formed by the glycosylation at N54, which is not found in RBD-receptor interactions of other coronaviruses. The different interaction modes of the three ACE2-using coronaviruses indicate a history of multiple independent receptor acquisition events during evolution22. The evolutionary advantage of ACE2 usage in different CoVs remains enigmatic.

Our results support the previous hypothesis that the origin of MERS-CoV might be a result of an intra-spike recombination event between a NeoCoV like virus and a DPP4-using virus26. RNA recombination can occur during the co-infection of different coronaviruses, giving rise to a new virus with different receptor usage and host tropisms3940. It remains unclear whether the event took place in bats or camels, and where the host switching happened. Although bat merbecoviruses are geographically widespread, the two known ACE2-using merbecoviruses are inhabited in Africa. Moreover, most camels in the Arabian Peninsula showing serological evidence of previous MERS-CoV infection are imported from the Greater Horn of Africa with several Neoromicia species41. Considering both viruses are inefficient in infecting human cells in their current form, the acquisition of the hDPP4 binding domain would be a critical event driving the emergence of MERS-CoV. Further studies will be necessary to obtain more evidence about the origin of MERS-CoV.

The host range determinants on ACE2 are barriers for cross-species transmission of these viruses. Our results show NeoCoV and PDF-2180-CoV favor ACE2 from bats of the Yangochiroptera group, especially vesper bats (Vespertilionidae), where their host belongs to, but not ACE2 orthologs from bats of the Yinpterochiroptera group. Interestingly, most merbecoviruses were found in species belonging to the Vespertilionidae group, a highly diverse and widely distributed family9. Although the two viruses could not use hACE2 efficiently, our study also reveals that single residue substitution increasing local hydrophobicity around site 510 could enhance their affinity for hACE2 and enable them to infect human cells expressing ACE2. Considering the extensive mutations in the RBD regions of the SARS-CoV-2 variants, especially the heavily mutated omicron variant, these viruses may hold a latent potential to infect humans through further adaptation via antigenic drift4243. It is also very likely that their relatives with human emergence potential are circulating somewhere in nature.

Overall, we identified ACE2 as a long-sought functional receptor of the potential MERS-CoV ancestors in bats, facilitating the in-depth research of these important viruses with zoonotic emergence risks. Our study adds to the knowledge about the complex receptor usage of coronaviruses, highlighting the importance of surveillance and research on these viruses to prepare for potential outbreaks in the future.

Supplementary Information

Methods

Receptor and virus sequences

The acquisition of sequences of 46 bat ACE2 and hACE were described in our previous study31. The five bat DPP4 and hDPP4 sequences were directly retrieved from the GenBank database (human DPP4, NM_001935.3; Bat37, Pipistrellus pipistrellus, KC249974.1) or extracted from whole genome sequence assemblies of the bat species retrieved from the GenBank database (Bat25, Sturnira hondurensis, GCA_014824575.2; Bat29, Mormoops blainvillei, GCA_004026545.1; Bat36, Aeorestes cinereus, GCA_011751065.1; Bat40, Antrozous pallidus, GCA_007922775.1). The whole genome sequences of different coronaviruses were retrieved from the GenBank database. The accession numbers are as follows: MERS-CoV (JX869059.2), Camel MERS-CoV KFU-HKU 19Dam (KJ650296.1), HKU4 (NC_009019.1), HKU5 (NC_009020.1), ErinaceusCoV/HKU31 strain F6 (MK907286.1), NeoCoV (KC869678.4), PDF-2180-CoV (NC_034440.1), ErinaceusCoV/2012-174 (NC_039207.1), BtVs-BetaCoV/SC2013 (KJ473821.1), BatCoV/H.savii/Italy (MG596802.1), BatCoV HKU25 (KX442564.1), BatCoV ZC45(MG772933.1) and SARS-CoV-2 (NC_045512.2), NL63 (JX504050.1) 229E (MT797634.1).

All gene sequences used in this study were commercially synthesized by Genewiz. The sources, accession numbers, and sequences of the receptors and viruses were summarized in Supplementary Table 1.

SARS-CoV-2 anti-sera collection

All the vaccinated sera were collected from volunteers at about 21 days post the third dose of the WHO-approved inactivated SARS-COV-2 vaccine (CorovaVac, Sinovac, China). All volunteers were provided informed written consent forms, and the whole study was conducted following the requirements of Good Clinical Practice of China.

Bioinformatic analysis

Protein sequence alignment was performed using the MUSCLE algorithm by MEGA-X software (version 10.1.8). For phylogenetic analysis, nucleotide or protein sequences of the viruses were first aligned using the Clustal W and the MUSCLE algorithm, respectively. Then, the phylogenetic trees were generated using the maximal likelihood method in MEGA-X (1000 Bootstraps). The model and the other parameters used for phylogenetic analysis were applied following the recommendations after finding best DNA/Protein Models by the software. The nucleotide similarity of coronaviruses was analyzed by SimPlot software (version 3.5.1) with a slide windows size of 1000 nucleotides and a step size of 100 nucleotides using gap-stripped alignments and the Kimura (2-parameter) distance model.

Plasmids

Human codon-optimized sequences of various ACE2 or DPP4 orthologs and their mutants were cloned into a lentiviral transfer vector (pLVX-IRES-puro) with a C-terminal 3×Flag tag (DYKDHD-G-DYKDHD-I-DYKDDDDK). The DNA sequences of human codon-optimized NeoCoV S protein (AGY29650.2), PDF-2180-CoV S protein (YP_009361857.1), HKU4-CoV S protein (YP_001039953.1), HKU5-CoV S protein (YP_001039962.1), HKU31 S protein (QGA70692.1), SARS-CoV-2 (YP_009724390.1), and MERS-CoV S protein (YP_009047204.1) were cloned into the pCAGGS vector with a C-terminal 13-15-amino acids deletion (corresponding to 18 amino-acids in SARS-CoV-2) or replacement by an HA tag (YPYDVPDYA) for higher VSV pseudotyping efficiency44. The plasmids expressing coronavirus RBD-IgG-hFc fusion proteins were generated by inserting the coding sequences of NeoCoV RBD (aa380-585), PDF-2180-CoV RBD (aa381-586), HKU4-CoV (aa382-593), HKU5-CoV RBD (aa385-586), HKU31-CoV RBD (aa366-575), SARS-CoV-2 RBD (aa331-524) and MERS-CoV RBD (aa377-588) into the pCAGGS vector with an N-terminal CD5 secretion leading sequence (MPMGSLQPLATLYLLGMLVASVL). The plasmids expressing soluble bat ACE2 and DPP4 proteins were constructed by inserting the ectodomain coding sequences into the pCAGGS vector with N-terminal CD5 leader sequence and C-terminal twin-strep tag and 3×Flag tag tandem sequences (WSHPQFEKGGGSGGGSGGSAWSHPQFEK-GGGRS-DYKDHDGDYKDHDIDYKDDDDK).

Virus spike proteins or receptor-related mutants or chimeras were generated by overlapping PCR. For Dual split protein (DSP) based cell-cell fusion assay, the dual reporter split proteins were expressed by pLVX-IRES-puro vector expressing the RLucaa1-155-GFP1-7(aa1-157) and GFP8-11(aa158-231)-RLuc-aa156-311 plasmids, which were constructed in the lab based on a previously study3245. The plasmids expressing the codon-optimized anti-ACE2 antibody (H11B11; GenBank accession codes MZ514137 and MZ514138) were constructed by inserting the heavy-chain and light-chain coding sequences into the pCAGGS vector with N-terminal CD5 leader sequences, respectively34. For anti-MERS-CoV nanobody-hFc fusion proteins, nanobody coding sequences were synthesized and cloned into the pCAGGS vector with N-terminal CD5 leader sequences and C-terminal hFc tags 35.

Protein expression and purification

The RBD-hFc (S1-CTD-hFc) fusion proteins of SARS-CoV-2, MERS-CoV, HKU4-CoV, HKU5-CoV, HKU31-CoV, NeoCoV, and PDF-2180-CoV, and the soluble ACE2 proteins of human, Bat25, Bat29, Bat36, Bat37, Bat38, and Bat40 were expressed by 293T by transfecting the corresponding plasmids by GeneTwin reagent (Biomed, TG101-01) following the manufacturers’ instructions. Four hrs post-transfection, the culture medium of the transfected cells was replenished by SMM 293-TII Expression Medium (Sino Biological, M293TII). The supernatant of the culture medium containing the proteins was collected every 2-3 days. The recombinant RBD-hFc proteins were captured by Pierce Protein A/G Plus Agarose (Thermo Scientific, 20424), washed by wash buffer W (100 mM Tris/HCl, pH 8.0, 150 mM NaCl, 1mM EDTA), eluted with pH 3.0 Glycine buffer (100mM in H2O), and then immediately balanced by UltraPure 1M Tris-HCI, pH 8.0 (15568025, Thermo Scientific). The twin-strep tag containing proteins were captured by Strep-Tactin XT 4Flow high capacity resin (IBA, 2-5030-002), washed by buffer W, and eluted with buffer BXT (100 mM Tris/HCl, pH 8.0, 150 mM NaCl, 1mM EDTA, 50mM biotin). The eluted proteins can be concentrated and buffer-changed to PBS through ultra-filtration. Protein concentrations were determined by Omni-Easy Instant BCA Protein Assay Kit (Epizyme, ZJ102). The purified proteins were aliquoted and stored at -80℃. For Cryo-EM analysis, NeoCoV RBD (aa380-588), PDF-2018-CoV RBD (381-589), and Bat37ACE2 (aa21-730) were synthesized and subcloned into the vector pCAGGS with a C-terminal twin-strep tag. Briefly, these proteins were expressed by transient transfection of 500 ml HEK Expi 293F cells (Gibco, Thermo Fisher, A14527) using Polyethylenimine Max Mw 40,000 (polysciences). The resulting protein samples were further purified by size-exclusion chromatography using a Superdex 75 10/300 Increase column (GE Healthcare) or a Superdex 200 10/300 Increase column (GE Healthcare) in 20mM HEPES, 100 mM NaCl, pH 7.5. For RBD-receptor complex (NeoCoV RBD-Bat37ACE2 / PDF-2180-CoV RBD-Bat37ACE2), NeoCoV RBD or PDF-2180-CoV RBD was mixed with Bat37ACE2 at the ratio of 1.2 :1, incubated for 30 mins on ice. The mixture was then subjected to gel filtration chromatography. Fractions containing the complex were collected and concentrated to 2 mg/ml.

Cell culture

293T (CRL-3216), VERO E6 cells (CRL-1586), A549 (CCL-185), BHK-21 (CCL-10), and Huh-7 (PTA-4583), Caco2 (HTB-37) and the epithelial bat cell line Tb 1 Lu (CCL-88) were purchased from American Type Culture Collection (ATCC) and cultured in Dulbecco’s Modified Eagle Medium, (DMEM, Monad, China) supplemented with 10% fetal bovine serum (FBS), 2.0 mM of L-glutamine, 110 mg/L of sodium pyruvate and 4.5 g/L of D-glucose. An I1-Hybridoma (CRL-2700) cell line secreting a neutralizing mouse monoclonal antibody against the VSV glycoprotein (VSVG) was cultured in Minimum Essential Medium with Earles’s balances salts and 2.0mM of L-glutamine (Gibico) and 10% FBS. All cells were cultures at 37℃ in 5% CO2 with the regular passage of every 2-3 days. 293T stable cell lines overexpressing ACE2 or DPP4 orthologs were maintained in a growth medium supplemented with 1 μg/ml of puromycin.

Stable cell line generation

Stable cell lines overexpressing ACE2 or DPP4 orthologs were generated by lentivirus transduction and antibiotic selection. Specifically, the lentivirus carrying the target gene was produced by cotransfection of lentiviral transfer vector (pLVX-EF1a-Puro, Genewiz) and packaging plasmids pMD2G (Addgene, plasmid no.12259) and psPAX2 (Addgene, plasmid no.12260) into 293T cells through Lip2000 Transfection Reagent (Biosharp, BL623B). The lentivirus-containing supernatant was collected and pooled at 24 and 48 hrs post-transfection. 293T cells were transduced by the lentivirus after 16 hrs in the presence of 8 μg/ml polybrene. Stable cells were selected and maintained in the growth medium with puromycin (1-2 μg/ml). Cells selected for at least ten days were considered stable cell lines and used in different experiments.

Cryo-EM sample preparation and data collection

For Cryo-EM sample preparation, the NeoCoV RBD-Bat37ACE2 or PDF-2018-CoV RBD-Bat37ACE2 complex was diluted to 0.5 mg/ml. Holy-carbon gold grid (Cflat R1.2/1.3 mesh 300) were freshly glow-discharged with a Solarus 950 plasma cleaner (Gatan) for 30s. A 3 μL aliquot of the mixture complex was transferred onto the grids, blotted with filter paper at 16℃ and 100% humidity, and plunged into the ethane using a Vitrobot Mark IV (FEI). For these complexes, micrographs were collected at 300 kV using a Titan Krios microscope (Thermo Fisher), equipped with a K2 detector (Gatan, Pleasanton, CA), using SerialEM automated data collection software. Movies (32 frames, each 0.2 s, total dose 60e−Å−2) were recorded at a final pixel size of 0.82 Å with a defocus of between -1.2 and -2.0 μm.

Image processing

For NeoCoV RBD-Bat37ACE2 complex, a total of 4,234 micrographs were recorded. For PDF-2018-CoV RBD-Bat37ACE2 complex, a total of 3,298 micrographs were recorded. Both data sets were similarly processed. Firstly, the raw data were processed by MotionCor2, which were aligned and averaged into motion-corrected summed images. Then, the defocus value for each micrograph was determined using Gctf. Next, particles were picked and extracted for two-dimensional alignment. The well-defined partial particles were selected for initial model reconstruction in Relion46. The initial model was used as a reference for three-dimensional classification. After the refinement and post-processing, the overall resolution of PDF-2018-CoV RBD-Bat37ACE2 complex was up to 3.8Å based on the gold-standard Fourier shell correlation (threshold = 0.143) 47. For the NeoCoV RBD-Bat37ACE2 complex, the C2 symmetry was expanded before the 3D refinement. Finally, the resolution of the NeoCoV RBD-Bat37ACE2 complex was up to 3.5Å. The quality of the local resolution was evaluated by ResMap48.

Model building and refinement

The NeoCoV RBD-Bat37ACE2 complex structures were manually built into the refined maps in COOT47. The atomic models were further refined by positional and B-factor refinement in real space using Phenix48. For the PDF-2018-CoV RBD-Bat37ACE2 complex model building, the refinement NeoCoV RBD-Bat37ACE2 complex structures were manually docked into the refined maps using UCSF Chimera and further corrected manually by real-space refinement in COOT. As the same, the atomic models were further refined by using Phenix. Validation of the final model was performed with Molprobity48. The data sets and refinement statistics are shown in Extended Data table 1.

Immunofluorescence assay

The expression levels of ACE2 or DPP4 receptors were evaluated by immunofluorescence assay detecting the C-terminal 3×FLAG-tags. The cells expressing the receptors were seeded in the 96-well plate (poly-lysine pretreated plates for 293T based cells) at a cell density of 1∼5×105/ml (100 μl per well) and cultured for 24 hrs. Cells were fixed with 100% methanol at room temperature for 10 mins, and then incubated with a mouse monoclonal antibody (M2) targeting the FLAG-tag (Sigma-Aldrich, F1804) diluted in 1% BSA/PBS at 37℃ for 1 hour. After one wash with PBS, cells were incubated with 2 μg/ml of the Alexa Fluor 594-conjugated goat anti-mouse IgG (Thermo Fisher Scientific, A32742) diluted in 1% BSA/PBS at room temperature for 1 hour. The nucleus was stained blue with Hoechst 33342 (1:5,000 dilution in PBS). Images were captured with a fluorescence microscope (Mshot, MI52-N).

Pseudovirus production and titration

Coronavirus spike protein pseudotyped viruses (CoV-psV) were packaged according to a previously described protocol based on a replication-deficient VSV-based rhabdoviral pseudotyping system (VSV-dG). The VSV-G glycoprotein-deficient VSV coexpressing GFP and firefly luciferase (VSV-dG-GFP-fLuc) was rescued by a reverse genetics system in the lab and helper plasmids from Karafast. For CoV-psV production, 293T or Vero E6 cells were transfected with the plasmids overexpressing the coronavirus spike proteins through the Lip2000 Transfection Reagent (Biosharp, BL623B). After 36 hrs, the transfected cells were transduced with VSV-dG-GFP-fLuc diluted in DMEM for 4 hrs at 37℃ with a 50 % tissue culture infectious dose (TCID50) of 1×106 TCID50/ml. Transduced cells were washed once with DMEM and then incubated with culture medium and I1-hybridoma-cultured supernatant (1:10 dilution) containing VSV neutralizing antibody to eliminate the infectivity of the residual input viruses. The CoV-psV-containing supernatants were collected at 24 hrs after the medium change, clarified at 4,000 rpm for 5 mins, aliquoted, and stored at -80℃. The TCID50 of pseudovirus was determined by a serial-dilution based infection assay on 293T-bat40ACE2 cells for NeoCoV and PDF-2180-CoV or 293T-hDpp4 cells for MERS-CoV and HKU4-CoV. The TCID50 was calculated according to the Reed-Muench method 4950. The relative luminescence unit (RLU) value ≥ 1000 is considered positive. The viral titer (genome equivalents) of HKU5-COV and HKU31-CoV without an ideal infection system was determined by quantitative PCR with reverse transcription (RT–qPCR). The RNA copies in the virus-containing supernatant were detected using primers in the VSV-L gene sequences (VSV-L-F: 5’-TTCCGAGTTATGGGCCAGTT-3’; VSVL-R: 5’-TTTGCCGTAGACCTTCCAGT-3’).

Pseudovirus entry assay

Cells for infection were trypsinized and incubated with different pseudoviruses (1×105 TCID50/well, or same genome equivalent) in a 96-well plate (5×104 /well) to allow attachment and viral entry simultaneously. For TPCK-trypsin treatment for infection boosting, NeoCoV and PDF-2180-CoV pseudovirus in serum-free DMEM were incubated with 100 μg/ml TPCK-treated trypsin (Sigma-Aldrich, T8802) for 10 mins at 25℃, and then treated with 100 μg/ml soybean trypsin inhibitor (Sigma-Aldrich, T6414) in DMEM+10% FBS to stop the proteolysis. At 16 hours post-infection (hpi), GFP images of infected cells were acquired with a fluorescence microscope (Mshot, MI52-N), and intracellular luciferase activity was determined by a Bright-Glo Luciferase Assay Kit (Promega, E2620) and measured with a SpectraMax iD3 Multi-well Luminometer (Molecular Devices) or a GloMax 20/20 Luminometer (Promega).

Pseudovirus neutralization Assay

For antibody neutralization assays, the viruses (2×105 TCID50/well) were incubated with the sera (50-fold diluted) or 10 μg/ml MERS-specific nanobodies at 37℃ for 30 mins, and then mixed with trypsinized BHK-21-Bat37ACE2 cells with the density of 2×104/well. After 16 hrs, the medium of the infected cells was removed, and the cells were lysed with 1×Bright-Glo Luciferase Assay reagent (Promega) for chemiluminescence detection with a SpectraMax iD3 Multi-well Luminometer (Molecular devices).

Western blot

After one wash with PBS, the cells were lysed by 2% TritonX-100/PBS containing 1mM fresh prepared PMSF (Beyotime, ST506) on ice for 10 mins. Then cell lysate was clarified by 12,000 rpm centrifugation at 4℃ for 5 mins, mixed with SDS loading buffer, and then incubated at 95 °C for 5 mins. After SDS-PAGE electrophoresis and PVDF membrane transfer, the membrane was blocked with 5% skim milk/PBST at room temperature for one hour, incubated with primary antibodies against Flag (Sigma, F1804), HA (MBL, M180-3), or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (AntGene, ANT011) at 1:10000 dilution in 1% milk/PBS overnight on a shaker at 4℃. After extensive wash, the membrane was incubated with the Horseradish peroxidase (HRP)-conjugated secondary antibody AffiniPure Goat Anti-Mouse IgG (H+L) (Jackson Immuno Reseach, 115-035-003) in 1% skim milk in PBST, and incubated for one hour. The blots were visualized using Omni-ECL Femto Light Chemiluminescence Kit (EpiZyme, SQ201) by ChemiDoc MP (Bio-Rad).

Coronavirus RBD-hFc live-cell binding assay

Recombinant coronavirus RBD-hFc proteins (1-16 μg/ml) were diluted in DMEM and then incubated with the cells for one hour at 37℃. Cells were washed once with DMEM and then incubated with 2 μg/ml of Alexa Fluor 488-conjugated goat anti-human IgG (Thermo Fisher Scientific; A11013) diluted in Hanks’ Balanced Salt Solution (HBSS) with 1% BSA for 1 hour at 37 ℃. Cells were washed twice with PBS and incubated with Hoechst 33342 (1:5,000 dilution in HBSS) for nucleus staining. Images were captured with a fluorescence microscope (MI52-N). For flow cytometry analysis, cells were detached by 5mM of EDTA/PBS and analyzed with a CytoFLEX Flow Cytometer (Beckman).

Biolayer interferometry (BLI) binding assay

The protein binding affinities were determined by BLI assays performed on an Octet RED96 instrument (Molecular Devices). Briefly, 20 μg/mL Human Fc-tagged RBD-hFc recombinant proteins were loaded onto a Protein A (ProA) biosensors (ForteBio, 18-5010) for 30s. The loaded biosensors were then dipped into the kinetic buffer (PBST) for 90s to wash out unbound RBD-hFc proteins. Subsequently, the biosensors were dipped into the kinetic buffer containing soluble ACE2 with concentrations ranging from 0 to 500 nM for 120s to record association kinetic and then dipped into kinetics buffer for 300s to record dissociation kinetics. Kinetic buffer without ACE2 was used to define the background. The corresponding binding affinity (KD) was calculated with Octet Data Analysis software 12.2.0.20 using curve-fitting kinetic analysis or steady-state analysis with global fitting.

Enzyme-linked immunosorbent assay (ELISA)

To evaluate the binding between viral RBD and the ACE2 in vitro, 96 well Immuno-plate were coated with ACE2 soluble proteins at 27 μg/ml in BSA/PBS (100 μl/well) overnight at 4℃. After three wash by PBS containing 0.1% Tween-20 (PBST), the wells were blocked by 3% skim milk/PBS at 37℃ for 2 hrs. Next, varying concentrations of RBD-hFc proteins (1-9 μg/ml) diluted in 3% milk/PBST were added to the wells and incubated for one hour at 37℃. After extensive wash, the wells were incubated with 1:2000 diluted HRP-conjugated goat anti-human Fc antibody (Sigma, T8802) in PBS for one hour. Finally, the substrate solution (Solarbio, PR1210) was added to the plates, and the absorbance at 450nm was measured with a SpectraMax iD3 Multi-well Luminometer (Molecular Devices).

Cell-cell fusion assay

Cell-cell fusion assay based on Dual Split proteins (DSP) was conducted on BHK-21 cells stably expressing different receptors32. The cells were separately transfected with Spike and RLucaa1-155-GFP1-10(aa1-157) expressing plasmids, and Spike and GFP11(aa158-231) RLuc-Caa156-311 expressing plasmids, respectively. At 12 hrs after transfection, the cells were trypsinized and mixed into a 96-well plate at 8×104/well. At 26 hrs post-transfection, cells were washed by DMEM once and then incubated with DMEM with or without 12.5 μg/ml TPCK-trypsin for 25 mins at RT. Five hrs after treatment, the nucleus was stained blue with Hoechst 33342 (1:5,000 dilution in HBSS) for 30min at 37℃. GFP images were then captured with a fluorescence microscope (MI52-N; Mshot). For live-cell luciferase assay, the EnduRen live cell substrate (Promega, E6481) was added to the cells (a final concentration of 30 μM in DMEM) for at least 1 hour before detection by a GloMax 20/20 Luminometer (Promega).

Statistical Analysis

Most experiments were repeated 2∼5 times with 3-4 biological repeats, each yielding similar results. Data are presented as MEAN±SD or MEAN±SEM as specified in the figure legends. All statistical analyses were conducted using GraphPad Prism 8. Differences between independent samples were evaluated by unpaired two-tailed t-tests; Differences between two related samples were evaluated by paired two-tailed t-tests. P<0.05 was considered significant. * p<0.05, ** p <0.01, *** p <0.005, and **** p <0.001.

Author contributions

H.Y. and X.X.W. conceived and designed the study. Q.X., L.C., C.B.M., C.L., J.Y.S., P.L., and F.T. performed the experiments. Q.X, L.C, C.B.M, C.L, C.F.Z., H.Y, and X.X.W analyzed the data. H.Y., X.X.W., Q.X, L.C, C.B.M, and C.L interpreted the results. H.Y and X.X.W wrote the initial drafts of the manuscript. H.Y., X.X.W., H.Y., X.X.W., L.C., and Q.X. revised the manuscript. C.B.M, C.L., P. L., M.X.G., C.L.W, L.L.S, F.T. M.L.H, J.L., C.S., Y.C., H.B.Z., and K.L. commented on the manuscript.

Competing interests

The authors declare no competing interests.

Data availability

The cryo-EM maps have been deposited at the Electron Microscopy Data Bank (www.ebi.ac.uk/emdb) and are available under accession numbers: EMD-32686 (NeoCoV RBD-Bat37ACE2 complex) and EMD-32693 (PDF-2180-CoV RBD-Bat37ACE2 complex). Atomic models corresponding to EMD-32686, EMD-32693 have been deposited in the Protein Data Bank (www.rcsb.org) and are available under accession numbers, PDB ID 7WPO, PDB ID 7WPZ, respectively. The authors declare that all other data supporting the findings of this study are available with the paper and its supplementary information files.

Additional Information

Supplementary Information is available for this paper.

Correspondence and requests for materials should be addressed to H.Y. (huanyan{at}whu.edu.cn)

Spike Proteins, COVID-19, and Vaccines

A new study further elucidates the role of spike proteins in COVID-19.

Authors: Steven Novella  May 5, 2021

A recent study looks at the effects of the SARS-CoV-2 spike proteins, showing that they can cause some of the harm of COVID-19 by themselves. This is an important advance in our understanding of the disease and hopefully will lead to new therapeutic interventions.

The spike protein is what gives the coronavirus family of viruses their name. The spikes jut out from the surface of the spherical virus, giving it a crown-like halo, hence “corona”. We have also known for a long time that the spike protein is the business end of these viruses, it is what gives the virus its ability to target, latch onto, and enter the cells that it infects. Mutations in the spike protein are also what determine different variants of SARS-CoV-2, and can alter its ability to infect and cause harm.

The new study, however, is the first to directly show that the spike proteins themselves are able to cause harm, and also confirms that COVID-19 is primarily a vascular disease that damages blood vessel walls.

What the researchers did was create a pseudovirus – a protein shell with spike proteins but no viral RNA inside. Therefore these pseudoviruses are unable to actually infect cells or replicate. The point was to isolate as much as possible the effects of the spike proteins themselves. They report:

We administered a pseudovirus expressing S protein (Pseu-Spike) to Syrian hamsters intratracheally. Lung damage was apparent in animals receiving Pseu-Spike, revealed by thickening of the alveolar septa and increased infiltration of mononuclear cells.

They had control animals with a mock virus that did not show this damage. The spike protein binds to the ACE2 receptor on cells, downregulates their function, and causes damage to the endothelium cells that line lung tissue and blood vessels. The damage is apparently caused by effects on the mitochondria (energy producing organelles) in the cells – they change their shape and have reduced function. They then reproduced these effects in vitro using a culture of lung endothelial cells exposed to the spike protein.

These results explain many of the clinical features of COVD-19. While the disease has been largely thought of as a respiratory illness, it is primarily a vascular disease. It affects the lungs, but also affects other organs in the body, and can cause strokes and blood clots. While the vascular effects of coronaviruses have long been known, this study demonstrates a clear mechanism of this injury. Knowing the precise mechanism may lead to treatments to prevent or limit the vascular damage from infection. The next step is to study exactly how downregulating the ACE2 receptor damages the mitochondria.

This study, of course, did not come out of the blue but was built on previous studies showing many of the same findings. It was known, for example, that the ACE2 receptor is important for coronavirus infection, and that it related to endothelium damage. In fact a comment to the FDA by Dr. Whelan nicely summarizes a lot of this research as of December 2020. This research, however, has raised some questions about the safety of the mRNA vaccines that produce spike proteins. To be clear, the safety data on the Pfizer and Moderna mRNA vaccines are now extensive, with hundreds of millions of doses give and months of data, without any significant side effects apparent.

The Pfizer and Moderna vaccines produce the full-length spike protein. Pfizer studied several formulations initially, but found that the full length protein vaccine had fewer side effects and was better tolerated than other vaccine candidates, so that is the one they went with. It is also likely that the full protein contains more epitopes (sites for immune activity) and therefore produces more thorough and longer lasting immunity. The proteins, however, are in a fixed state, they are unable to change their confirmation, which is necessary to bind to cells. So they function differently than spike proteins on infecting virus.

After the Pfizer vaccine full spike proteins are expressed on the vaccinated cells for presentation to the immune system. But the vaccine-induced proteins do not appear to cause any harmful effects. This may be because the vaccine is administered in the muscle, and so muscle cells are the ones taking up the mRNA and making spike proteins. There is a vigorous immune response which neutralizes the spike proteins before they can cause any harm. This is very different from a virus replicating throughout the body.

Unfortunately, the complexity of COVID, mRNA, immunity, and vaccines is such that those who wish to raise fears about the vaccine can exploit partial information. There is a tremendous amount of misinformation about the COVID vaccines, and the mRNA vaccines in particularly, which then has to be constantly rebutted and debunked. That has become almost a full-time job for David Gorski here at SBM. Meanwhile, there is legitimate complexity and concerns that scientists need to carefully sort out, which they are doing, transparently and vigorously.

It’s important not to confuse not knowing everything with knowing nothing. The safety data on the mRNA vaccines is robust. Most vaccine serious side effects occur within six weeks, which is why the FDA wanted at least 6 weeks of safety data before giving the vaccines an EUA. We now have more than 6 months of data, including several months with millions of doses. It is very unlikely there are any surprises still in store with either of the mRNA vaccines. The risk is vanishingly small, while the benefit is clear.